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3M2H
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Crystallographic and Single Crystal Spectral Analysis of the Peroxidase Ferryl Intermediate
|
Crystallographic and Single Crystal Spectral Analysis of the
Peroxidase Ferryl Intermediate
Yergalem T. Meharenna, Tzanko Doukovb, Huiying Lia, S. Michael Soltisb,*, and Thomas L.
Poulosa,*
aDepartments of Molecular Biology and Biochemistry, Pharmaceutical Sciences, and Chemistry,
University of California, Irvine, California 92697-3900
bMacromolecular Crystallographic Group, The Stanford Synchrotron Radiation Lightsource,
SLAC, Stanford University, Stanford, California 94025
Abstract
The ferryl (Fe(IV)O) intermediate is important in many heme enzymes and thus the precise nature
of the Fe(IV)-O bond is critical in understanding enzymatic mechanisms. The 1.40 Å crystal
structure of cytochrome c peroxidase Compound I has been solved as a function of x-ray dose
while monitoring the visible spectrum. The Fe-O bond increases linearly from 1.73 Å in the low x-
ray dose structure to 1.90 Å in the high dose structure. The low dose structure correlates well with
a Fe(IV)=O bond while we postulate that the high dose structure is the cryo-trapped Fe(III)-OH
species previously thought to be Fe(IV)-OH.
The ferryl, Fe(IV)O, species is a critically important intermediate in a number of
metalloproteins and especially heme enzymes. The high redox potential enables Fe(IV)O to
serve as a potent oxidant utilized by several heme enzymes including cytochromes P450,
nitric oxide synthase (NOS), cytochrome oxidase, and peroxidases. Since the ferryl
intermediate is quite stable in peroxidases, most of what we know about Fe(IV)O in heme
enzymes derives from studies with peroxidases.
In most heme peroxidases one H2O2 oxidizing equivalent is used to oxidize Fe(III) to
Fe(IV)O and the second is used to oxidize an organic group to give Fe(IV)R.+ (1) and this
activated intermediate is called Compound I. In most heme peroxidases such as horse radish
peroxidase (HRP) R is the porphyrin (2) although in yeast cytochome c peroxidase (CCP) R
is the active site Trp191 (3). A majority of studies find that the Fe(IV)-O bond is short,
somewhat less than 1.7 Å, thus indicating a Fe(IV)=O bond as opposed to a Fe(IV)-OH
bond (4). An empirical formula called Badger’s rule relates the calculated Fe-O bond with
the calculated vibrational frequency (5) and the experimental frequencies and EXAFS bond
distances fit very well to these plots (5) further supporting a Fe(IV)=O double bond.
However, a majority of x-ray crystal structures are distinct outliers giving distances closer to
1.8-1.9 Å (4, 6) with one exception being the HRP Compound I structure (7). These
differences are not trivial since the longer bond predicts that the ferryl species should be
protonated to give Fe(IV)-OH, while the shorter bond gives Fe(IV)=O. The chemistry of
each of these species is quite different (8) and knowing the correct structure is essential if
we are to understand details of heme enzyme mechanisms.
*To whom correspondences should be addressed. T.L.P.: poulos@uci.edu; phone (494) 824-7020; FAX, (949) 824-3280.
SUPPORTING INFORMATION AVAILABLE
Experimental details and Tables 1S and 2S . This material is available free of charge at http://pubs.acs.org.
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 October 27.
Published in final edited form as:
Biochemistry. 2010 April 13; 49(14): 2984–2986. doi:10.1021/bi100238r.
NIH-PA Author Manuscript
NIH-PA Author Manuscript
NIH-PA Author Manuscript
A serious problem encountered at high intensity synchrotron x-ray sources is rapid reduction
of metal centers, particularly high potential metal centers such as Fe(IV). As a result great
care must be taken to minimize reduction and the redox state should be verified during data
collection (for example with UV/VIS spectroscopy). We recently found that crystals of the
CCP N184R mutant diffract unusually well (9) and thus might provide an opportunity to
obtain a low x-ray dose Compound I structure but at sufficiently high resolution to resolve
the discrepancies between crystal structures and solution studies. Here we present single
crystal spectroscopy together with a composite data collection strategy that has allowed the
Fe-O bond distance to be measured as a function of x-ray dose.
Fig. 1A shows the single crystal spectrum of CCP Compound I as a function of x-ray dose.
Before data collection the spectrum in the 500-700 nm region is identical to the solution
spectrum of Compound I. After extensive x-ray exposure (inset to Fig. 1A) the spectrum
clearly is no longer that of Compound I nor is this similar to the Fe(III) high spin solution
spectrum of CCP. The nature of this species will be discussed further on. Fig. 1B shows the
estimated percentage of Compound I remaining in the crystal as a function of x-ray exposure
as monitored by changes in the visible spectrum. Based on this plot ~90% of Compound I
remains after receiving an estimated x-ray dose of 0.035 MGy (calculations were performed
using RADDOSE (10)) or just ~0.1% of the theoretical radiation damage limit for protein
crystals, ≈30 MGy (11). Therefore, a data collection strategy for obtaining predominantly
Compound I was employed using multiple crystals, none of which received more than 0.035
MGy.
With this maximum dose, we estimate that the resulting “integrated” structure is comprised
of ~90% Compound I. Crystallographic data collection was carried out at 65 K on SSRL
BL9-2 (~4×1011 photons/s at 13.0 KeV). Nearly 100 crystals were mounted and indexed in
an automated fashion. Exposures used for indexing were attenuated by 99% and did not
significantly contribute to reduction of Compound I. For each crystal, data collections were
carried out in 15 separate runs. Run 1 consisted of 5° of data, representing the first 0.035
MGy of x-ray exposure. Then the same 5° of scanning angle were recollected 12 more times
giving runs 2 through 13 with increased x-ray dose. In run 14 a full 120° of data were
collected in order to fully reduce the crystal followed by run 15 which again repeated the
same 5° representing the highest x-ray dose. The same 15-run data collection protocol was
adopted for similarly sized crystals and the scanning angles were chosen to optimize the
completeness of the data. Each composite data set was assembled by merging 5° of data
with identical run numbers from 19 crystals. A total of 15 structures at 1.40 Å resolution
were refined providing a picture of the structural changes associated with increasing x-ray
dose (Table S1).
In Fig. 2A we compare the structures of the low dose (set 1) and the ferric resting state 1.06
Å structure of the N184R mutant (3E2O) (9). In the ferric resting state a water molecule is
positioned ≈ 2.0 Å from the heme iron while in the low dose data set the Fe-O oxygen
distance is 1.73 Å. In both structures a water molecule is within H-bonding distance of the
Fe-linked oxygen. In the ferric state the heme iron is displaced from the porphyrin plane by
0.18 Å toward the proximal His ligand while in Compound I the iron is displaced by 0.07 Å
in the opposite direction toward the distal pocket. Thus the net movement of the iron is ≈
0.25 Å relative to the porphyrin plane owing to the oxidation of the iron from Fe(III) to
Fe(IV). Note that the water molecules in the distal pocket, including the one closest to the
iron, are located in nearly the same position relative to the heme while the His-Fe bond
increases from 2.07 Å to 2.12 Å upon oxidation to Fe(IV). Thus, the short Fe-O bond in the
Compound I structure is due in large part to motion of the iron. As in our previous work on
peroxide treated CCP (12) Arg48 in the distal pocket forms a 2.78 Å H-bond with the iron
linked O atom.
Meharenn et al.
Page 2
Biochemistry. Author manuscript; available in PMC 2011 October 27.
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We next compare the set 1 (low dose, Fig. 2C) and set 15 (high dose, Fig. 2D) structures. At
the 4.0 σ contour level the electron density between the Fe and O atoms is not continuous in
set 15 and the Fe-O bond length has increased from 1.73 Å to 1.90 Å. The local water
structure remains largely unchanged. The changes owing to x-ray induced reduction are
highlighted by examining a Fo(low dose)-Fo(high dose) electron density difference map
contoured at ±5σ (Fig. 2B). This map clearly shows that the iron is positioned quite
differently in each structure and is closer toward the distal pocket in the low dose structure.
In addition the His-Fe bond decreases from 2.12 Å to 2.07 Å upon photo reduction again
due to motion of the iron back into the porphyrin plane. The only other notable feature in the
Fo(low dose)-Fo(high dose) difference map is around the carbonyl O atom of the heme
ligand, His175. This group is slightly less than 0.1 Å closer to Trp191 in the low dose
structure and may reflect a local tightening of the structure around the Trp191 cation radical
that provides additional electrostatic stability. The various heme parameter distances are
provided in Table S2.
The structures of set 1 through set 13 next were used to assess how the Fe-O bond changes
as a function of x-ray dose and the results are shown in Fig. 3. The fit to a simple straight
line equation is remarkably good and extrapolates to zero dose at a Fe-O bond distance of
1.72 Å. Raman data (13) coupled with Badger’s rule (4) gives a Fe-O bond of 1.68 Å.
Therefore, the low dose Compound I crystal structure agrees within 0.04 Å with the Raman
data and the ferryl center in CCP Compound I can best be described as Fe(IV)=O and not
Fe(IV)-OH.
The nature of the ferryl center after extensive x-ray exposure is intriguing: the short Fe-O
bond (1.90 Å) compared to the ≈ 2.0 - 2.3 Å observed in Fe(III) high spin peroxidase
structures and the total lack of similarity between the high dose spectrum (Fig. 1) and the
solution spectrum of Fe(III) CCP shows that the high dose structure is not that of Fe(III)
high spin CCP. The spectrum is similar to that of HRP Fe(II) in both the crystal and solution
except in HRP there is no ligand coordinated to the iron (7). Since we clearly see a ligand
coordinated to the iron in the high dose structure we very likely have trapped either Fe(II)-
OH or Fe(III)-OH. Unfortunately we cannot compare single crystal and solution spectra
since formation of Fe(III)-OH, and presumably Fe(II)-OH, requires an increase in pH and
CCP is not stable above pH 8.0.
Our first goal in this study was to further develop the necessary methods and protocols
required to obtain x-ray structures of high potential intermediates in metalloproteins. This
requires isomorphous crystals that diffract well in order to have sufficient resolution to
obtain the level of accuracy required for estimating subtle bond parameter differences (7).
Coupling data collection with on-line single crystal spectroscopy to monitor the redox state
is also essential. Our second goal was to obtain a very low dose x-ray structure of CCP
Compound I at high resolution in order to reconcile the long standing differences observed
in the Fe(IV)-O bond distance between most available x-ray structures and other biophysical
techniques. The low dose CCP Compound I structure agrees within 0.04 Å of previous
experimental estimates indicating that the ferryl species in Compound I is Fe(IV)=O and not
Fe(IV)-OH. It should be noted that from the perspective of the heme, CCP Compound I is
equivalent to HRP Compound II since both contain Fe(IV) with no porphyrin radical. Thus
it is likely that other crystal structures where the Fe(IV)-O bond in Compound II was
estimated to be 1.8 Å (7, 14) or longer may also have a significant amount of a reduced iron
species.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Meharenn et al.
Page 3
Biochemistry. Author manuscript; available in PMC 2011 October 27.
NIH-PA Author Manuscript
NIH-PA Author Manuscript
NIH-PA Author Manuscript
Acknowledgments
We thank Aina Cohen, John Kovarick and Michael Hollenbeck for their contribution to the design and
implementation of the single-crystal microspectrophotometer. Portions of this research were carried out at the
Stanford Synchrotron Radiation Lightsource, a national user facility operated by Stanford University on behalf of
the U.S. Department of Energy, Office of Basic Energy Sciences. The SSRL Structural Molecular Biology Program
is supported by the Department of Energy, Office of Biological and Environmental Research, and by the National
Institutes of Health, National Center for Research Resources, Biomedical Technology Program, and the National
Institute of General Medical Sciences. Work at UCI was supported by NIH grant GM42614 (TLP).
References
1. Poulos TL, Kraut J. J Biol Chem. 1980; 255:8199–8205. [PubMed: 6251047]
2. Dolphin D, Forman A, Borg DC, Fajer J, Felton RH. Proc Natl Acad Sci USA. 1971; 68:614–618.
[PubMed: 5276770]
3. Sivaraja M, Goodin DB, Smith M, Hoffman BM. Science. 1989; 245:738–740. [PubMed: 2549632]
4. Behan RK, Green MT. J Inorg Biochem. 2006; 100:448–459. [PubMed: 16500711]
5. Green MT. J Am Chem Soc. 2006; 128:1902–1906. [PubMed: 16464091]
6. Hersleth HP, Hsiao YW, Ryde U, Gorbitz CH, Andersson KK. Chem Biodivers. 2008; 5:2067–
2089. [PubMed: 18972498]
7. Berglund GI, Carlsson GH, Smith AT, Szoke H, Henriksen A, Hajdu J. Nature. 2002; 417:463–468.
[PubMed: 12024218]
8. Green MT, Dawson JH, Gray HB. Science. 2004; 304:1653–1656. [PubMed: 15192224]
9. Meharenna YT, Oertel P, Bhaskar B, Poulos TL. Biochemistry. 2008; 47:10324–10332. [PubMed:
18771292]
10. Paithankar KS, Owen RL, Garman EF. J Synchr Radiat. 2009; 16:152–162.
11. Owen RL, Rudino-Pinera E, Garman EF. Proc Natl Acad Sci U S A. 2006; 103:4912–4917.
[PubMed: 16549763]
12. Bonagura CA, Bhaskar B, Shimizu H, Li H, Sundaramoorthy M, McRee D, Goodin DB, Poulos
TL. Biochemistry. 2003; 42:5600–5608. [PubMed: 12741816]
13. Reczek CM, Sitter AJ, Terner J. J Molec Struc. 1989; 214:27–41.
14. Hersleth HP, Dalhus B, H GC, A KK. J Biol Inorg Chem. 2002; 7:299–304. [PubMed: 11935353]
Meharenn et al.
Page 4
Biochemistry. Author manuscript; available in PMC 2011 October 27.
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Figure 1.
Single crystal spectra of CCP Compound I as a function of x-ray dose. Prior to x-ray
exposure the spectrum is identical to the solution spectrum of Compound I. The estimated
percentage of Compound I remaining in the crystal as a function of x-ray dose in panel B
was based on the decrease in the absorbance peak at 634 nm.
Meharenn et al.
Page 5
Biochemistry. Author manuscript; available in PMC 2011 October 27.
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Figure 2.
A) Superposition of the low dose structure (red) on the Fe(III) structure (cyan). Note that the
iron is displaced below the plane of the heme in the Fe(III) structure and above the plane of
the heme in the low dose structure; B) Fo(low dose)-Fo(high dose) electron density
difference map using phases obtained from the low dose structure. The map is contoured at
-5.0σ (green) and +5.0σ (blue); C and D) 2Fo-Fc electron density maps contoured at 4.0σ for
the dose data set 1 (panel C) and high dose data set 15 (panel D). Oxygen and water
molecules are represented by the small spheres.
Meharenn et al.
Page 6
Biochemistry. Author manuscript; available in PMC 2011 October 27.
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Figure 3.
Plot of the Fe-O distance as a function of x-ray dose. Each of the 13 structures was refined
exactly the same way using the same starting structure and two different protocols. In the
first the distances between the Fe and N atoms (4 pyrrole and 1 His closed circles) were
restrained while in the second protocol no restraints were applied (open circles). At no time
were restraints imposed on the Fe-O distance. The estimated error in the Fe-O bond distance
is ≈0.017Å (see Supporting Information).
Meharenn et al.
Page 7
Biochemistry. Author manuscript; available in PMC 2011 October 27.
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|
3M2I
|
Crystallographic and Single Crystal Spectral Analysis of the Peroxidase Ferryl Intermediate
|
Crystallographic and Single Crystal Spectral Analysis of the
Peroxidase Ferryl Intermediate
Yergalem T. Meharenna, Tzanko Doukovb, Huiying Lia, S. Michael Soltisb,*, and Thomas L.
Poulosa,*
aDepartments of Molecular Biology and Biochemistry, Pharmaceutical Sciences, and Chemistry,
University of California, Irvine, California 92697-3900
bMacromolecular Crystallographic Group, The Stanford Synchrotron Radiation Lightsource,
SLAC, Stanford University, Stanford, California 94025
Abstract
The ferryl (Fe(IV)O) intermediate is important in many heme enzymes and thus the precise nature
of the Fe(IV)-O bond is critical in understanding enzymatic mechanisms. The 1.40 Å crystal
structure of cytochrome c peroxidase Compound I has been solved as a function of x-ray dose
while monitoring the visible spectrum. The Fe-O bond increases linearly from 1.73 Å in the low x-
ray dose structure to 1.90 Å in the high dose structure. The low dose structure correlates well with
a Fe(IV)=O bond while we postulate that the high dose structure is the cryo-trapped Fe(III)-OH
species previously thought to be Fe(IV)-OH.
The ferryl, Fe(IV)O, species is a critically important intermediate in a number of
metalloproteins and especially heme enzymes. The high redox potential enables Fe(IV)O to
serve as a potent oxidant utilized by several heme enzymes including cytochromes P450,
nitric oxide synthase (NOS), cytochrome oxidase, and peroxidases. Since the ferryl
intermediate is quite stable in peroxidases, most of what we know about Fe(IV)O in heme
enzymes derives from studies with peroxidases.
In most heme peroxidases one H2O2 oxidizing equivalent is used to oxidize Fe(III) to
Fe(IV)O and the second is used to oxidize an organic group to give Fe(IV)R.+ (1) and this
activated intermediate is called Compound I. In most heme peroxidases such as horse radish
peroxidase (HRP) R is the porphyrin (2) although in yeast cytochome c peroxidase (CCP) R
is the active site Trp191 (3). A majority of studies find that the Fe(IV)-O bond is short,
somewhat less than 1.7 Å, thus indicating a Fe(IV)=O bond as opposed to a Fe(IV)-OH
bond (4). An empirical formula called Badger’s rule relates the calculated Fe-O bond with
the calculated vibrational frequency (5) and the experimental frequencies and EXAFS bond
distances fit very well to these plots (5) further supporting a Fe(IV)=O double bond.
However, a majority of x-ray crystal structures are distinct outliers giving distances closer to
1.8-1.9 Å (4, 6) with one exception being the HRP Compound I structure (7). These
differences are not trivial since the longer bond predicts that the ferryl species should be
protonated to give Fe(IV)-OH, while the shorter bond gives Fe(IV)=O. The chemistry of
each of these species is quite different (8) and knowing the correct structure is essential if
we are to understand details of heme enzyme mechanisms.
*To whom correspondences should be addressed. T.L.P.: poulos@uci.edu; phone (494) 824-7020; FAX, (949) 824-3280.
SUPPORTING INFORMATION AVAILABLE
Experimental details and Tables 1S and 2S . This material is available free of charge at http://pubs.acs.org.
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 October 27.
Published in final edited form as:
Biochemistry. 2010 April 13; 49(14): 2984–2986. doi:10.1021/bi100238r.
NIH-PA Author Manuscript
NIH-PA Author Manuscript
NIH-PA Author Manuscript
A serious problem encountered at high intensity synchrotron x-ray sources is rapid reduction
of metal centers, particularly high potential metal centers such as Fe(IV). As a result great
care must be taken to minimize reduction and the redox state should be verified during data
collection (for example with UV/VIS spectroscopy). We recently found that crystals of the
CCP N184R mutant diffract unusually well (9) and thus might provide an opportunity to
obtain a low x-ray dose Compound I structure but at sufficiently high resolution to resolve
the discrepancies between crystal structures and solution studies. Here we present single
crystal spectroscopy together with a composite data collection strategy that has allowed the
Fe-O bond distance to be measured as a function of x-ray dose.
Fig. 1A shows the single crystal spectrum of CCP Compound I as a function of x-ray dose.
Before data collection the spectrum in the 500-700 nm region is identical to the solution
spectrum of Compound I. After extensive x-ray exposure (inset to Fig. 1A) the spectrum
clearly is no longer that of Compound I nor is this similar to the Fe(III) high spin solution
spectrum of CCP. The nature of this species will be discussed further on. Fig. 1B shows the
estimated percentage of Compound I remaining in the crystal as a function of x-ray exposure
as monitored by changes in the visible spectrum. Based on this plot ~90% of Compound I
remains after receiving an estimated x-ray dose of 0.035 MGy (calculations were performed
using RADDOSE (10)) or just ~0.1% of the theoretical radiation damage limit for protein
crystals, ≈30 MGy (11). Therefore, a data collection strategy for obtaining predominantly
Compound I was employed using multiple crystals, none of which received more than 0.035
MGy.
With this maximum dose, we estimate that the resulting “integrated” structure is comprised
of ~90% Compound I. Crystallographic data collection was carried out at 65 K on SSRL
BL9-2 (~4×1011 photons/s at 13.0 KeV). Nearly 100 crystals were mounted and indexed in
an automated fashion. Exposures used for indexing were attenuated by 99% and did not
significantly contribute to reduction of Compound I. For each crystal, data collections were
carried out in 15 separate runs. Run 1 consisted of 5° of data, representing the first 0.035
MGy of x-ray exposure. Then the same 5° of scanning angle were recollected 12 more times
giving runs 2 through 13 with increased x-ray dose. In run 14 a full 120° of data were
collected in order to fully reduce the crystal followed by run 15 which again repeated the
same 5° representing the highest x-ray dose. The same 15-run data collection protocol was
adopted for similarly sized crystals and the scanning angles were chosen to optimize the
completeness of the data. Each composite data set was assembled by merging 5° of data
with identical run numbers from 19 crystals. A total of 15 structures at 1.40 Å resolution
were refined providing a picture of the structural changes associated with increasing x-ray
dose (Table S1).
In Fig. 2A we compare the structures of the low dose (set 1) and the ferric resting state 1.06
Å structure of the N184R mutant (3E2O) (9). In the ferric resting state a water molecule is
positioned ≈ 2.0 Å from the heme iron while in the low dose data set the Fe-O oxygen
distance is 1.73 Å. In both structures a water molecule is within H-bonding distance of the
Fe-linked oxygen. In the ferric state the heme iron is displaced from the porphyrin plane by
0.18 Å toward the proximal His ligand while in Compound I the iron is displaced by 0.07 Å
in the opposite direction toward the distal pocket. Thus the net movement of the iron is ≈
0.25 Å relative to the porphyrin plane owing to the oxidation of the iron from Fe(III) to
Fe(IV). Note that the water molecules in the distal pocket, including the one closest to the
iron, are located in nearly the same position relative to the heme while the His-Fe bond
increases from 2.07 Å to 2.12 Å upon oxidation to Fe(IV). Thus, the short Fe-O bond in the
Compound I structure is due in large part to motion of the iron. As in our previous work on
peroxide treated CCP (12) Arg48 in the distal pocket forms a 2.78 Å H-bond with the iron
linked O atom.
Meharenn et al.
Page 2
Biochemistry. Author manuscript; available in PMC 2011 October 27.
NIH-PA Author Manuscript
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NIH-PA Author Manuscript
We next compare the set 1 (low dose, Fig. 2C) and set 15 (high dose, Fig. 2D) structures. At
the 4.0 σ contour level the electron density between the Fe and O atoms is not continuous in
set 15 and the Fe-O bond length has increased from 1.73 Å to 1.90 Å. The local water
structure remains largely unchanged. The changes owing to x-ray induced reduction are
highlighted by examining a Fo(low dose)-Fo(high dose) electron density difference map
contoured at ±5σ (Fig. 2B). This map clearly shows that the iron is positioned quite
differently in each structure and is closer toward the distal pocket in the low dose structure.
In addition the His-Fe bond decreases from 2.12 Å to 2.07 Å upon photo reduction again
due to motion of the iron back into the porphyrin plane. The only other notable feature in the
Fo(low dose)-Fo(high dose) difference map is around the carbonyl O atom of the heme
ligand, His175. This group is slightly less than 0.1 Å closer to Trp191 in the low dose
structure and may reflect a local tightening of the structure around the Trp191 cation radical
that provides additional electrostatic stability. The various heme parameter distances are
provided in Table S2.
The structures of set 1 through set 13 next were used to assess how the Fe-O bond changes
as a function of x-ray dose and the results are shown in Fig. 3. The fit to a simple straight
line equation is remarkably good and extrapolates to zero dose at a Fe-O bond distance of
1.72 Å. Raman data (13) coupled with Badger’s rule (4) gives a Fe-O bond of 1.68 Å.
Therefore, the low dose Compound I crystal structure agrees within 0.04 Å with the Raman
data and the ferryl center in CCP Compound I can best be described as Fe(IV)=O and not
Fe(IV)-OH.
The nature of the ferryl center after extensive x-ray exposure is intriguing: the short Fe-O
bond (1.90 Å) compared to the ≈ 2.0 - 2.3 Å observed in Fe(III) high spin peroxidase
structures and the total lack of similarity between the high dose spectrum (Fig. 1) and the
solution spectrum of Fe(III) CCP shows that the high dose structure is not that of Fe(III)
high spin CCP. The spectrum is similar to that of HRP Fe(II) in both the crystal and solution
except in HRP there is no ligand coordinated to the iron (7). Since we clearly see a ligand
coordinated to the iron in the high dose structure we very likely have trapped either Fe(II)-
OH or Fe(III)-OH. Unfortunately we cannot compare single crystal and solution spectra
since formation of Fe(III)-OH, and presumably Fe(II)-OH, requires an increase in pH and
CCP is not stable above pH 8.0.
Our first goal in this study was to further develop the necessary methods and protocols
required to obtain x-ray structures of high potential intermediates in metalloproteins. This
requires isomorphous crystals that diffract well in order to have sufficient resolution to
obtain the level of accuracy required for estimating subtle bond parameter differences (7).
Coupling data collection with on-line single crystal spectroscopy to monitor the redox state
is also essential. Our second goal was to obtain a very low dose x-ray structure of CCP
Compound I at high resolution in order to reconcile the long standing differences observed
in the Fe(IV)-O bond distance between most available x-ray structures and other biophysical
techniques. The low dose CCP Compound I structure agrees within 0.04 Å of previous
experimental estimates indicating that the ferryl species in Compound I is Fe(IV)=O and not
Fe(IV)-OH. It should be noted that from the perspective of the heme, CCP Compound I is
equivalent to HRP Compound II since both contain Fe(IV) with no porphyrin radical. Thus
it is likely that other crystal structures where the Fe(IV)-O bond in Compound II was
estimated to be 1.8 Å (7, 14) or longer may also have a significant amount of a reduced iron
species.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Meharenn et al.
Page 3
Biochemistry. Author manuscript; available in PMC 2011 October 27.
NIH-PA Author Manuscript
NIH-PA Author Manuscript
NIH-PA Author Manuscript
Acknowledgments
We thank Aina Cohen, John Kovarick and Michael Hollenbeck for their contribution to the design and
implementation of the single-crystal microspectrophotometer. Portions of this research were carried out at the
Stanford Synchrotron Radiation Lightsource, a national user facility operated by Stanford University on behalf of
the U.S. Department of Energy, Office of Basic Energy Sciences. The SSRL Structural Molecular Biology Program
is supported by the Department of Energy, Office of Biological and Environmental Research, and by the National
Institutes of Health, National Center for Research Resources, Biomedical Technology Program, and the National
Institute of General Medical Sciences. Work at UCI was supported by NIH grant GM42614 (TLP).
References
1. Poulos TL, Kraut J. J Biol Chem. 1980; 255:8199–8205. [PubMed: 6251047]
2. Dolphin D, Forman A, Borg DC, Fajer J, Felton RH. Proc Natl Acad Sci USA. 1971; 68:614–618.
[PubMed: 5276770]
3. Sivaraja M, Goodin DB, Smith M, Hoffman BM. Science. 1989; 245:738–740. [PubMed: 2549632]
4. Behan RK, Green MT. J Inorg Biochem. 2006; 100:448–459. [PubMed: 16500711]
5. Green MT. J Am Chem Soc. 2006; 128:1902–1906. [PubMed: 16464091]
6. Hersleth HP, Hsiao YW, Ryde U, Gorbitz CH, Andersson KK. Chem Biodivers. 2008; 5:2067–
2089. [PubMed: 18972498]
7. Berglund GI, Carlsson GH, Smith AT, Szoke H, Henriksen A, Hajdu J. Nature. 2002; 417:463–468.
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Meharenn et al.
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Biochemistry. Author manuscript; available in PMC 2011 October 27.
NIH-PA Author Manuscript
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Figure 1.
Single crystal spectra of CCP Compound I as a function of x-ray dose. Prior to x-ray
exposure the spectrum is identical to the solution spectrum of Compound I. The estimated
percentage of Compound I remaining in the crystal as a function of x-ray dose in panel B
was based on the decrease in the absorbance peak at 634 nm.
Meharenn et al.
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Biochemistry. Author manuscript; available in PMC 2011 October 27.
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Figure 2.
A) Superposition of the low dose structure (red) on the Fe(III) structure (cyan). Note that the
iron is displaced below the plane of the heme in the Fe(III) structure and above the plane of
the heme in the low dose structure; B) Fo(low dose)-Fo(high dose) electron density
difference map using phases obtained from the low dose structure. The map is contoured at
-5.0σ (green) and +5.0σ (blue); C and D) 2Fo-Fc electron density maps contoured at 4.0σ for
the dose data set 1 (panel C) and high dose data set 15 (panel D). Oxygen and water
molecules are represented by the small spheres.
Meharenn et al.
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Biochemistry. Author manuscript; available in PMC 2011 October 27.
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Figure 3.
Plot of the Fe-O distance as a function of x-ray dose. Each of the 13 structures was refined
exactly the same way using the same starting structure and two different protocols. In the
first the distances between the Fe and N atoms (4 pyrrole and 1 His closed circles) were
restrained while in the second protocol no restraints were applied (open circles). At no time
were restraints imposed on the Fe-O distance. The estimated error in the Fe-O bond distance
is ≈0.017Å (see Supporting Information).
Meharenn et al.
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3M2K
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Crystal Structure of fluorescein-labeled Class A -beta lactamase PenP in complex with cefotaxime
|
RESEARCH ARTICLE
Open Access
Structural studies of the mechanism for
biosensing antibiotics in a fluorescein-
labeled β-lactamase
Wai-Ting Wong, Ho-Wah Au, Hong-Kin Yap, Yun-Chung Leung*, Kwok-Yin Wong* and Yanxiang Zhao*
Abstract
Background: β-lactamase conjugated with environment-sensitive fluorescein molecule to residue 166 on the
Ω-loop near its catalytic site is a highly effective biosensor for β-lactam antibiotics. Yet the molecular mechanism of
such fluorescence-based biosensing is not well understood.
Results: Here we report the crystal structure of a Class A β-lactamase PenP from Bacillus licheniformis 749/C with
fluorescein conjugated at residue 166 after E166C mutation, both in apo form (PenP-E166Cf) and in covalent
complex form with cefotaxime (PenP-E166Cf-cefotaxime), to illustrate its biosensing mechanism. In the apo
structure the fluorescein molecule partially occupies the antibiotic binding site and is highly dynamic. In the PenP-
E166Cf-cefatoxime complex structure the binding and subsequent acylation of cefotaxime to PenP displaces
fluorescein from its original location to avoid steric clash. Such displacement causes the well-folded Ω-loop to
become fully flexible and the conjugated fluorescein molecule to relocate to a more solvent exposed environment,
hence enhancing its fluorescence emission. Furthermore, the fully flexible Ω-loop enables the narrow-spectrum
PenP enzyme to bind cefotaxime in a mode that resembles the extended-spectrum β-lactamase.
Conclusions: Our structural studies indicate the biosensing mechanism of a fluorescein-labelled β-lactamase. Such
findings confirm our previous proposal based on molecular modelling and provide useful information for the
rational design of β-lactamase-based biosensor to detect the wide spectrum of β-lactam antibiotics. The
observation of increased Ω-loop flexibility upon conjugation of fluorophore may have the potential to serve as a
screening tool for novel β-lactamase inhibitors that target the Ω-loop and not the active site.
Background
β-Lactamase is one of the major mechanisms of antibio-
tic resistance in bacteria. Enzymes of this family deacti-
vate β-lactam antibiotics by hydrolyzing the conserved
β-lactam moiety in the antibiotics and rendering them
ineffective to bind to their target proteins, the penicillin-
binding proteins (PBPs), which are essential for bacterial
cell wall synthesis and survival [1,2]. Detailed mechanis-
tic studies of these enzymes over the past decades have
revealed a conserved mechanism of β-lactam hydrolysis
that consists of two steps, the acylation step in which
the β-lactam ring is “opened” and acylated to the side
chain hydroxyl group of Ser70 through nucleophilic
attack to form the enzyme-substrate acyl adduct ES*;
followed by the deacylation step in which the ES* inter-
mediate is hydrolyzed and released as E + P facilitated
by Glu166 (residue numbering according to the most
conserved Class A β-lactamases) [3].
The substrate profile of a β-lactamase in hydrolyzing
diverse β-lactam antibiotics is strongly influenced by a
structural element termed Ω-loop, a short stretch of
residues on the surface of the β-lactamase structure that
forms part of the outer part of the antibiotic binding
site [4-9]. For narrow-spectrum β-lactamases such as
the PenP used in this study and the clinically significant
TEM-1 or SHV-1 enzymes, Ω-loop is tightly packed
onto the enzyme active site through hydrophobic and
electrostatic interactions with residues lining the
* Correspondence: bctleung@inet.polyu.edu.hk; bckywong@inet.polyu.edu.hk;
bcyxzhao@inet.polyu.edu.hk
Department of Applied Biology and Chemical Technology, Central
Laboratory of the Institute of Molecular Technology for Drug Discovery and
Synthesis, The Hong Kong Polytechnic University, Hung Hom, Hong Hong,
China
Wong et al. BMC Structural Biology 2011, 11:15
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© 2011 Wong et al; licensee BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons
Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in
any medium, provided the original work is properly cited.
catalytic site, posing as steric hindrance for binding of
second- or third-generation antibiotics with bulky side
chains attached onto the β-lactam nucleus. Many
mutant strains of TEM- and SHV-like β-lactamases
overcome this inefficiency and broaden their hydrolytic
profile by acquiring mutations in the Ω-loop region to
render this region more flexible to accommodate large-
sized antibiotics [4-10]. Many extended-spectrum
β-lactamases have significantly extended Ω-loop, result-
ing in an enlarged active site that readily binds to and
hydrolyzes almost all antibiotics [11-13].
Exploiting the proximity of Ω-loop to the antibiotic
binding site and its structural flexibility, we have suc-
cessfully converted a β-lactamase PenPC from Bacillus
cereus 569/H into a biosensor for β-lactam antibiotics
by mutating the catalytically critical residue Glu166 on
the Ω-loop to cysteine and conjugating an environment-
sensitive fluorescein molecule to its reactive side chain
thiol group to form PenPC-E166Cf as reported in pre-
vious studies [14-16]. Fluorescein is an environment-
sensitive fluorophore with suppressed fluorescence in a
hydrophobic environment but fluoresces strongly in a
polar aqueous environment [17]. The mutation of
Glu166 to cysteine severely reduces the efficiency of the
deacylation step of β-lactamase catalysis, rendering the
enzyme to stall at the acylation step and form a stable
ES* acyl adduct that enhances the fluorescence emission
of the conjugated fluorescein [16]. We have speculated
that the fluorescein molecule is positioned near the cat-
alytic site so that the binding and subsequent acylation
of β-lactam antibiotics would displace it to a more polar
environment, enhancing its fluorescence intensity [16].
Here we report structural studies of fluorescein-
conjugated PenP β-lactamase from Bacillus licheniformis
749/C to validate our proposed biosensing mechanism.
The structural findings suggest an important role of Ω-
loop in the biosensing process, which will help the
rational design of improved biosensors for β-lactam
detection as well as for novel antibiotics discovery.
Results and Discussion
The biosensing profile of PenP-E166Cf
The biosensing profile of fluorescein conjugated PenP
(PenP-E166Cf) for detecting β-lactam antibiotics have
never been reported before. In our previous study, a
highly similar enzyme, PenPC from Bacillus cereus 569/
H with 58% amino acid sequence identity to PenP, was
successfully engineered into a biosensor using the same
design scheme (PenPC-E166Cf) [15,16]. We chose to
work with PenP in this study for the advantage of its
easy propensity for crystallization, which would enable
structural studies to understand its biosensing mechan-
ism at atomic resolution. PenPC, on the other hand, has
poor thermal stability and is difficult to crystallize.
Because of the high sequence similarity between these
two proteins, as well as the general sequence conserva-
tion among all Class A β-lactamase enzymes we expect
that PenP can serve as a good model system to under-
stand the biosensing mechanism of fluorescein-based
biosensing.
Indeed the biosensing profile of PenP-E166Cf is highly
similar to that of PenPC-E166Cf. The conjugation of
fluorescein to the mutated Cys166 residue through thiol
linkage is highly efficient for PenP. The ESI-MS profile
confirmed that over 90% of PenP was labelled by the
fluorophore and converted to PenP-E166Cf, with little
unlabelled PenP remaining (Figure 1a). The fluorescence
scanning spectrum of PenP-E166Cf shows an increase of
~25% in emitted intensity when the antibiotic cefotaxime
is present at 10 μM concentration (Figure 1b). A variety
of β-lactam antibiotics, including the first-, second- and
third-generation compounds with diverse chemical struc-
tures in addition to the conserved β-lactam core, induce
significant fluorescence enhancement in PenP-E166Cf at
concentration as low as 1 μM (Figure 1c). Lastly the
time-dependent spectra of PenP-E166Cf in the presence
of cefotaxime at different concentrations ranging from
0.01 μM to 10 μM shows that PenP-E166Cf can detect
cefotaxime at concentration as low as 0.01 μM and the
fluorescence response is saturated at 1 μM (Figure 1d).
The structure of PenP-E166Cf in apo form
PenP-E166Cf readily crystallized in the form of clustered
needles. These crystals were tinted in bright yellow col-
our, indicating the presence of fluorescein (data not
shown YZ). To confirm that fluorescein remaining con-
jugated to the protein in the crystal form we harvested
and thoroughly washed these yellow-coloured crystals
and analyzed the dissolved crystals on SDS-PAGE gel
under both visible and UV light. A band corresponding
to PenP (~30.5 kDa) is clearly visible under both condi-
tions, confirming that the crystals are indeed of PenP-
E166Cf (data not shown YZ).
The structure of PenP-E166Cf was solved by molecu-
lar replacement using the known structure of PenP
(PDB ID 4BLM) as search model. Two molecules of
PenP-E166Cf are found in each asymmetric unit. Struc-
ture rebuilding and refinement were done in CCP4 pro-
gram [18]. The overall structure of PenP-E166Cf is
largely identical to that of the wild-type unlabeled PenP.
The RMSD of all 4011 protein atoms between the
labeled and wild-type structures is just ~1.5 Å. For main
chain atoms, the RMSD is only 0.8 Å. Key residues lin-
ing the catalytic site, including Ser70 and mutated
Cys166 are virtually identical between the labeled and
wild-type structures (Figure 2a). In summary the conju-
gation of fluorescein to PenP does not alter its overall
structural folding.
Wong et al. BMC Structural Biology 2011, 11:15
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The fluorescein molecule was modeled onto the PenP
structure after careful inspection of the fo-fc and 2fo-fc
electron density map. These maps are not of high qual-
ity at regions around the fluorescein conjugation site,
with only pieces of discontinuous density visible at 2.0 s
contour level in the fo-fc map (Figure 2a). We tried our
best to fit fluorescein into these pieces of electron den-
sity, particularly matching the melaimide group to a
piece of electron density near the thiol side chain of
Cys166, as well as matching the xanthene group at the
end of the fluorescein molecule to a large piece of elec-
tron density near the catalytic site (Figure 2a). This
modeled structure is stable after rounds of structural
refinement, showing good electron density for the
Ω-loop residues and the fluorescein molecule at 1.0 s
contour level in the 2fo-fc map, suggesting that our
fitting is reasonable (Figure 2b). However, no electron
density was visible for the benzoic group in the
30053 Da
(a)
(d)
PenP(29608Da)+fluorescein(427Da)+water(18Da)=30053Da
(c)
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1
505
525
545
565
585
Wavelength (nm)
Relative Fluorescence Intensity
E166Cf only
10-5M cefotaxime
(b)
10-5
10-6
10-7
5x10-8
10-8
E166Cf only
Figure 1 Biosensing of b-lactam antibiotics by fluorescein-labelled PenP. (a) De-convoluted ESI mass spectrum of PenP-E166Cf. The add-up
at the bottom confirms the correct mass of the labelled protein. (b) Fluorescence scanning spectra of PenP-E166Cf in the presence of 10-5M
cefotaxime in 50 mM phosphate buffer (pH 7.0). (c) Change in fluorescence emission of PenP-E166Cf after incubation with different antibiotics
(cefotaxime, ceftriaxone, ceftazidime, cephaloridine, cephalothin, cefoxitin, cefuroxime, penicillin G and ampicillin) at 10-6 M for 100 s. (d) Time-
dependent fluorescence spectra in the presence of different concentrations (1 × 10-8 M - 1 × 10-5 M) of cefotaxime monitored at 515 nm.
Wong et al. BMC Structural Biology 2011, 11:15
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Page 3 of 8
mid-region of the fluorophore molecule, indicating that
this region is more disordered as compared to other
parts of the fluorophore molecule.
In our PenP-E166Cf structure the fluorescein molecule
partially occupies the outer edge of the antibiotic binding
region and is in close contact with several residues at the
catalytic site. The maleimide moiety near the thiol link-
age site is inserted into the catalytic core, located within
2.5 Å from the side chain of Ser70 on one side and 3.5 Å
away from Ω-loop on the other side. The xanthene group
near the other end of the fluorescein molecule extends
toward the solvent (Figure 2b), loosely packed against
β-strand B3 that forms part of the extended substrate
binding area involved in coordinating antibiotics as
shown in the extended-spectrum class A β-lactamase,
Toho-1, in complex with cefotaxime, cephalothin, and
benzylpenicillin [19]. No specific interactions were
observed between fluorescein and the protein. Total sol-
vent accessible area is 188 Å2, 33% of the total surface
area, indicating that fluorescein is partially packed against
the PenP molecule and not fully solvent exposed. The
fluorescein molecule is highly dynamic, as reflected by
the poor electron density map as well as high average
temperature factor (~72.3). In contrast, the rest of the
structure shows excellent electron density and low aver-
age temperature factor (~23.5) that is typical of the 2.2 Å
data set. The Ω-loop, on which the fluorescein molecule
is conjugated, was little affected by the dynamic fluoro-
phore and adopts the same conformation as that of the
unlabelled PenP (Figure 2b).
The structure of PenP-E166Cf in complex with cefotaxime
We chose to determine the PenP-E166Cf-cefotaxime
structure, using cefotaxime as a representative of the
many β-lactam antibiotics because of its positive fluores-
cence response induced in PenP-E166Cf as well as its
chemical structure that contains functional groups typi-
cal of both second- and third-generation antibiotics.
Cefotaxime was soaked into the PenP-E166Cf crystals
by incubating the crystals in the reservoir solution with
0.01 M cefotaxime added for 20 minutes. The PenP-
E166C structure, without the conjugated fluorescein
molecule, was used as the starting model for structure
determination. After initial rounds of refinement both
the fo-fc and 2fo-fc electron density maps were carefully
inspected for evidence of cefotaxime and fluorescein, as
well as for any structural changes on PenP.
The cefotaxime was clearly visible in fo-fc electron
density map as covalently bonded through its carbonyl
carbon atom C7 to the Og atom of Ser70, which repre-
sents the acylated ES* adduct (Figure 3a). But we could
not identify any electron density in either fo-fc or 2fo-fc
:-loop
fluorescein
Ser70
Cys166
2.5 Å
(b)
Ser70
Cys166
:-loop
(a)
Figure 2 Crystal structure of PenP-E166Cf. (a) The fo-fc omit map of fluorescein-5-maleimide contoured at 2.0 s. (b) The 2fo-fc map of
Phe165 to Asn170 and fluorescein-5-maleimide contoured at 1.0 s. Side chains of Phe165 to Asn170 and Ser70 are shown in cpk cylinder
model. Fluorescein is shown in green cylinder model.
Wong et al. BMC Structural Biology 2011, 11:15
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map that would be accountable for fluorescein molecule
around the location seen in PenP-E166Cf or anywhere
nearby. Furthermore, the fo-fc map showed strong nega-
tive signal for a large segment of Ω-loop (residues 164
to 174) and the 2fo-fc map showed no electron density
for this region at all, indicating this region became
highly
disordered
upon
acylation
of
cefotaxime
(Figure 3b). Based on these observations we did not
include fluorescein molecule or the disordered region of
Ω-loop in our final refined structure of PenP-E166Cf-
cefotaxime.
The overall structure folding of fluorescein-labeled
and cefotaxime-bound PenP is nearly identical to that of
the wild-type unlabeled PenP and the fluorescein-labeled
PenP-E166Cf. From the calculation result by the CCP4
program, it was found that the B factor of Glu163,
(c)
:-loop
GC1
Toho-1
PenP-E166Cf
(c)
:-loop
GC1
Toho-1
PenP-E166Cf
Ser70
cefotaxime
Ω-loop
Cys166
Ser70
cefotaxime
fluorescein
:-loop
Cys166
Ser70
cefotaxime
fluorescein
:-loop
(a)
(b)
(c)
Figure 3 Crystal structure of PenP-E166Cf-cefotaxime. (a) The fo-fc map of cefotaxime in PenP-E166Cf-cefotaxime complex contoured at 2.0
s. The light blue dash line represents the disordered Arg164 to Pro174 due to the poor electron density. (b) Comparison of PenP-E166Cf-
cefotaxime complex with apo PenP-E166Cf structure. The two structures are superimposed by main chain atoms. Key residues including Cys166,
Ser70 and cefotaxime are also shown in cpk cylinder model. (c) Comparison of binding mode of cefotaxime in PenP-E166Cf with that of Toho-1
and GC-1. PenP-E166Cf, Toho-1 and GC1 are superimposed by aligning on overall main chain atoms. Cefotaxime is in cylinder model colored in
cpk (PenP-E166Cf-cefotaxime), golden (Toho-1) and red (GC1) respectively.
Wong et al. BMC Structural Biology 2011, 11:15
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Gly175, Glu176 on Ω-loop, which are next to the disor-
dered region, is significantly higher (~65 Å2) than other
parts of the protein (~20 Å2). The refinement statistic
for this set of crystal structure has different values from
that of the apo PenP-E166Cf structure due to the cefo-
taxime and the difference in Ω-loop.
To investigate why the binding and acylation of cefotax-
ime causes the Ω-loop and the conjugated fluorescein
molecule to become highly flexible and structural disor-
dered, we superposed the PenP-E166Cf structure onto the
PenP-E166Cf-cefotaxime complex structure. Fluorescein is
seen as occupying a site that partially overlaps with the
acylated cefotaxime; particularly the benzoic group of
fluorescein molecule is in direct steric clash with the
7-amino substituent of cefotaxime (Figure 3b). Thus the
binding and acylation of cefotaxime to PenP would dis-
place fluorescein from its original position to avoid steric
clash. It is likely that the Ω-loop, in order to accommodate
such displacement, loses its well-folded structure and
becomes highly flexible. As a consequence the fluorescein
molecule conjugated to the flexible Ω-loop becomes fully
exposed to the polar aqueous environment, leading to
enhanced fluorescence. Thus our structural findings con-
firmed our initial proposal of a biosensing mechanism
based on displacement of fluorescein [15,16].
To understand the impact of conjugated fluorescein
molecule on the substrate binding kinetics of PenP we
compared the PenP-E166Cf-cefotaxime structure to two
other β-lactamase structures in complex with cefotaxime,
including the narrow-spectrum Toho-1 and the
extended-spectrum GC1 [19,20]. In Toho-1 structure the
methoxyimino side chain points away from the active site
and is solvent-exposed (Figure 3c). Such an orientation
packs the methoxyimino side chain tightly against the
thiozolyl ring, leading to a distorted configuration of the
cephem nucleus that is catalytically incompetent for dea-
cylation [19]. In GC1 structure the transition analog of
cefotaxime binds to GC1 in a fully extended conforma-
tion, with oxyimino group inserted to active site and
extended away from the thiozolyl ring (Figure 3c). This
conformation is regarded as catalytically competent to
facilitate deacylation because the distortion on the
cephem nucleus is released [20]. Importantly, the binding
mode of cefotaxime in our PenP-E166Cf-cefotaxime
structure closely resembles that of GC1 (Figure 3c), sug-
gesting that with its Ω-loop fully flexible the naturally
narrow-spectrum PenP can accommodate cefotaxime in
a manner that resembles the extended-spectrum GC1.
Conclusions
Our structural studies indicate the molecular mechan-
ism how fluorescein-labeled β-lactamase detects β-lac-
tam antibiotics. The conjugated fluorescein molecule is
located near the catalytic site and partially occupies the
antibiotic binding region. The binding and acylation of
β-lactam antibiotics such as cefotaxime would expel the
fluorescein molecule from its original position and leads
to increased flexibility of the Ω-loop, to which the fluor-
ophore is linked. As a result, the fluorophore is relo-
cated from its original position with partial solvent
exposure to become fully solvent exposed, leading to
enhanced fluorescence emission. These findings confirm
our previous proposal based on structural modeling.
Furthermore the Ω-loop demonstrates the propensity
of becoming highly flexible and unstructured if its tight
packing against the catalytic site is disturbed. Such
increased flexibility enables PenP to bind and acylate
cefotaxime, a naturally poor substrate, in a manner that
resembles the extended-spectrum cefotaxime-resistant
β-lactamases. This finding could be valuable in the
future design of novel antibiotics that resist the binding
or hydrolysis by β-lactamases.
Methods
Protein expression and purification
Two constructs of PenP protein were used for our
experiments, the maltose binding protein (MBP)-fusion
construct for time-dependent fluorescence measure-
ments and the His6-tagged construct for crystallization
and structural studies, as well as scanning fluorescence
spectra. The MBP fusion has been shown not to inter-
fere with fluorescence measurements in our previous
studies (data not shown). The MBP-fusion construct
was cloned into pMAL-c2X vector (NEB). The His6-
tagged PenP enzyme was cloned into a modified pRset-
A vector (Invitrogen) with a TEV protease cleavage site
upstream of the PenP gene. The E166C mutation was
constructed using QuikChange Site-Directed Mutagen-
esis Kits (Strategene).
The MBP-fusion construct was expressed in E. coli
strain BL21 (DE3) at 37°C for overnight after induction
by 300 μM IPTG when A600 reached 0.5-0.7. The har-
vested cells were centrifuged and lysed by sonication.
The supernatant after sonication was passed through
amylose affinity chromatography. The eluted fractions
were pooled and buffer exchanged to 20 mM ammo-
nium bicarbonate. The protein was freeze-dried for sto-
rage afterwards.
The His6-tagged PenP protein was expressed in E. coli
strain BL21 (DE3) at 37°C for overnight after induction
by 200 μM IPTG when A600 reached 0.8-1.2. The har-
vested cells were centrifuged and the supernatant was
passed through Nickel affinity chromatography, followed
by DEAE anion exchange chromatography. The frac-
tions containing the target protein were pooled and con-
centrated by Amicon® Ultra-15 Centrifugal Filter
Devices (Millipore NMWL = 10,000). The His6-tag was
cleaved by adding the TEV protease in 1:20 molar ratio
Wong et al. BMC Structural Biology 2011, 11:15
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to the concentrated PenP-E166C protein (2 mg/ml). The
mixture was incubated at 30°C for 6 hours and was
further purified by Nickel affinity chromatography to
remove uncleaved protein.
Fluorescein labeling of PenP-E166C to form PenP-E166Cf
A ten-fold molar excess of fluorescein, with concentra-
tion of 20 mM, was dissolved in DMF (Dimethyl forma-
mide) and added to the concentrated PenP-E166C
protein solution drop by drop. The labelling reaction
was allowed to proceed in darkness with stirring for
1 hour, and then dialysed against 50 mM potassium phos-
phate buffer (pH 7.0) at 4°C for several times in order to
remove excess fluorescein. The labelled PenP-E166Cf pro-
tein was concentrated to less than 1 ml and further puri-
fied by Superdex™75 gel filtration column (GE
Healthcare). The running buffer contains 20 mM Tris-
HCl, 50 mM NaCl, pH 7.5. The target fractions were
pooled and concentrated by Amicon Ultra to 25 mg/ml.
The labelling efficiency was confirmed by ESI-MS.
Fluorescence spectra of PenP-E166Cf for antibiotic
detection
Fluorescence profile of PenP-E166Cf alone, as well as in
presence of various β-lactams were measured using Per-
kin-Elmer LS50B spectrofluorimeter. Both scanning
spectra and time-dependent spectra were measured. Dif-
ferent β-lactam antibiotics, including cefotaxime, cef-
triaxone, ceftazidime, cephaloridine, cephalothin,
cefoxitin, cefuroxime, penicillin G, and ampicillin, were
incubated with PenP-E166Cf for 100 s at 1 μM to allow
sufficient acylation of the antibiotic to form ES* adduct.
The product after acylation was subjected to fluores-
cence measurement as previously described [15].
Crystallization, structure determination and refinement
Crystals of PenP-E166Cf were grown by hanging-drop
vapour diffusion method after mixing 1 μl of protein
and 1 μl of reservoir solution containing 25% (w/v) PEG
4000, 0.1 M Hepes pH 7.2, 0.4 M NH4Acetate and 0.2
M K2HPO4. Small crystals in the form of clustered nee-
dles appeared readily. For data collection, single crystals
were obtained after separating them from the clustered
needles. Crystals were harvested and cryoprotected in its
reservoir solution supplemented with 20% ethylene gly-
col for one minute prior to flash freeze and data collec-
tion on the Rigaku MicroMax™-007HF x-ray machine.
For PenP-E166Cf-cefotaxime data set, crystals were
soaked in its growth solution added with 0.01 M of
cefotaxime for 15 minutes and then mounted to the
x-ray machine. Data were integrated and scaled by Crys-
talClear™1.3.5 SP2 (Rigaku Inc.).
The crystals belong to the monoclinic group P21 with
cell parameter: a = 43.43 Å, b = 92.3 Å, c = 66.43 Å and
β = 104°. The PenP-E166Cf crystals diffracted to 2.15 Å
resolution, while the PenP-E166Cf-cefotaxime crystal
diffracted to 2.8 Å. Both structures were determined by
molecular replacement using PenP structure as the
search model (PDB ID 4BLM) [21]. The program
COOT was used for inspection of electron density maps
and model building [22]. There are two molecules per
asymmetric unit. The fluorescein and cefotaxime mole-
cules were built by PRODRG [23] and appended to the
PenP structure for refinement. Structure determination
and refinement of PenP-E166Cf and PenP-E166Cf-
cefotaxime were done using the CCP4 program suite
[18]. A summary of the crystallographic data and refine-
ment statistics are given in Table 1. The coordinates
and structure factors from this study have been
Table 1 X-ray data-collection and structure refinement
statistics.
E166Cf
E166Cf+cefotaxime
PDB code
3M2J
3M2K
Data collection
Space group
P21
P21
Unit cell parameters (Å)
a
43.3
43.5
b
92.3
91.4
c
66.3
66.1
b
104.82
104.52
Resolution range (Å)
52-2.15
(2.24-2.15)
45-2.80
(2.95-2.80)
No. of total reflections
79750
40611
No. of unique reflections
29537
12412
I/s
7.1 (2.7)
6.3 (2.4)
Completeness (%)
97.0 (99.5)
99.8 (99.9)
Rmerge (%)
9.7 (27.1)
11.8 (32.0)
Structure refinement
Resolution (Å)
50.0-2.20
45.0-2.80
Rcryst/Rfree (%)
20.0/23.2
21.2/27.7
r.m.s.d. bonds (Å)/angles (°)
0.018/1.784
0.010/1.672
No. of reflections
Working set
24217
11749
Test set
1291
647
No. of atoms
Protein atoms
4011
3706
Water molecules
254
29
Average B-factor (Å2)
Main chain
24.7
16.96
Ligand molecules
48.4
42.46
Water
32.7
10.7
Wong et al. BMC Structural Biology 2011, 11:15
http://www.biomedcentral.com/1472-6807/11/15
Page 7 of 8
deposited into Protein Data Bank (PDB) under accession
codes 3M2J (PenP-E166Cf apo structure) and 3M2K
(PenP-E166Cf-cefotaxime).
Acknowledgements
This work was supported by the Research Grants Council (PolyU 5463/05 M,
PolyU 5017/06P, PolyU 5641/08 M, and PolyU 5639/09M), the Area of
Excellence Fund of the University Grants Committee (AoE/P-10/01) and the
Research Committee of the Hong Kong Polytechnic University. We thank
Shanghai Synchrotron Radiation Facility (SSRF) for access to beam time. Mr.
C.H. Cheng is acknowledged for technical assistance with in-house x-ray
crystallography facility.
Authors’ contributions
WTW performed experiments, analyzed data and drafted manuscript. HWA
and HKY assisted in experiments. YXZ, KYW and YCL designed project,
analyzed data and drafted manuscript. All authors read and approved the
final manuscript.
Received: 21 September 2010 Accepted: 28 March 2011
Published: 28 March 2011
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Cite this article as: Wong et al.: Structural studies of the mechanism for
biosensing antibiotics in a fluorescein-labeled β-lactamase. BMC
Structural Biology 2011 11:15.
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|
3M2L
|
Crystal structure of the M113F mutant of alpha-hemolysin
|
Molecular bases of cyclodextrin adapter interactions
with engineered protein nanopores
Arijit Banerjeea, Ellina Mikhailovaa, Stephen Cheleya, Li-Qun Gub,2, Michelle Montoyac,3, Yasuo Nagaokaa,4,
Eric Gouauxd, and Hagan Bayleya,1
aDepartment of Chemistry, University of Oxford, Oxford, OX1 3TA, United Kingdom; bDepartment of Medical Biochemistry & Genetics, Texas A&M
University System Health Science Center, College Station, TX 77843-1114; cDepartment of Biochemistry and Molecular Biophysics and Howard Hughes
Medical Institute, Columbia University, New York, NY 10032; and dVollum Institute and Howard Hughes Medical Institute, Oregon Health and Science
University, Portland, OR 97239
Edited by Gregory A. Petsko, Brandeis University, Waltham, MA, and approved March 3, 2010 (received for review December 15, 2009)
Engineered protein pores have several potential applications in
biotechnology: as sensor elements in stochastic detection and
ultrarapid DNA sequencing, as nanoreactors to observe single-
molecule chemistry, and in the construction of nano- and micro-
devices. One important class of pores contains molecular adapters,
which provide internal binding sites for small molecules. Mutants
of the α-hemolysin (αHL) pore that bind the adapter β-cyclodextrin
(βCD) ∼104 times more tightly than the wild type have been ob-
tained. We now use single-channel electrical recording, protein en-
gineering including unnatural amino acid mutagenesis, and high-
resolution x-ray crystallography to provide definitive structural in-
formation on these engineered protein nanopores in unparalleled
detail.
alpha-hemolysin ∣single molecule ∣stochastic sensing ∣structure ∣
unnatural amino acid
M
any research groups have used protein engineering to
obtain enzymes and antibodies with new properties suited
for specific tasks (1–6). Fewer groups have taken on the difficult
problem of engineering membrane proteins (7). We have engi-
neered the α-hemolysin protein pore, mindful of several potential
applications in biotechnology, including its ability to act as a de-
tector in stochastic sensing (8) and ultrarapid DNA sequencing
(9), to serve as a nanoreactor for the observation of single-
molecule chemistry (10) and to act as a component for the con-
struction of nano- and microdevices (11).
An important breakthrough in this area, which enabled the sto-
chastic sensing of organic molecules including the detection of
DNA bases in the form of nucleoside monophosphates (12, 13),
was the discovery of internal molecular adapters, a form of non-
covalent protein modification (14). Most useful have been cyclo-
dextrin (CD) adapters, which have until now been used in the
absence of detailed structural information about how they work.
The present paper is a definitive investigation, which provides
such information through the application of a wide variety of
technical approaches: single-channel electrical recording, protein
engineering including unnatural amino acid mutagenesis, and
x-ray crystallography. The studies employing mutagenesis show
that the striking interactions seen in the crystal structures also
occur in individual pores in lipid bilayers.
We reveal that the tight-binding αHL mutants (15) M113N7
and M113F7 bind βCD in different orientations within the hep-
tameric pore. In the case of M113N7, the top (primary hydroxyls)
of the CD ring faces the trans entrance of the pore. In the case of
M113F7, the bottom (secondary hydroxyls) of the CD ring faces
the trans entrance, while the top of the ring is bonded to the pore
through remarkable CH-π interactions. Another tight-binding
mutant, M113V7, can bind the CD in both orientations. These
results illustrate the exquisite level of engineering that can be
achieved with protein nanopores, which is, for example, far be-
yond what is possible with solid-state pores. The work also pro-
vides information valuable for the design of new binding sites
within the lumen of the αHL pore or within other β-barrel pro-
teins. Our results will be of interest to others exploring the inter-
actions of CDs with the αHL pore (16, 17), including groups
involved in computational studies (18, 19). In addition CDs bind
to a variety of other pores, including porins (20, 21) and connex-
ins (22), and are being tested in vivo as blockers of the anthrax
protective antigen pore (23, 24). The CD adapter concept has
also been incorporated into other formats, e.g., with glass nano-
pores (25), and artificial pores based on CDs have been made by
several groups (26–28). Our work is pertinent to these studies.
Results
Kinetics and Thermodynamics of the Interactions of βCD with αHL
Pores Containing Met, Phe and Asn at Position 113. We showed earlier
that position 113 in the αHL pore (Fig. 1A) is critical for the bind-
ing of βCD (14). Subsequently, residue 113, which is Met in the
WT protein, was changed to each of the remaining 19 naturally
occurring amino acids by site-directed mutagenesis (15). We
found that 11 of these mutants, expressed as homoheptamers,
bound βCD with a similar affinity and with similar kinetics to
the WT homoheptamer. Two mutants (P, W) bound βCD about
10 times more strongly than the WT homoheptamer, while six of
them (V, H, Y, D, N, F) bound with high affinity, i.e., with a Kd
value 103 to 104 times lower than the WT.
Remarkably, the side chains of the latter six amino acids bear
little resemblance to one another, and this issue is addressed in the
present paper. We first examined the two amino acids with the
most disparate side chains (Fand N) by making heteromeric pores
containing WT (Met-113), M113F, and M113N subunits. Three
series of heteroheptamers were produced: WT7−nM113Nn,
WT7−nM113Fn, and M113F7−nM113Nn. The heteroheptamers
were separated by SDS-polyacrylamide gel electrophoresis aided
by an oligoaspartate (D8) tail on the first of the two types of sub-
unit (Fig. 1B) (29). All 21 combinations of WT, M113F, and
M113N subunits formed αHL pores that interacted with βCD as
shown by single-channel current recordings, which revealed the
extent of block by βCD (Fig. S1), the association and dissociation
Author contributions: A.B., S.C., E.G., and H.B. designed research; A.B., E.M., S.C., L.-Q.G.,
M.M., and Y.N. performed research; A.B., E.G., and H.B. analyzed data; and A.B. and
H.B. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
1To whom correspondence should be addressed. E-mail: hagan.bayley@chem.ox.ac.uk.
2Present address: Department of Biological Engineering and Dalton Cardiovascular
Research Center, University of Missouri, Columbia, MO 65211.
3Present address: Nature Structural & Molecular Biology, 75 Varick Street, 9th Floor, New
York NY 10013-1917.
4Present address: Department of Biotechnology, Faculty of Engineering, Kansai University,
3-3-35 Yamate-cho, Suita, Osaka 564-8680, Japan.
This article contains supporting information online at www.pnas.org/cgi/content/full/
0914229107/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.0914229107
PNAS ∣May 4, 2010 ∣vol. 107 ∣no. 18 ∣8165–8170
BIOCHEMISTRY
rate constants for βCD (kon and koff), and (from the latter) the
equilibrium dissociation constant for βCD (Kd ¼ koff∕kon) (15).
The kon values for βCD for the 21 combinations of subunits
were all similar at ∼5 × 105 M−1 s−1 (Fig. 1C, Upper). By contrast,
the koff values differed widely, ranging from ∼5 × 10−2 s−1 to
∼103 s−1. For WT7−nM113Nn and WT7−nM113Fn, the koff values
decreased as M113N or M113F subunits were added. In the case
of M113N, there was a steep drop in the value of koff after the
fifth subunit had been incorporated. In the case of M113F,
the decrease in the value of koff occurred less precipitously as the
M113F subunits were added (Fig. 1C, Lower). Intriguingly, with
M113F7−nM113Nn, koff first increased as M113N subunits were
added to M113F7 until n ¼ 4 (M113F3M113N4) and then de-
creased for larger values of n (Fig. 1C, Lower). We recognize that
there is more than one permutation of heteromers containing two
to five mutant subunits (Fig. 1B), but we have ignored this fact
here because no significant differences in the properties of indi-
vidual heteromers were observed. For example, 42 recordings
were made of WT5M113N2, which has three permutations.
Because, kon showed little variation with subunit composition,
the variation in Kd was similar to the variation in koff (Fig. 1C).
While these studies were in progress, the crystal structures of
βCD complexed to M113N7 (Fig. 2B) and M113F7 (Fig. 2C) were
solved (Table S1) (30). High-resolution structures could be
obtained because the CD and the αHL pore have the same C7
symmetry. In the case of M113N7, βCD is bound with the second-
ary hydroxyl face “upward” (Fig. 2B). In each glucose unit of the
βCD, the 2-hydroxyl is hydrogen bonded to the side-chain amide
of an Asn-113 (the residue introduced by mutagenesis) and the
3-hydroxyl is hydrogen bonded to the ϵ-amino group of Lys-147.
In the case of M113F7, two βCDs are bound to the αHL pore
(Fig. 2C). It is the top βCD in the structure that concerns us, be-
cause it is in contact with the Phe-113 residues introduced by mu-
tagenesis. It is immediately apparent that the top βCD in M113F7
is in the opposite orientation to the βCD in M113N7 with each
6-hydroxyl group in a CH-π bonding interaction (31–35) with a
Phe-113 side chain. The opposite orientations of the βCDs in
M113N7 and M113F7 immediately explain why heteromers
formed from similar numbers of M113N and M113F subunits
(e.g., M113N4M113F3) bind βCD weakly (see also Discussion).
Unnatural Amino Acid Mutagenesis. To further explore the range of
noncovalent interactions that are available when βCD binds to
the αHL pore, five unnatural amino acids (Fig. 3A and Fig. S2)
were incorporated at position 113, by using the in vitro nonsense
codon suppression method (36). In particular, we had noted that
M113V7 containing the β-branched Val binds βCD tightly (15),
and therefore we compared cyclopropylglycine (Cpg) and cyclo-
propylalanine (Cpa). We also further examined the means by
which M113F7 binds βCD tightly, by comparing the properties of
4-fluorophenylalanine (f1F), pentafluorophenylalanine (f5F),
and cyclohexylalanine (Cha) at position 113.
The five homomeric pores all produced single-channel cur-
rents with unitary conductance values in the range expected
for properly assembled heptamers (Fig. S3). All five bound βCD
(Fig. 3B, Level 2), either tightly (f1F, Cpg) or weakly (f5F, Cha,
Cpa) as described in detail below. During the long βCD binding
events, additional current spikes were seen (Fig. 3B). Similar
Fig. 1.
Binding of βCD by heteromeric αHL pores formed by WT, M113F and M113N subunits. (A) Crystal structure of WT-αHL (61) showing residue 113 (Met,
yellow). Left panel, side view and right panel, top view. (B) Separation of 35S-labeled αHL heteroheptamers by SDS-polyacrylamide electrophoresis. The
separation of the M113F7-nM113Nn heteromers is shown as detected by autoradiography of a dried gel. The M113F subunits carried a D8 tail. Lane 1,
M113N7; lane 2, M113F7−nM113Nn (the heteromers formed from several preparations made with differing ratios of M113F and M113N subunits were mixed
to give roughly equal amounts of each subunit combination); lane 3, M113F7. A diagram of the eight different combinations of subunits and their permuta-
tions is shown to the right of the autoradiogram. The various permutations are not separated by electrophoresis. (C) Kinetics of the interaction of βCD with
single heteromeric αHL pores as determined by bilayer recording. Values of kon were calculated by using kon ¼ 1∕ðτon½βCDÞ, where τon is the mean interevent
interval. Values of koff were determined by using koff ¼ 1∕τoff, where τoff is the mean dwell time of βCD in the pore. Values of Kd were calculated by using
Kd ¼ koff∕kon. Each point represents the mean s:d: for three or more determinations. Where they cannot be seen, the s.d. values lie within the symbol. Black
squares, WT7−nM113Nn; gray squares, M113F7−nM113Nn; empty squares, M113F7−nWTn.
8166
∣
www.pnas.org/cgi/doi/10.1073/pnas.0914229107
Banerjee et al.
events had been observed previously with certain Met-113 repla-
cement mutants and may represent movement of the βCD at its
binding site (e.g., rotation about axes perpendicular to the C7
axis) (15). The additional current spikes were more prevalent
for M113V7 and M113Cpg7, which may take part in more con-
formationally labile interactions with βCD, compared with say
M113F7 (Fig. S4).
Interactions of βCD with Homoheptamers Bearing Aromatic Residues
at Position 113. To further understand the nature of the binding of
βCD to aromatic side chains, we examined the kinetics of βCD
binding to the homoheptamers containing f1F or f5F at position
113, M113f1F7 and M113f5F7 (Fig. 3C). For both mutants, the
value of kon was very similar to that of WT7, but the values of koff
and therefore Kd for M113f1F7 differed dramatically from WT7
and were close to the values for the tight-binding mutant M113F7
(Table S2A). By contrast, koff and Kd for M113f5F7 were similar
to the values for WT7 (Table S2A).
To determine whether M113f1F7 binds βCD in the same orien-
tation as M113F7 (Fig. 2C), we made heteromers of the M113f1F
subunit with M113N or M113F and examined M113F4M113f1F3
and M113N4M113f1F3. M113F4M113f1F3 binds βCD as strongly
as either M113F7 or M113f1F7, but M113N4M113f1F3 binds
βCD weakly with a similar affinity to WT7 (Fig. 3D and
Table S3). Therefore, it is reasonable to infer that M113F7
and M113f1F7 bind βCD in the same orientation with the 6-
hydroxyl groups of the CD in proximity to the aromatic rings
on the protein.
Cyclohexylalanine (Cha) was used to replace the aromatic side
chains with a roughly isosteric hydrophobic group. Again the va-
lue of kon for βCD was little changed, but koff for M113Cha7 had
an intermediate value of 42 6 s−1. Therefore, M113Cha7 binds
βCD more weakly than M113F7 but distinctly more strongly than
the WT7 pore (Table S2A and Fig. 3C).
Interactions of βCD with Homoheptamers Bearing Hydrophobic Resi-
dues at Position 113. M113V7 binds βCD very strongly, and there-
fore we compared αHL pores with Cpg or Cpa at position 113.
Cpg is roughly isosteric with Val, and like Val has a β-branched
side chain. Gratifyingly, M113Cpg7 has a kon value similar to the
other αHL pores, and koff and Kd values close to those of
M113V7 (Table S2B and Fig. 3E). Cyclopropylalanine (Cpa), with
an additional methylene group compared to Cpg, is roughly
isosteric with Leu, a weak binder, and M113Cpa7 also binds
βCD weakly with kon, koff and Kd values similar to those of
WT7 (Table S2B and Fig. 3E). M113I7 and M113T7, which are
β-branched, are also weak binders, but Ile and Thr are less closely
related to Val than Cpg.
To determine whether M113V7 binds βCD in the same orien-
tation as M113F7 or M113N7 (Fig. 2), we made heteromers of
M113V and the M113N or M113F subunits. M113V3M113F4,
M113V4M113F3, M113V3M113N4, and M113V4M113N3 were
examined in detail. All four heteroheptamers bound βCD more
weakly than M113V7, M113F7 or M113N7 (Fig. 3F and Table S4),
suggesting that Val at position 113 interacts with βCD strongly but
in a different manner to either Phe or Asn. Each heteromer
exhibited a range of Kd values, perhaps reflecting the various pos-
sible permutations of the two different subunits around the cen-
tral axis of the heptamer, although this heterogeneity was not
seen for heteromers made from WT, M113F and M113N (Fig. 1).
Discussion
Soon after we discovered that βCD binds to the WT-αHL pore for
around a millisecond, we found a mutant pore, M113N7, that re-
leases βCD ∼104 times more slowly (14). This prompted us to
examine all 19 mutants in which residue 113 is replaced by a nat-
ural amino acid, with the surprising result that a collection of ami-
no acids with structurally unrelated side chains (V, H, Y, D, N, F)
are tight binders (15). Here, we have examined the nature of the
binding interactions more closely by single-channel electrical re-
cording, protein engineering including unnatural amino acid mu-
tagenesis, and high-resolution x-ray crystallography, and we
provide the first definitive structural information on an engi-
neered protein nanopore.
We find that βCD can bind tightly to the αHL pore in three
different ways depending on the residue at 113, as exemplified
by Asn, Phe, and Val. Because Asn and Phe have quite different
side chains, we first compared the ability of M113N and M113F
subunits to take part in binding the CD. The examination of het-
eromeric proteins containing WT (Met-113), M113N and M113F
subunits showed that the replacement of WT subunits in WT7
with M113N or M113F subunits led to increased affinity for
βCD. The more M113N or M113F subunits that were added, the
tighter binding became. By contrast, when subunits in M113N7
were replaced with M113F subunits, binding became weaker,
reaching a minimum at three to four M113F subunits, and then
increasing in strength with five M113F subunits or more (Fig. 1C).
Parallel structural studies (30) revealed the basis of the “oppos-
ing” effects of the M113N and M113F subunits. βCD binds to
M113N7 in the opposite orientation to that in which it binds
to M113F7. In M113N7, the secondary hydroxyls in the βCD ring
are hydrogen bonded to Lys-147 and Asn-113 (Fig. 2A). By con-
trast, βCD interacts with M113F7 through its primary hydroxyl
face (Fig. 2B).
It seemed likely that M113V7, bound βCD in yet another way,
and this was examined by forming heteromers between M113V
and M113N or M113F. The presence of three or four subunits
of either M113N or M113F greatly decreases the affinity of
the pore for βCD (Fig. 3F), with an average koff of 7.3 × 101 s−1,
indicating that a third binding mode is indeed operating
Fig. 2.
X-ray structures of M113N and
M113F homoheptamers with βCD bound.
(A) Side view of heptameric αHL. βCD binds
in the blue highlighted region. (B) βCD
bound to M113N7 (dotted lines indicate hy-
drogen bonding). The side chains of Lys-147
are in pale brown and the side chains of Asn-
113 in yellow. (C) βCD bound to M113F7
(dotted lines indicate CH-π bonding). The
side chains of Phe-113 are in yellow. The sec-
ond βCD in the M113F7 · ðβCDÞ2 structure is
hydrogen bonded to the top βCD in a head-
to-head arrangement and has no apparent
interactions with the protein. For both (B)
and (C), four β strands were omitted from
the barrel to give a better view.
Banerjee et al.
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(Table S4). In summary, the three groups of tight-binding mutants
comprise αHL pores incorporating, at position 113: (i) the hydro-
gen-bonding amino acids N, D (the latter would have to be largely
in the protonated form), and possibly H; (ii) the aromatics F, Y,
f1F, and possibly H, and more weakly W; (iii) the β-branched ami-
no acids V, Cpg. There may be yet other means by which CDs can
bind to the αHL pore. For example, we earlier found that hepta-
6-sulfato-βCD can bind tightly to αHL pores containing the
N139Q mutation (37). Presumably, this CD is bound at a site low-
er down in the β barrel in a fashion that includes hydrogen bond-
ing to the Gln at position 139. While the various mutants
exhibited widely different koff values, the value of kon was almost
invariant and averaged ∼2.3 × 105 M−1 s−1 (Table S2) (15). Ap-
parently, transport to the binding site is rate limiting, through
a route unaffected by mutagenesis.
koff increased precipitously with the addition of WTsubunits to
M113N7 (Fig. 1C). Crystal structures of M113N7 show that resi-
dues
111,
113,
and
147
are
reorganized
by
compari-
son with WT7 and then undergo a more limited rearrangement
when βCD binds (Fig. S5). For example, the side chain of
Lys-147 shifts position to form a bifurcated hydrogen bond with
a 3-hydroxyl group of βCD and the side chain carbonyl of an Asn-
113 (Fig. S6). Therefore, the side chains of residues 111, 113, and
147 might be in a variety of conformations in WT7−nM113Nn het-
eromers and offer less well preorganized binding sites for βCD
than they do in M113N7. Further, the intramolecular hydrogen
bonds of the secondary hydroxyls in βCD (38) must be disrupted
upon binding as both hydroxyls on each glucose ring form hydro-
gen bonds to the mutant subunits (Fig. 2B). Because the hydrogen
bonds that are broken in βCD are arranged in a circle, the break-
age of bonds involving a single glucose (three bonds in all) will be
relatively more disruptive than those involving adjoining glucose
residues or the entire circle. The overall binding cooperativity in
M113N7 could be attributed to enthalpic cooperativity outweigh-
ing entropic penalties to binding (39). Positive cooperativity has
been observed previously in fairly rigid model systems (40).
By contrast with M113N7, there is little movement of side
chains in ðM113FÞ7 by comparison with WT7 and little move-
ment, including Phe-113, upon binding βCD (Fig. S7A). Further,
the crystal structure of M113F7 · βCD suggests that each Phe re-
sidue interacts independently with the βCD through what appear
to be CH-π interactions (Fig. S7B). These interactions are ex-
pected to be weak and not strongly directional and hence offer
less enthalpic cooperativity, as supported by the B-factors (crys-
tallographic temperatures factors) at the primary βCD binding
site, which are between ∼40 and 50. Positive cooperativity is ob-
served, but it is less pronounced than in the case of M113N7
(Table S5). In the case of M113N7, the B-factors of the residues
that bind βCD are in the 20s implying that the βCD is more rigidly
held than it is in M113F7.
The binding of sugars to aromatic residues in proteins can in-
clude CH-π bonding (41) or OH-π bonding or a finely balanced
complement of both (42, 43). However, we have dismissed the
possibility of an OH-π interaction between Phe-113 and the
6-hydroxyl groups of βCD as the distance between the center
of the phenyl rings to the nearest hydroxyl oxygen is higher
(5.2 0.65 Å, n ¼ 7) than that expected for a favorable OH-π
interaction (33). While we propose that βCD binds to Phe-113
through a C-6 CH-π interaction (Fig. S7B), the distances between
the center of the Phe-113 ring and the nearest C-6 of βCD ob-
served in the M113F7 · βCD structure (4.66 0.24 Å, n ¼ 7)
are in the upper range of the expected distance for a strong inter-
action, which is ∼4.5 Å (33). The angle between the normal to the
aromatic rings and the line connecting the C-6 atoms to the aro-
matic midpoint is 8.0 5.6°, which is well within the expected
range (44). The measurements with M113f5F7 argue against a
hydrophobic interaction between Phe residues at position 113
and the βCD ring. In f5F, the hydrophobicity of the phenyl ring
is significantly increased (45) yet M113f5F7 binds βCD weakly,
like WT7 (Fig. 3C and Table S2A).
By contrast with F, f1F, Y and N, homomeric αHL pores with
f5F and W at position 113 bound βCD relatively weakly (Fig. 3C
and Table S2A). In the case of f5F, the powerful electron with-
drawing action of the five fluorine atoms leaves a highly increased
positive charge at the center of the ring (46, 47), mitigating
against a hydrogen-bonding interaction. The electron-rich Trp
Fig. 3.
Properties of pores containing natural and unnatural amino acid sub-
stitutions at position 113. The data were recorded at þ40 mV in 1.0 M NaCl,
10 mM sodium phosphate, pH 7.5. (A) Unnatural amino acids used in this
study: 4-fluorophenylalanine, f1F; pentafluorophenylalanine, f5F; cyclohex-
ylalanine, Cha; cyclopropylglycine, Cpg; cyclopropylalanine, Cpa. (B) Repre-
sentative
current
traces
from
single
homoheptameric
αHL
pores,
containing unnatural amino acids at position 113, in the presence of βCD.
βCD (40 μM final) was added to the trans chamber. Level 1, open pore current;
level 2, pore occupied by βCD. The broken line indicates zero current. (C) In-
teraction of βCD with homomeric αHL pores containing aromatic amino acids
at position 113. Kd values for the interaction between βCD and the αHL pore
were calculated by using Kd ¼ koff∕kon. Each column represents the mean
s:d: for 10 or more determinations: dark gray, natural amino acids; light gray,
unnatural amino acids. Data adapted from Gu and colleagues (15) are
marked (*). (D) Representative current traces from single-channel recordings
of βCD binding to M113F4M113f1F3 and M113N4M113f1F3. βCD (40 μM final)
was added to the trans chamber. The broken line indicates zero current. (E)
Interaction of βCD with homomeric αHL pores containing hydrophobic amino
acids at position 113. Kd values for the interaction between βCD and the αHL
pore were calculated by using Kd ¼ koff∕kon. Each column represents the
mean s:d: for ten or more determinations: dark gray, natural amino acids;
light gray, unnatural amino acids. Data adapted from Gu and colleagues (15)
are marked (*). (F) koff values for βCD from heteroheptamers formed with
M113F and M113V subunits and with M113N and M113V subunits. βCD
(40 μM final) was added to the trans chamber. The kon values for βCD for
all these mutants are similar, at ∼3 × 105 M−1 s−1. Empty square: average
koff values for the mutant (bar is s:d). Filled square: M113V3M113F4; filled
circle: M113V4M113F3; filled upright triangle: M113V3M113N4; filled in-
verted triangle: M113V4M113N3.
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Banerjee et al.
ring (44, 46, 47) should favor hydrogen bonding, but here we can-
not make a direct comparison with the crystal structure of
M113F7 as the indole ring is far larger than benzene. It is possible
that it cannot become oriented in the same manner and that it is
misaligned for hydrogen bonding.
Our experiments suggest that M113V7 and M113Cpg7 bind
βCD in a third way. In heteromers with M113V, both M113F
and M113N reduce the affinity of the pore for βCD suggesting that
neither the CH-π interaction with Phe-113 nor the hydrogen-
bonding interactions with Asn-113 and Lys-147 are compatible
with binding to Val. Close interactions of Val with glucose rings
have been noted previously (48). Therefore, we propose that the
Val side-chain interacts with the side of the glucose ring. This in-
teraction might occur in one or both orientations of the CD
ring (Fig. 4).
Conclusion
We provide structural information on engineered protein nano-
pores and describe three distinct ways in which βCD can bind
within the lumen of mutant αHL pores in atomic detail. Our re-
sults will be useful in several areas of basic science and biotech-
nology. By using host molecules lodged within the αHL pore,
host-guest interactions can be investigated in fine detail at the
single-molecule level (17, 49). The present work will now permit
us to examine binding events at a known face of a CD. The work
also provides information for designing new binding sites within
the lumen of the αHL pore (37) or within other β barrel proteins
(21, 50) and for using molecular design to devise ways in which to
covalently attach CDs within pores (13, 51). These areas impact
practical applications of nanopore technology including stochas-
tic sensing (8), single-molecule DNA sequencing (9, 12, 13, 52),
the use of nanoreactors for the observation of single-molecule
chemistry (10), and the construction of nano- and microdevices
(11, 53), as well as the design of CDs as therapeutic agents
(23, 24).
Methods
Full details of the experimental procedures can be found in SI Appendix.
Materials
L-Amino acids were obtained as follows: 4-fluorophenylalanine (f1F) (Fluka);
pentafluorophenylalanine (f5F) (PepTech Corp.); cyclopropylglycine (Cpg) (Ty-
ger); cyclopropylalanine (Cpa) (Tyger). 4-N-benzoyl-5′-O-(4,4′-dimethoxytri-
tyl)-2′-deoxycytidine-3′-[(2-cyanoethyl)-(N,N-diisopropyl)]-phosphoramidite
and bis(2-cyanoethyl)-N,N-diisopropylphosphoramidite for the synthesis of
pdCpA were purchased from Glen Research and Toronto Research Chemicals,
respectively.
Preparation
of
NVOC-Protected
Aminoacyl-pdCpA.
NVOC-protected
aminoacyl-pdCpAs were prepared as reported previously by reacting the
dinucleotide pdCpA with N-protected, carboxylic acid-activated, amino
acids (54–56).
Preparation of NVOC-Protected Aminoacyl-tRNA. NVOC-protected aminoacyl-
pdCpAs were ligated enzymatically with a truncated tRNA, prepared by using
methods described elsewhere (57, 58).
Genetic Constructs and Mutagenesis. All new αHL constructs were verified by
DNA sequencing. Details of each construct can be found in SI Appendix.
Synthesis, Assembly, and Purification of Mutant αHL pores. αHL monomers (WT
and mutants) were prepared in vitro by coupled transcription and translation
(IVTT) and assembled into homoheptamers on rabbit red blood cell
membranes followed by purification by SDS–PAGE as described earlier
(59). Heteroheptamers were prepared by mixing the two required DNAs
(one encoding an αHL with a D8 tail) before IVTT and then oligomerizing
the mixed translation products on rabbit red blood cell membranes. Pores
with the desired combinations of subunits were purified by SDS–PAGE (59).
Synthesis, Assembly, and Purification of αHL Mutants Containing Unnatural Ami-
no Acids. αHL polypeptides containing unnatural amino acids were synthe-
sized by IVTT in the presence of rabbit red blood cell membranes. The
plasmid with a stop codon (TAG) at position 113 was used. Deprotected ami-
noacyl-tRNAs (SI Appendix) were added to the IVTT mixtures. For heterohep-
tamers with subunits containing unnatural amino acids in combination with
M113N or M113F, monomers were first made, which were then coassembled
on rabbit red blood cell membranes and subsequently purified by SDS–PAGE.
Single-Channel Current Recordings in Planar Lipid Bilayers. (15, 60) Recordings
were made with 1.0 M NaCl, 10 mM sodium phosphate, pH 7.5, in both cham-
bers, at an applied potential of þ40 mV. Data were recorded at 22 2°C. The
bilayer was formed from 1,2-diphytanoyl-sn-glycero-phosphocholine (Avanti
Polar Lipids). Proteins were added to the cis chamber, and βCD to the trans
chamber. Single-channel currents were recorded with an Axopatch 200B
patch-clamp amplifier (Axon Instruments) and filtered at 2 kHz with a
built-in 4-pole low-pass Bessel Filter. The data were acquired at a sampling
rate of 10 kHz. For mutants that bind βCD strongly, the data were acquired
for at least 30 min and for weak-binding mutants for at least 10 min.
Kinetic Data Analysis. Current amplitude and dwell-time histograms were
made by using ClampFit 9.0. The mean dwell times, τoff, were determined
by fitting the dwell-time histograms to single exponentials. Values of kon
and koff were obtained by using the mean dwell times and mean interevent
intervals, as described previously (15, 60). This analysis assumes a binary in-
teraction, which was supported in all cases examined by the finding of only
one major blockade level and a single exponential distribution of dwell
times (τoff).
Fig. 4.
Molecular model showing the three classes of interaction between
the αHL pore and βCD identified in this work. The model identifies the region
of βCD responsible for each interaction (H atoms interacting with Phe-113 or
Asn-113 and Lys-147: gray). The first class of interaction is with aromatic
residues and involves the seven -CH2OH groups of the βCD. The second class
is typified by the interactions with Asn at position 113, which involve hydro-
gen-bonds to the secondary 2-hydroxyls of the βCD. Structural studies show
that this interaction is supported by hydrogen bonding between Lys-147 and
the secondary 3-hydroxyls of the βCD. Structural studies and experiments
with heteromers suggest that the βCD in M113F7 is in the opposite orienta-
tion to the βCD in M113N7, in support of the model shown here. As the inter-
action with Val is hydrophobic, it is not directional and βCD may not bind at
the same position inside the β barrel as it does in M113F7 or M113N7.
Banerjee et al.
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Protein Crystallography. Details can be found in SI Appendix. Protein Data
Bank: The coordinates and structure factors of the described structures have
been deposited with accession codes 3M2L ðM113F7Þ, 3M3R ðM113F7 · βCDÞ,
3M4D ðM113N7Þ, 3M4E ðM113N7 · βCDÞ.
ACKNOWLEDGMENTS. We thank Dennis Dougherty for the plasmid pTHG73.
This work was funded by a Royal Society Wolfson Research Merit Award
(to H.B.), the Medical Research Council (H.B.), the National Institutes of
Health (H.B.), and the Howard Hughes Medical Institute (E.G.).
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|
3M2R
|
Structural Insight into Methyl-Coenzyme M Reductase Chemistry using Coenzyme B Analogues
|
Structural Insight into Methyl-Coenzyme M Reductase Chemistry
using Coenzyme B Analogues,†,‡
Peder E. Cedervall§,#, Mishtu Dey||,#,⊥, Arwen R. Pearson§,⊥, Stephen W. Ragsdale||, and
Carrie M. Wilmot*,||,§
§ Department of Biochemistry, Molecular Biology and Biophysics, University of Minnesota,
Minneapolis, Minnesota 55455
|| Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109
Abstract
Methyl-coenzyme M reductase (MCR) catalyzes the final and rate-limiting step in methane
biogenesis; the reduction of methyl-coenzyme M (methyl-SCoM) by coenzyme B (CoBSH) to
methane and a heterodisulfide (CoBS-SCoM). Crystallographic studies show that the active site is
deeply buried within the enzyme, and contains a highly reduced nickel-tetrapyrrole, coenzyme
F430. Methyl-SCoM must enter the active site prior to CoBSH, as species derived from analogues
of methyl-SCoM are always observed bound to the F430 nickel in the deepest part of the 30 Å long
substrate channel that leads from the protein surface to the active site. The seven-carbon
mercaptoalkanoyl chain of CoBSH binds within a 16 Å predominantly hydrophobic part of the
channel close to F430, with the CoBSH thiolate lying closest to the nickel at a distance of 8.8 Å. It
has previously been suggested that binding of CoBSH initiates catalysis by inducing a
conformational change that moves methyl-SCoM closer to the nickel promoting cleavage of the C-
S bond of methyl-SCoM. In order to better understand the structural role of CoBSH early in the
MCR mechanism, we have determined crystal structures of MCR in complex with four different
CoBSH analogues; pentanoyl-, hexanoyl-, octanoyl- and nonanoyl- derivatives of CoBSH
(CoB5SH, CoB6SH, CoB8SH and CoB9SH respectively). The data presented here reveal that the
shorter CoB5SH mercaptoalkanoyl chain overlays with that of CoBSH, but terminates two units
short of the CoBSH thiolate position. In contrast, the mercaptoalkanoyl chain of CoB6SH adopts a
different conformation, such that its thiolate is coincident with the position of the CoBSH thiolate.
†This work was supported by a Department of Energy grant DE-FG02-08ER15931 to S.W.R and supplement to C.M.W. and a
Minnesota Partnership for Biotechnology and Medical Genomics grant SPAP-05-0013-P-FY06.
‡Co-ordinates and structure factors have been deposited in the Protein Data Bank as entries 3m1v (MCRHSCoM); 3m2r
(MCRCoB5SH); 3m2u (MCRCoB6SH); 3m2v (MCRCoB8SH); 3m30 (MCRCoB9SH).
*Address correspondence to: Carrie M. Wilmot, Tel: (612) 624-2406, wilmo004@umn.edu.
⊥Current addresses: Mishtu Dey, Department of Chemistry, Massachusetts Institute of Technology, 77 Massachusetts Ave.,
Cambridge, Massachusetts 02139; Arwen Pearson, Astbury Centre for Structural Molecular Biology, Institute for Molecular and
Cellular Biology, Astbury Building. Leeds, LS2 9JT, U.K.
#These authors contributed equally to this work.
Supporting Informational material is available free of charge via the Internet at http://pubs.acs.org and contains the following:
MATERIAL AND METHODS for single crystal UV-visible microspectrophotometry, X-ray photoreduction experiment and X-ray
crystallography of the MCR-heterodisulfide product complex (MCRCoBSH + methyl-SCoM); RESULTS AND DISCUSSION for
redox changes and MCRCoBSH + methyl-SCoM crystal structure; Table S1, X-ray Data Collection, Processing and Refinement
Statistics for MCRCoBSH + methyl-SCoM; Figure S1, use of Fo-Fc electron density in modelling MCRCoB8SH; Figure S2,
illustration of electron density quality of MCRCoB5SH; Figure S3, solution and single crystal UV-visible spectra; Figure S4,
modeling of CoB6SH and CoBSH into the electron density of MCRCoB6SH; Figure S5, partially occupied HSCoM in MCRCoB6SH;
Figure S6, the active site and substrate channel of MCRCoBSH + methyl-SCoM; Figure S7, alternative conformation of Valα482 in
MCRHSCoM; Figure S8, the two conformations of the gly-rich loop in MCRHSCoM; Figure S9, propagation of conformational
changes in MCRHSCoM; Figure S10, the two conformations of CoB9SH in MCRCoB9SH; Figure S11, EPR spectra of MCRred1
sample; Scheme S1, scheme of the characterized forms of MCR.
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Biochemistry. Author manuscript; available in PMC 2011 September 7.
Published in final edited form as:
Biochemistry. 2010 September 7; 49(35): 7683–7693. doi:10.1021/bi100458d.
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This is consistent with the observation that CoB6SH is a slow substrate. A labile water in the
substrate channel was found to be a sensitive indicator for the presence of CoBSH and HSCoM.
The longer CoB8SH and CoB9SH analogues can be accommodated in the active site through
exclusion of this water. These analogues react with Ni(III)-methyl; a proposed MCR catalytic
intermediate of methanogenesis. The CoB8SH thiolate is 2.6 Å closer to the nickel than that of
CoBSH, but the additional carbon of CoB9SH only decreases the nickel thiolate distance a further
0.3 Å. Although the analogues did not induce any structural changes in the substrate channel, the
thiolates appeared to preferentially bind at two distinct positions in the channel; one being the
previously observed CoBSH thiolate position, and the other being at a hydrophobic annulus of
residues that lines the channel proximal to the nickel.
INTRODUCTION
Methanogenic archaea are organisms that under strict anaerobic conditions derive energy by
reducing compounds such as carbon dioxide, methylamine, acetate and methanol, to
methane (1, 2). The global production of methane by these organisms is estimated at one
billion tons annually. Microbially produced methane is not only a potential source of
renewable energy but also a potent greenhouse gas, and as such study of this process has
environmental ramifications. In these microorganisms, methyl-coenzyme M reductase
(MCR)1 is the enzyme that catalyzes the final step in methanogenesis, in which the
substrates methyl-coenzyme M (methyl-SCoM, 2-(methylthio)ethanesulfonate) and
coenzyme B (CoBSH, N-7-mercaptoheptanoylthreonine phosphate) are converted to
methane and a heterodisulfide (CoBS-SCoM) (Scheme 1) (3).
MCR is a 300 kDa protein with six subunits arranged in a α2β2γ2 oligomer (4). The known
crystal structures show that MCR has two active sites approximately 50 Å apart that are
deeply buried within the enzyme (5). The active site pocket is comprised of residues from
subunits α, α′, β and γ, with a 30 Å long substrate channel leading to the enzyme surface
(Figure 1). At the heart of the active site pocket is coenzyme F430, which is a highly reduced
nickel-containing tetrapyrrole (6–8). Currently sixteen distinct enzymatic and complexed
states of MCR have been spectroscopically characterized (Supporting Information, Scheme
S1) (9–31). In the resting active state of the enzyme, denoted MCRred1, the redox-active
nickel of F430 is present in the Ni(I) state (9, 16, 32). MCR is extremely oxygen sensitive
and upon oxygen exposure the enzyme enters an inactive Ni(II) state, denoted MCRred1-silent
(6). In this state it cannot be converted back to the active Ni(I) form by any known reducing
agent making this a challenging system to study. Additional complications involve the tight
association of coenzymes to purified MCR that are not easily displaced as demonstrated by
X-ray crystallographic and kinetic studies (5, 33–35).
Despite the fact that MCR has been studied for decades, no true catalytic intermediate has
been observed, and the actual mechanism remains elusive. Currently three general
mechanistic schemes for the enzymatic reaction have been proposed, each of which posit
different chemistry to initiate catalysis. Mechanism I involves Ni(I) acting as a nucleophile
in an SN2-type reaction that generates Ni(III)-methyl as an intermediate (Scheme 2A) (35–
38). Mechanism II starts with methyl-SCoM undergoing homolytic cleavage at the Ni(I) to
generate a methyl radical and a Ni(II)-SCoM species (Scheme 2B) (39–41). A more recently
proposed mechanism III suggests protonation of coenzyme F430 promotes reductive
cleavage of the methyl-SCoM thioether bond (42).
1Abbreviations: MCR, Methyl-coenzyme M reductase; methyl-SCoM, methyl-coenzyme M; CoBSH, coenzyme B; HSCoM,
coenzyme M; CoBS-SCoM, heterodisulfide of coenzyme B and coenzyme M; APS, Advanced Photon Source; ASU, asymmetric unit;
BPS, bromopropanesulfonate.
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Due to the stringent requirement to exclude O2, the available MCR crystal structures are all
in the inactive Ni(II) state. All but one contain CoBSH and HSCoM (demethylated methyl-
SCoM, an inhibitor and substrate analogue) in the active site (PDB codes 1hbn, 1hbo, 1hbu,
1e6y, 1e6v) (5, 33, 34). Another crystal structure has bound heterodisulfide product, CoBS-
SCoM (MCRsilent, PDB code 1hbm, Scheme 1 and Supporting Information, Scheme S1) (5,
33). All these structures reveal that both substrates access the active site through the same
channel (Figure 1). The binding site of HSCoM (and presumably methyl-SCoM) is more
deeply buried within the enzyme, and so it must enter prior to CoBSH for productive
chemistry to occur. As binding of CoBSH in the absence of co-substrate would be
inhibitory, it was suggested that a conformational change upon methyl-SCoM binding might
lower the Kd for CoBSH, and thus promote an ordered mechanism. Furthermore it has been
suggested that CoBSH binding induces a conformational change that brings the methyl-
SCoM substrate into closer proximity to the nickel, and this promotes C-S bond cleavage.
To investigate the proposed structural role of CoBSH in initiating catalysis, we have solved
the X-ray crystal structures of MCR in complex with four different CoBSH analogues.
CoBSH has a heptanoyl moiety linked to the thiol group, and the analogues are pentanoyl-,
hexanoyl-, octanoyl- or nonanoyl-containing derivatives of CoBSH (CoB5SH, CoB6SH,
CoB8SH and CoB9SH respectively; Figure 2) (3, 35, 43–47). In addition, we present a
structure in which the substrate channel predominantly lacks either CoBSH or
heterodisulfide product.
MATERIALS AND METHODS
Materials
The organism Methanothemobacter marburgensis (catalog OCM82) was obtained from the
Oregon Collection of Methanogens (Portland, OR). All buffers and media reagents were
obtained from Sigma-Aldrich (St. Louis, MO). The gases N2 (99.98%), H2/CO2 (80%/20%),
and ultra high purity H2 (99.999%) were obtained from Cryogenic Gases (Grand Rapids,
MI). Ti(III) citrate solutions were prepared from a stock solution of 246 mM Ti(III) citrate,
which was synthesized by adding sodium citrate to Ti(III) trichloride (30 wt % solution in 2
N hydrochloric acid) (Acros Organics, Morris Plains, NJ) under anaerobic conditions and
adjusting pH to 7.0 with sodium bicarbonate (48). The concentration of Ti(III) citrate was
determined by titrating against a solution of methyl viologen.
Synthesis of methyl-SCoM, CoB5SH, CoB6SH, CoBSH, CoB8SH, and CoB9SH
Methyl-SCoM was prepared from HSCoM and methyl iodide (49). The homodisulfides,
CoB5S-SCoB5, CoB6S-SCoB6, CoBS-SCoB, CoB8S-SCoB8, CoB9S-SCoB9 were prepared
as described from 5-bromovaleric acid, 6-bromohexanoic acid (Sigma-Aldrich, St. Louis,
MO), 7-bromoheptanoic acid (Karl Industries, Aurora, OH), 8-bromooctanoic acid, and 9-
bromononanoic acid (Matrix Scientific, Columbia, SC), respectively (43, 46). The free thiol
forms of CoB5SH, CoB6SH, CoBSH, CoB8SH, and CoB9SH were generated by the
reduction of the homodisulfides as previously described (45). The purity of the CoBSH
analogues was determined by 1H NMR spectroscopy. All compounds synthesized were
stored in a Vacuum Atmospheres chamber maintained at an oxygen level below 1 ppm, as
monitored continually with an oxygen analyzer (model 317, Teledyne Analytical
Instruments, City of Industry, CA) until use.
M. marburgensis Growth and MCRred1 Purification
Buffer preparations and all manipulations were performed under strict anaerobic conditions
in a Vacuum Atmospheres chamber maintained at an oxygen level below 1 ppm, as
monitored continually with an oxygen analyzer (model 317, Teledyne Analytical
Instruments, City of Industry, CA). MCRred1 was isolated from M. marburgensis cultured on
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H2/CO2 (80%/20%) at 65 °C in a 14-L fermentor (New Brunswick Scientific Co., Inc. New
Brunswick, NJ). Culture media were prepared as previously described (20, 24). MCRred1
was generated in vivo and purified as described previously (20). The purification procedure
routinely generates 65–75% MCRred1 as determined by UV-visible and EPR spectroscopy.
Spectroscopy of MCR
UV-visible spectra of the Ni(I)-containing MCRred1 were recorded in the anaerobic chamber
using a spectrophotometer (Model USB4000-UV-VIS, Ocean Optics, Dunedin, FL). EPR
spectra were recorded on a Bruker EMX spectrometer (Bruker Biospin Corp., Billerica,
MA), equipped with an Oxford ITC4 temperature controller, a Hewlett-Packard Model 5340
automatic frequency counter and Bruker gaussmeter. The EPR spectroscopic parameters
included: temperature, 70 K; microwave power, 10 mW; microwave frequency, 9.43 GHz;
receiver gain, 2 × 104; modulation amplitude, 10.0 G; modulation frequency, 100 kHz.
Double integrations of the EPR spectra were performed and referenced to a 1 mM copper
perchlorate standard. The NMR data were acquired at 298 K on a Bruker Avance DRX 500
MHz instrument equipped with a TXI cryoprobe.
Preparation of MCRred1 for Crystallization
All crystallization experiments were performed in the anaerobic chamber in which MCR
was purified unless otherwise noted. MCRred1 was prepared in 50 mM Tris, pH 7.6 and
excess HSCoM was removed by buffer exchange using an Amicon Ultra centrifuge filter
with a 50 kDa cut-off membrane (Millipore). Typically, 2–3 ml of MCRred1 was exchanged
with 10–15 ml of 50 mM Tris, pH 7.6. The enzyme was concentrated to 500–600 μl, and
this process was repeated three times. The fraction of MCRred1 in the purified MCR sample
was calculated from the UV-visible spectrum using extinction coefficients of 27.0
mM−1cm−1 at 385 nm for Ni(I)-MCRred1, and 9.15 mM−1cm−1 at 420 nm for Ni(II)-
MCRred1-silent (20). The amount of MCRred1 in samples used for crystallization was
determined to be ~80% and the concentration of total enzyme used was in the range of about
120–150 μM (~32–40 mg/ml). All crystallization experiments were performed anaerobically
by incubating 2.0 μl of enzyme solution in 50 mM Tris, pH 7.5 and 2.0 μl of reservoir
solution (100 mM Hepes-Na, pH 7.3/7.5/8.0; 150 mM magnesium acetate (Mg(CH3COO)2),
and 20/22% (w/v) PEG 400) in a sitting drop over 1 ml reservoir solution at 9 ºC. Triangular
and rectangular prismatic crystals with a bright yellowish-green color confirmed the
presence of nickel coenzyme F430. The crystals grew to a size of approximately 100–200 μm
in 4–5 days. CoBSH-depleted crystals were obtained by incubating 2 μl of a reaction
mixture containing 139 μM MCRred1 and 13 mM HSCoM with 2 μl of reservoir solution
(100 mM Hepes-Na pH 7.5, 150 mM magnesium acetate (Mg(CH3COO)2), 22% PEG 400).
Crystals of MCR complexed with the CoBSH analogues were grown by co-crystallization.
The CoB5SH co-crystals were obtained by incubating 2μl enzyme solution containing 124
μM MCRred1, 10 mM methyl-SCoM and 1 mM CoB5SH with 2μl of reservoir solution (100
mM Hepes-Na, pH 7.5, 150 mM (Mg(CH3COO)2), 22 % PEG 400). The crystals with
bound CoB6SH and CoB9SH were obtained by co-crystallization of 1 mM of analogue with
142 μM MCRred1 and equilibrated with 2μl of reservoir solution (100 mM Hepes-Na, pH
7.5, 150 mM Mg(CH3COO)2, 20 % PEG 400 for CoB6SH and 100 mM Hepes-Na, pH 7.3,
150 mM Mg(CH3COO)2), 22 % PEG 400 for CoB9SH). Crystals were cryoprotected in
reservoir solution containing 25 % (v/v) PEG 400 by soaking for 2–5 minutes before
cryocooling in liquid nitrogen in the anaerobic chamber. Crystals of CoB8SH bound to MCR
were obtained by incubating 2 μl of a mixture of 119 μM MCRred1 and 1 mM CoB8SH with
2 μl of reservoir solution (100 mM Hepes-Na, pH 7.3, 150 mM Mg(CH3COO)2), 20 % PEG
400). Before cryoprotection, the crystals were soaked for 5–10 minutes in a 100 mM
solution of methyl iodide2. The methyl iodide solution used for soaking was prepared by
adding a concentrated stock of methanolic solution of methyl iodide to the reservoir
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solution. Soaked crystals were quickly cryoprotected as described above and cryocooled in
the anaerobic chamber.
X-ray Diffraction Data Collection, Processing and Refinement
X-ray diffraction data were collected at 100 K on a ADSC Quantum-315 detector at the APS
Beamline 14-BM-C (BioCARS). The wavelength of X-rays was 0.979 Å. Data were
processed using HKL2000 (50). As in the previous X-ray crystallographic studies, the
crystals belong to the monoclinic space group P21 (a = 82 Å, b = 118 Å, c = 122 Å, β = 92°),
with one MCR molecule (two active sites) per asymmetric unit (5, 33). For refinement,
REFMAC in the Collaborative Computational Project Number 4 (CCP4) program suite was
used (51). A random sample of 5 % of the data across all resolution shells was chosen to
check refinement progress through calculation of an Rfree. The same reflections were used to
calculate Rfree for all structures, thus preventing bias due to high structural identity. The
remaining reflections were used in refinement (Rwork). Model building was done using the
Crystallographic Object-Oriented Toolkit (COOT) (52). The diffraction data and their
models are designated as MCRCoBXSH, where X is the number of carbons in the alkanoyl
portion of the analogue. Library files in CCP4 for F430 and CoBSH were incorrect, and these
were modified in Monomer Library Sketcher in the CCP4 program suite by comparison with
schematic drawings from Grabarse et al. (33). Co-ordinate and CCP4 library files for the
different CoBSH analogues were created in Monomer Library Sketcher. The general model
building and refinement strategy for all structures was as follows. It was clear from the
electron density in the substrate channel and at the active site that mixtures of species were
present in all datasets. These could be visualized with Fo-Fc and Fo-Fo difference electron
density maps (Supporting Information, Figure S1). The known positions of CoBSH and
HSCoM from the published Ni(II)-MCR crystal structures (PDB codes 1hbn, 1hbo, 1hbu
(33)) were used as guides to determine which species could be present in each dataset, and
these were then simultaneously modeled into the electron density. By alteration of their
relative occupancies (in 10% increments) followed by refinement, the ratio of occupancy
between different species was determined using the assumption that the average B-factors
for all molecular species bound should be similar to that of F430 and adjacent well-ordered
protein atoms within the active site and substrate channel. The combinations of modeled
ligands were constantly reassessed throughout refinement based on the remaining difference
electron density. This included test refinements of different ligand combinations during the
latter stages, thus using the optimized phases to check whether a different combination of
ligands could also explain the electron density. Sensible chemical structures and
interactions, along with keeping the combined occupancies of sterically mutually exclusive
species ≤ 100%, were maintained throughout refinement. The model was finally accepted
when the difference electron density map was minimal and the B-factors for the models
converged.
In practice the first structure refined was that of MCRCoB5SH. Initial phases were generated
by difference Fourier using a previously determined crystal structure (PDB code 1mro (5))
but with all non-bonded molecules, including water, removed from the model except F430.
Initial rigid body refinement followed by restrained refinement of MCRCoB5SH reduced the
Rwork to 26.5%. After model building and subsequent rounds of restrained refinement the
Rwork was 14.3% (Rfree 16.6%). Of the five structures only the CoB5SH analogue is
completely coincident with CoBSH, and so particular care had to be used in teasing apart the
ratios of the two species in modeling the MCRCoB5SH electron density. This was done by
2Methyl iodide was added with the intention of creating a Ni(III)-methyl species, which CoB8SH stabilizes. This was not achieved,
but the diffraction quality of this crystal was significantly better than crystals co-crystallized with CoB8SH alone, and so has been
included in this study.
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initially refining 100% CoBSH in the substrate channel. Positive Fo-Fc difference density
located at the carbon where the shorter CoB5SH thiol might be expected to be indicated the
presence of a more electron-rich species than carbon, which is consistent with the presence
of the CoB5SH sulfur. The refinement converged at a model containing a 50:50 mix of
CoBSH and the analogue. However, positive Fo-Fc difference density was still present at the
position of the CoBSH thiol, therefore a water molecule was added to the CoB5SH model at
50% occupancy and upon refinement this accounted for the electron density. An illustration
of the electron density quality from this structure is shown in Supporting Information,
Figure S2. An HSCoM-, CoBSH- and CoBSH analogue-free version of the refined
MCRCoB5SH structure was used as the starting model to generate initial phases for the four
other structures. After the initial round of restrained refinement the Rwork for these structures
were reduced to 14.5–15.6 %.
RESULTS AND DISCUSSION
Crystal Structures of MCR
Five crystal structures were determined, four of which are in complex with CoBSH
analogues differing in the number of carbons atoms in the alkanoyl portion of the molecule.
CoBSH is an N-7-mercaptoheptanoyl-containing molecule, whereas the four CoBSH
analogues contain N-5-mercaptopentanoyl-, N-6-mercaptohexanoyl-, N-8-mercaptooctanoyl-
or N-9-mercaptononanoyl-moieties (Figure 2). The corresponding crystal structures are
designated as MCRCoBXSH, where X is the number of carbons in the alkanoyl portion of the
analogue. The other crystal structure is of MCRred1c-silent (MCR in the Ni(II) state in
complex with HSCoM, designated here as MCRHSCoM) that is CoBSH-depleted. The
datasets have resolutions in the range from 1.35 – 1.8 Å. Although the crystallizations were
set up with the MCR solution predominantly in the Ni(I)-MCRred1 state, by the time X-ray
diffraction data were collected they had been oxidized to the Ni(II)-MCRred1-silent state
(Supporting Information). Following data collection there was no evidence for
photoreduction of the Ni(II) back to Ni(I) in any of the crystals, as assessed by single crystal
UV-visible microspectrophotometry (Supporting Information and Figure S3). Attempts to
photoreduce the crystals using different wavelengths and temperatures were unsuccessful
(Supporting Information).
Overall, the resulting structures are very similar to each other and to the previously
published structures of MCR, with differences mainly localized to the active site and
substrate channel. The two active sites in the ASU were refined independently. Unless
otherwise stated there was no difference between them. All five datasets contain a mixture
of species bound to the enzyme. There is always a background of CoBSH and HSCoM,
which co-purify with MCR and cannot be fully removed by extensive buffer exchange or by
the addition of a CoBSH analogue. HSCoM is added during purification of MCR, as it
stabilizes the resting active Ni(I) state (unpublished data), and this leads to HSCoM
occupancies between 50–100% amongst the structures (Table 1). In contrast CoBSH, which
is not added during purification, has occupancies ranging from 30–50%. As these
confounding species have all been described at high occupancy in other crystallographic
studies, the structural data of interest could be isolated (5, 33). In each case, the additional
electron density could be explained by inclusion of the appropriate CoBXSH model used in
that experiment at 50% or higher occupancy. The resulting models, along with 2Fo-Fc
electron density, are shown in Figure 3. The Rwork for the final structures range from 13.0 to
15.0 % (Rfree 15.5 to 19.5 %). The X-ray data collection, processing, refinement and model
building statistics are given in Table 1.
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Analogues shorter than CoBSH; CoB5SH and CoB6SH
CoB5SH is two methylene groups shorter than CoBSH, the MCR substrate. The
MCRCoB5SH structure is to 1.35 Å resolution. As expected the pentanoyl chain follows the
path of the CoBSH heptanoyl carbons down the substrate channel, and thus its thiol is
positioned in the same place as the second carbon preceding the CoBSH thiol (Figure 3A
and 4). There are no published MCR kinetic studies using CoB5SH, but as it binds in the
substrate channel, it is likely to be an inhibitor.
CoB6SH is one methylene shorter than CoBSH, and is a slow substrate of MCR. In this case
the 1.4 Å resolution electron density of MCRCoB6SH indicates that the analogue
unexpectedly binds in the substrate channel such that its thiol is virtually in the same
position as the thiol of CoBSH (Figure 3B and 4). The hexanoyl chain is oriented so that it
takes a shorter route down the substrate channel between carbons 2 and 5 (with the carbonyl
carbon labeled as carbon 1) than CoBSH (Figure 4, Supporting Information, Figure S4).
This short-cut is not seen in any of the other CoBXSH complex crystal structures, but
presumably arises because this CoB6SH binding conformer is energetically more favorable,
although it is not clear from the structure why this might be the case. CoB6SH binds very
tightly to MCR, with an apparent Ki value of 0.1 μM (3).
Water structure in the absence of HSCoM
The electron density for the MCRCoB6SH crystal structure only supported the modeling of 50
% bound HSCoM. In the fraction of MCR molecules where HSCoM is absent, the HSCoM
binding site is occupied by a network of four water molecules (Supporting Information,
Figure S5). Two waters are positioned close to the absent sulfonate oxygen positions of
HSCoM. Based on the presence of positive difference electron density, a third water was
modeled ligated to the Ni, and refined to a distance of ~2 Å (2.0 Å and 2.1 Å in the two
active sites of the ASU) with no distance restraint imposed between the Ni and water. This
water is in a similar position as the Ni coordinating sulfonate oxygen of the heterodisulfide
product in MCRsilent (Supporting Information, Figure S6 and PDB codes 3m32, 1hbm) (5,
33). The fourth water was in the vicinity of the expected position of a bridging water (W1)
seen in other structures (Figure 1, 3A and 3C).
Water structure in the absence of CoBSH
The 1.45 Å resolution electron density obtained for MCRHSCoM indicates that the substrate
channel contains only 30 % CoBSH. Nine ordered waters (W1–W9), along with an acetate
ion from the crystallization solution occupy the channel, with the acetate positioned where
the phosphothreonine linkage of CoBSH would be (Figure 3C). Presumably 1–2 further
waters would replace the acetate under physiological conditions. Other than W3 and W7, the
waters form hydrogen bonds with protein (Figure 5). One water (W2) occupies the same site
as the CoBSH thiol. Presumably due to the loss of favorable interactions that exist when
CoBSH is present, the hydrophobic side-chain of Valα482 adopts a second conformation
modeled at 60 % occupancy (Supporting Information, Figure S7).
Position of the “bridging” water, W1
The equivalent of W1 has previously been observed in MCRred1-silent and MCRox1-silent
crystal structures where, in the presence of CoBSH and HSCoM, it is sited equidistant (3.2
Å) between the two coenzyme thiols (PDB codes: 1hbn, 1hbo, 1hbu) and thus been termed
the “bridging water” (Figure 1) (5, 33). However, in the MCRHSCoM structure, due to the
presence of the more polarized W2 water, W1 is displaced away from HSCoM to maximize
the hydrogen bond interaction with W2 (2.9 Å to W2; 3.5 Å to HSCoM thiol, Figure 5). In
the MCRCoB5SH structure that also contained W2, the electron density indicated that this
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repositioning of W1 towards W2 also occurred. In contrast, the MCRCoB6SH structure
contained 100% thiol at the CoBSH position, but a partial occupancy of HSCoM (50%). In
this case the electron density for W1 indicated it had moved towards the nickel to form an
optimal hydrogen bond with a Ni-ligating water that was only present in the absence of
HSCoM (3.7 Å to CoB(6)SH thiol; 3.0 Å to Ni-ligating water, Supporting Information,
Figure S5). In all structures reported here, W1 (if present) appears to be a sensitive indicator
of the relative electronegativity of the Ni-ligated atom to that occupying the position of the
CoBSH thiol, and was a useful check in the crystallographic modeling and refinement
process.
Flexibility in the substrate channel: Alternative protein conformers
The binding site of HSCoM (and presumably methyl-SCoM) is more deeply buried within
the enzyme, and so it must enter prior to CoBSH for productive chemistry to occur. As
binding of CoBSH in the absence of co-substrate would be inhibitory, it was suggested that
a conformational change upon methyl-SCoM binding might lower the Kd for CoBSH, and
thus promote an ordered mechanism. Compared to the 1hbn MCRox1-silent and 1hbu
MCRred1-silent crystal structures, which both have full occupancy HSCoM, the lower
occupancy of HSCoM in the 1hbo MCRred1-silent structure was associated with significantly
greater flexibility within the channel, and the ability to model a second conformation of a
Gly-rich amino acid stretch that formed part of the CoBSH channel. This suggested that
methyl-SCoM binding might cause the channel to become more ordered, increasing the
affinity of MCR for CoBSH by conformational restriction rather than a switch mechanism
where the structure reorganizes from one well-defined conformer to another (33). In the
MCRHSCoM data containing 30 % CoBSH and 100% HSCoM, the Fo-Fc difference electron
density map at one of the two independent active sites in the ASU contained positive peaks
that suggested the presence of an alternate conformation also involving this part of the
polypeptide (Supporting Information, Figure S8). Using this as a guide, a similar second
conformation involving seven contiguous amino acid residues of the same Gly-rich amino
acid stretch (β366–372) could be modeled and refined at 20 % occupancy leaving no
residual difference density. Parts of the α′ subunit (α′111–129 and α′237–242) that are in
close proximity to this stretch of amino acids also exhibit second conformations, with the
main-chain carbonyl of α′243 in van der Waals contact with the B ring of F430 tetrapyrrole
(Supporting Information, Figure S9). Modeling these at 20% occupancy accounted for the
weak positive Fo-Fc difference electron density peaks observed in these areas. The evidence
of alternate conformers in these areas lends support to the proposal that increased flexibility
in the substrate channel propagates through the protein (33).
The MCRCoB6SH crystal structure contains 50 % CoB6SH, 50% CoBSH and 50% HSCoM.
In this case there is no evidence of an alternate loop conformation in either active site of the
ASU. However, as CoBSH and CoB6SH combined are at 100% occupancy, it is not
surprising their favorable interactions with the substrate channel would reduce
conformational disorder, despite the partial occupancy of HSCoM.
Analogues longer than CoBSH; CoB8SH and CoB9SH
Both analogues could be accommodated in the MCR substrate channel (Figures 3D and 3E).
The electron density supported final models containing 50% CoB8SH for MCRCoB8SH (1.8
Å resolution) and 60 % CoB9SH for MCRCoB9SH (1.45 Å resolution). The phosphate head-
groups are in identical positions to those of CoBSH, CoB5SH, CoB6SH (Figure 4) (5, 33).
Both analogues follow the crystallographically observed chain path of bound CoBSH, with
the extra atoms displacing the W1 water and placing the thiols closer to the nickel (Figure
6). CoB9SH does have a second conformer that deviates from the CoBSH path, but the thiol
position for this conformer and the CoBSH-tracking conformer are identical (Figure 3E and
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Supporting Information, Figure S10). Interestingly, the thiol of CoB8SH is not coincident
with the CoB9SH carbon that precedes the CoB9SH thiol (Figure 6). CoB8SH is an MCR
inhibitor with an apparent Ki of 15 μM (3). CoB9SH has never been tested for inhibition of
MCR-catalyzed methane formation, but it is reasonable to assume that it would be an
inhibitor.
CoBXSH thiol-to-nickel spatial relationship
The position of CoBSH in previous crystal structures poses a conundrum (5, 33). In all the
proposed catalytic mechanisms, CoBSH must interact with species generated at the nickel.
Perplexingly, the sulfur of the CoBSH substrate is 8.8 Å from the Ni(II) in the MCRox1-silent
and both MCRred1-silent crystal structures, and 6.4 Å from the thiol of the substrate analogue
HSCoM (Figure 1). Modeling studies demonstrated that the addition of a methyl group to
HSCoM did not bridge this gap (35, 45, 53). Therefore, a conformational change has been
postulated that would enable CoBSH to penetrate deeper into the substrate channel, and thus
approach closer to any nickelbound species. The heterodisulfide product in the MCRsilent
crystal structure has the CoBSH portion in virtually the same place as in MCRox1-silent,
giving no clue to possible structural changes that might occur to facilitate CoBSH reacting
with nickel-associated intermediates (5, 33).
Trigonometry suggests that if the alkanoyl chain of CoBSH or its analogue is in an extended
conformation, each additional unit in the chain would lead to the thiol moving ~1.2 Å
towards the Ni. Until this study there have been no crystal structures of CoBSH analogues in
complex with MCR, so mechanistic studies using different chain length analogues of
CoBSH assumed that shorter analogues would trace the observed path of CoBSH, and
longer analogues would penetrate about ~1.2 Å deeper per additional chain unit into the
channel. In the case of the shortest analogue CoB5SH, it does indeed follow the path of
CoBSH, with the thiol of CoB5SH being 2.8 Å away from the thiol position of CoBSH.
However, due to the conformation CoBSH adopts when bound in the substrate channel, the
difference in the S-Ni distance is small; the CoB5SH thiol being only 0.5 Å farther from the
Ni than CoBSH (8.8 Å for CoBSH vs. 9.3 Å for CoB5SH) (Table 2). This is due to the
alkanoyl chain of CoBSH not being in an extended conformation from carbons 4 to 6
(carbon 1 is the carbonyl carbon). CoB6SH on the other hand adopts a conformation that
places its thiol in virtually the same position as the thiol of CoBSH (Figure 4 and Table 2).
This is consistent with CoB6SH being a substrate. However, the kcat is 1000-fold lower than
for CoBSH (3, 35) although its Km value (180 μM, Dey & Ragsdale, in preparation) is
similar to that of CoBSH (Km = 75 μM (3)). The reason for this may be that the shorter
alkanoyl chain may not enable the analogue thiol to approach the nickel close enough for
efficient catalysis, and thus explain why CoB6SH is such a poor substrate.
In the case of the longer CoBXSH analogues, the sulfur of CoB8SH is 2.6 Å closer to the Ni
ion of F430 than that of CoBSH, and 2.5 Å closer to the thiol of HSCoM (Figure 6 and Table
2). The CoB9SH molecule follows the path of CoBSH, and reaches only a little further into
the substrate channel than CoB8SH, with the CoB9SH thiol positioned 2.9 Å closer to the Ni
than the thiol of CoBSH (Figure 6 and Table 2). This is only 0.3 Å closer than the distance
observed for the CoB8SH thiol, even though they are non-coincident. The distance to the
thiol of HSCoM is 2.6 Å closer than that of the substrate, CoBSH; only 0.1 Å closer than the
CoB8SH thiol. The two analogue thiols are above an annular hydrophobic aromatic
environment created by Pheα330, Tyrα333, Pheα443, Pheβ361 and Tyrβ367 that lies
between them and F430 (Figure 6). As a result, penetrating further into the channel may be
energetically unfavorable, consistent with the small difference in relative distances between
the CoB8SH/CoB9SH thiols and the HSCoM thiol/F430 nickel. The annulus is proposed to
be catalytically important in positioning methyl-SCoM and stabilizing the methane product,
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and the tyrosines have been proposed to be proton donors associated with mechanism II
(Scheme 2B) (5, 33).
Thus, there appear to be three preferential distances for thiols (including that of HSCoM)
within the MCR substrate channel; HSCoM at 2.4 Å, CoB6SH/CoBSH at 8.7 – 8.8 Å and
CoB8SH/CoB9SH at 5.9 – 6.2 Å from the nickel of F430 (Table 2).
Recent ENDOR, high-field continuous and pulse EPR work has identified changes in nickel
co-ordination when CoBSH is added to MCRred1c (active Ni(I)-MCRred1 + HSCoM) (14,
15, 18, 31). This generates up to 50% MCRred2 which is comprised of two distinct nickel co-
ordination geometries; an axial MCRred2a formally assigned as a Ni(III)-hydride, and a
rhombic MCRred2r in which the thiol of HSCoM is a Ni(I) ligand (Supporting Information,
Scheme S1). Formation of MCRred2 could also be induced by addition of CoBSH substrate
analogues CoBS-CH3 and CoBS-CF3, which have a chain length one unit longer than
substrate CoBSH (18, 53). The CoBS-CF3 enabled 19F-ENDOR studies to be performed,
and demonstrated that following CoBS-CF3 addition the remaining MCRred1 species had
Ni(I)-19F distances of 6.2 – 7.7 Å. This distance range agreed with a CoBS-CF3 model
created using the CoBSH position observed in the MCRox1-silent crystal structure (53).
However, in the MCRred2 species the Ni(I)-19F distances had shortened indicating a
movement of ~ 2 Å towards the nickel (MCRred2a Ni(I)-19 F, 4.0 – 5.5 Å; MCRred2r
Ni(I)-19F, 4.5 – 5.7 Å). In the case of the CoB8SH analogue (the closest equivalent to CoBS-
CF3 used in this study) the thiol to Ni(II) distance lies between the distance ranges observed
in the CoBS-CF3 studies, and so the fluorine(s) of the CoBS-CF3 in MCRred2 might
penetrate a little further into the hydrophobic annulus in the MCRred2 species. As the
alkanoyl chain of CoBSH is not fully extended it could easily undergo a similar
conformation change to that observed in the MCRred2 state.
Reaction of MCR Ni(III)-alkyl species’ with CoB8SH and CoB9SH
The two longer CoBXSH analogues have been shown to undergo alkylation when reacted
with MCRPS, a [Ni(III)-alkyl ↔ Ni(II)-alkylsulfonate radical] formed from reaction of
Ni(I)-MCRred1 with bromopropanesulfonate (BPS) (Supporting Information, Scheme S1)
(20, 23, 30, 45, 54). BPS is a substrate of MCRred1 in a reaction that involves a rapid
CoBSH-independent nucleophilic attack by Ni(I) on BPS to displace bromide and generate
MCRPS at a rate ~60-fold faster than generation of methane from CoBSH and methyl-
HSCoM (20, 45). Certain thiols can eliminate the propylsulfonate to yield a thioether
product and regenerate MCRred1, although at a rate 1000-fold slower than methane
formation (45). Both CoB8SH and CoB9SH can react with MCRPS to regenerate MCRred1,
but CoBSH cannot. The overall second-order rate constant for the reactivation of MCR by
CoB8SH is 160 M−1s−1, whereas for CoB9SH the reaction is more sluggish (12 M−1s−1).
CoB9SH might be expected to be closer to the proximal Ni ligand. It was therefore proposed
that this caused steric interference and explained why CoB9SH was a poorer reactivator of
MCR than CoB8SH. Our study has shown that the thiols of these two analogues are placed
such that they are approximately the same distance (~3.7 Å) from the thiol of HSCoM
ligated to the Ni atom (Table 2). The Ni(II)-HSCoM bond is 2.4 Å, whereas an Ni(III)-alkyl
bond is expected to be ~2 Å (24, 33), thus indicating that a conformational change is
required to effect the nucleophilic attack of the CoB8SH and CoB9SH thiols on an alkyl-
bound species. It would thus appear that a conformational change, such as observed in
MCRred2, is required for this chemistry also (53).
A Ni(III)-alkyl species is akin to the first intermediate in mechanism I of MCR-catalyzed
methane formation, Ni(III)-methyl (MCRMe, Supporting Information, Scheme S1, Scheme
2A) (11, 27). MCRMe has been shown to be capable of generating MCRred1 and methyl-
SCoM upon addition of HSCoM (which is the reverse of mechanism I, step 1, Scheme 2A);
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similar chemistry to the observed formation of a thioether product from the Ni(III)-alkyl.
Further addition of CoBSH following HSCoM treatment of MCRMe led to methane and
heterodisulfide formation, the natural products of methanogenesis. Although this lends
credence to mechanism I, it should be noted that like MCRPS, MCRMe in these experiments
was generated artificially. However, the MCRCoB9SH crystal structure demonstrates that the
two additional methylene units in the alkanoyl chain c.f. CoBSH, do not necessarily translate
into direct interaction of the thiol with the nickel proximal ligand. However, this could
represent the favorable position for a CoBSH thiol interacting with the methyl group of
methyl-SCoM. Just as the alkanoyl chain of CoB6SH has a more extended conformation
than CoBSH in the substrate channel, CoBSH could also adopt a more extended
conformation so that its thiol was in a similar position as the thiol of CoB8SH, priming it for
reaction with a nickel bound species.
If a significant conformational change is required early in MCR-catalyzed chemistry, which
would be a requirement of mechanism I, catalysis may well involve a rearrangement of the
aromatic amino acid annulus due to the presence of the methyl of methyl-SCoM, and this
might enable deeper penetration of CoBSH into MCR (Figure 6). All the crystal structures in
this study, and those solved previously, are of the inactive Ni(II)-MCR, which disfavors
close approach to the nickel in the absence of Ni(I)-bound methyl-SCoM, even in the case of
CoB9SH.
Conclusion
The goal of this study was to induce structural changes within the substrate channel and
active site of MCR using analogues of coenzyme CoBSH. It was hoped that this would shed
light on the nature of conformational changes that have been proposed to occur in MCR
catalysis. We have shown that that the CoBXSH analogues do not lead to any significant
conformational changes within the context of inactive Ni(II)-MCR. Therefore, it may be that
methyl-SCoM is the key coenzyme, in combination with a nickel oxidation state of 1 (and
3), that triggers a conformational change bringing the thiol of CoBSH closer to the nickel.
Thus, the crystal structure of the Ni(I)-methyl-SCoM/MCR complex may be required to
structurally define the conformational changes required for MCR-mediated chemistry.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
X-ray data were collected at beam-line 14-BM-C and photoreduction studies at 14-ID-B, BioCARS at the
Advanced Photon Source (APS), Argonne National Laboratory, Argonne, IL. We thank Vukica Srajer and Yu-
Sheng Chen for valuable assistance during data collection. Use of the Advanced Photon Source was supported by
the U.S. Department of Energy, Basic Energy Sciences, Office of Science under Contract No. DE-
AC02-06CH11357. Use of the BioCARS Sector 14 was supported by the National Institutes of Health, National
Center for Research Resources, under grant number RR007707. Computer resources were provided by the Basic
Sciences Computing Laboratory of the University of Minnesota Supercomputing Institute, and we thank Can
Ergenekan for his support. We also thank Ed Hoeffner at the Kahlert Structural Biology Laboratory (KSBL) at the
University of Minnesota. Use of the KSBL was supported by a Minnesota Partnership for Biotechnology and a
Medical Genomics Grant SPAP-05-0013-P-FY06.
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53. Ebner S, Jaun B, Goenrich M, Thauer RK, Harmer J. Binding of coenzyme B induces a major
conformational change in the active site of methyl-coenzyme M reductase. J Am Chem Soc. 2010;
132:567–575. [PubMed: 20014831]
54. Goenrich M, Mahlert F, Duin EC, Bauer C, Jaun B, Thauer RK. Probing the reactivity of Ni in the
active site of methyl-coenzyme M reductase with substrate analogues. J Biol Inorg Chem. 2004;
9:691–705. [PubMed: 15365904]
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Figure 1.
The active site and substrate channel in the MCRox1-silent crystal structure (PDB code 1hbn)
(9). Coenzyme F430, CoBSH and HSCoM are drawn as stick colored by atom (carbon: dark
grey). The nickel is displayed as a green sphere, water as a red sphere. Interactions are
drawn as dashed lines, and the corresponding distance is indicted in Angstroms (Å). The
path of the substrate channel was defined in the absence of F430, CoBSH, HSCoM and
water, with the surface closest to the viewer cut away. The figure was generated using
PyMOL (http://www.pymol.org).
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Figure 2.
Drawing of CoBSH analogues; (A) N-5-mercaptopentanoylthreonine phosphate (CoB5SH);
(B) N-6-mercaptohexanoylthreonine phosphate (CoB6SH); (C) N-8-
mercaptooctanoylthreonine phosphate (CoB8SH); (D) N-9-mercaptononanoylthreonine
phosphate (CoB9SH).
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Figure 3.
The active sites and substrate channels of the MCR crystal structures.; (A) MCRCoB5SH; (B)
MCRCoB6SH; (C) MCRHSCoM; (D) MCRCoB8SH; (E) MCRCoB9SH. 2Fo-Fc electron density
map around the CoBXSH analogues, waters in the CoBSH binding part of the channel and
the acetate ion (contoured at 1σ) is shown as a blue mesh. The protein is drawn as cartoon.
CoBXSH and acetate are drawn as stick and colored by atom (carbon: CoB5SH orange;
CoB6SH pale yellow; CoB8SH light blue; CoB9SH magenta; acetate white). Coenzyme F430
and HSCoM are drawn as stick colored by atom (carbon: F430 dark grey; HSCoM medium
grey). The nickel is displayed as a green sphere, and waters as red spheres. The figure was
generated using PyMOL (http://www.pymol.org/).
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Figure 4.
Overlay of CoBSH (from PDB code 1hbn) and the different CoBSH analogues. CoBXSH are
drawn as stick with the thiol represented by a sphere, and colored CoB5SH orange; CoB6SH
pale yellow; CoBSH light green; CoB8SH light blue; CoB9SH magenta. The protein is
drawn as cartoon. Coenzyme F430 and HSCoM are drawn as stick colored by atom (carbon:
F430 dark grey; HSCoM medium grey). The nickel is displayed as a green sphere. The figure
was generated using PyMOL (http://www.pymol.org/).
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Figure 5.
Hydrogen bonding diagram for the water structure modeled in MCRHSCoM. The water
molecules are named as in Figure 3C (W1–W9); WA, WB and WC are water molecules that
are present in all structures (i.e. in concert with the substrate CoBSH and the CoBSH
analogues). Interactions between surrounding residues and the water molecules are drawn as
dashed lines, and the corresponding distance is indicated in Angstroms (Å).
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Figure 6.
Stereo image of the annulus of aromatic amino acids distal of coenzyme F430. The protein is
drawn as cartoon with the side-chains of the aromatic residues drawn as white stick.
CoBSH, (from PDB code 1hbn (9)), CoB8SH and CoB9SH are drawn as stick with the thiols
represented by spheres, and colored CoBSH light green; CoB8SH light blue; CoB9SH
magenta. Coenzyme F430 and HSCoM are drawn as stick colored by atom (carbon: F430
dark grey; HSCoM medium grey). The nickel is displayed as a green sphere. The figure was
generated using PyMOL (http://www.pymol.org/).
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Scheme 1.
Reaction catalyzed by methyl-coenzyme M reductase
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Scheme 2.
Two of the proposed catalytic mechanisms for methyl-coenzyme M reductase; (A)
mechanism I; (B) mechanism II.
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Table 1
X-ray Data Collection, Processing and Refinement Statistics
Data collection and processing statistics
Name of data set
MCRCoB5SH
MCRCoB6SH
MCRHSCoM
MCRCoB8SH
MCRCoB9SH
Measured reflections
1969388
2427498
1440665
1160543
1425506
Unique reflections
553755
446253
405349
211803
401701
Resolution (Å) a
50.0–1.30 (1.35–1.30)
50.0–1.40 (1.45–1.40)
50.0–1.45 (1.50–1.45)
50.0–1.80 (1.86–1.80)
50.0–1.45 (1.50–1.45)
Completeness (%) a
97.1 (78.1)
99.9 (100.0)
99.5 (99.7)
99.8 (100.0)
98.1 (95.4)
R-sym (%) a,b
5.5 (32.9)
7.3 (44.7)
6.2 (44.0)
8.4 (47.7)
5.6 (42.5)
I/σI a
22.3 (3.6)
20.4 (4.0)
20.2 (3.2)
21.8 (3.9)
24.3 (3.2)
Space group
P21
P21
P21
P21
P21
Refinement and model building statistics
Resolution (Å) a
20.49–1.30 (1.33–1.30)
19.89–1.40 (1.44–1.40)
20.15–1.45 (1.49–1.45)
19.93–1.80 (1.84–1.80)
20.07–1.45 (1.48–1.45)
No. of reflection in working set a
525817 (30239)
423854 (25833)
384868 (25791)
201128 (11193)
381474 (23611)
No. of reflection in test set a
27777 (1576)
22348 (1331)
20362 (1319)
10625 (557)
20163 (1210)
R-work (%) c
14.32
13.04
13.47
14.95
13.58
R-free (%) d
16.56
15.53
16.22
19.54
16.44
ESU (Å) R-work/R-free
0.044/0.046
0.049/0.051
0.056/0.059
0.121/0.119
0.057/0.060
No. protein atoms
20087
19960
20265
19750
20036
No. coenzyme atoms
218
220
180
224
272
No. ligand atoms
37
62
52
26
49
No. water molecules
2443
2352
2516
1893
2432
RMS
bond lengths (Å)
0.033
0.033
0.032
0.028
0.032
bond angles (deg.)
2.693
2.625
2.468
2.059
2.549
Ramachandran plot (%)
favored
97.8
97.5
97.6
97.2
97.7
allowed
2.1
2.4
2.3
2.7
2.1
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disallowed
0.1
0.1
0.1
0.1
0.1
Average B-factor (Å2)
protein
12.42
13.35
12.12
17.22
12.73
coenzymes
8.20
9.24
7.25
11.24
8.27
ligands
31.95
35.48
28.29
33.76
32.92
waters
22.95
24.89
23.85
26.79
24.09
over all
13.54
14.57
13.40
18.02
13.93
Occupancy of HSCoM per active site (%)e
90/90
50/50
100/100
90/90
90/85
Occupancy of CoBSH per active site (%) e
50/50
50/50
30/30
50/50
40/40
CoBSH analogue, occupancy per active site (%) e
CoB5SH, 50/50
CoB6SH, 50/50
CoB8SH, 50/50
CoB9SH, 60/60
Other molecule, occupancy per active site (%) e
Acetate, 70/70
aValues in brackets correspond to the highest resolution shell.
bR-sym = ΣhklΣNj=1|Ihkl−Ihkl(j)|/ΣhklN*Ihkl, sum over all reflections and all observations N, with Ihkl(j) intensity of the jth observation of reflection hkl and Ihkl mean intensity of the reflection hkl.
cR-work = Σ||Fo| − |Fc||/Σ|Fo|, where |Fo| = observed structure factor amplitude and |Fc| = calculated structure factor amplitude.
dR-free, R-factor based on 5% of the data excluded from refinement.
eOccupancy of model in each of the two crystallographically independent active sites in the ASU
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Table 2
Distances from analogue thiols.
CoBXS - SCoM distance (Å)
CoBXS - Ni distance (Å)
CoB5SH
7.11/7.11a
9.30/9.30
CoB6SH
6.26/6.26
8.70/8.70
CoB7SH (substrate) b
6.37/6.39
8.73/8.77
CoB8SH
3.75/3.78
6.16/6.17
CoB9SH
3.71/3.68
5.96/5.91
aDistances in the two crystallographically independent active sites in the ASU
bDistances in the 1.16 Å resolution MCRox1-silent structure (PDB code 1hbn)(33)
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|
3M2U
|
Structural Insight into Methyl-Coenzyme M Reductase Chemistry using Coenzyme B Analogues
|
Structural Insight into Methyl-Coenzyme M Reductase Chemistry
using Coenzyme B Analogues,†,‡
Peder E. Cedervall§,#, Mishtu Dey||,#,⊥, Arwen R. Pearson§,⊥, Stephen W. Ragsdale||, and
Carrie M. Wilmot*,||,§
§ Department of Biochemistry, Molecular Biology and Biophysics, University of Minnesota,
Minneapolis, Minnesota 55455
|| Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109
Abstract
Methyl-coenzyme M reductase (MCR) catalyzes the final and rate-limiting step in methane
biogenesis; the reduction of methyl-coenzyme M (methyl-SCoM) by coenzyme B (CoBSH) to
methane and a heterodisulfide (CoBS-SCoM). Crystallographic studies show that the active site is
deeply buried within the enzyme, and contains a highly reduced nickel-tetrapyrrole, coenzyme
F430. Methyl-SCoM must enter the active site prior to CoBSH, as species derived from analogues
of methyl-SCoM are always observed bound to the F430 nickel in the deepest part of the 30 Å long
substrate channel that leads from the protein surface to the active site. The seven-carbon
mercaptoalkanoyl chain of CoBSH binds within a 16 Å predominantly hydrophobic part of the
channel close to F430, with the CoBSH thiolate lying closest to the nickel at a distance of 8.8 Å. It
has previously been suggested that binding of CoBSH initiates catalysis by inducing a
conformational change that moves methyl-SCoM closer to the nickel promoting cleavage of the C-
S bond of methyl-SCoM. In order to better understand the structural role of CoBSH early in the
MCR mechanism, we have determined crystal structures of MCR in complex with four different
CoBSH analogues; pentanoyl-, hexanoyl-, octanoyl- and nonanoyl- derivatives of CoBSH
(CoB5SH, CoB6SH, CoB8SH and CoB9SH respectively). The data presented here reveal that the
shorter CoB5SH mercaptoalkanoyl chain overlays with that of CoBSH, but terminates two units
short of the CoBSH thiolate position. In contrast, the mercaptoalkanoyl chain of CoB6SH adopts a
different conformation, such that its thiolate is coincident with the position of the CoBSH thiolate.
†This work was supported by a Department of Energy grant DE-FG02-08ER15931 to S.W.R and supplement to C.M.W. and a
Minnesota Partnership for Biotechnology and Medical Genomics grant SPAP-05-0013-P-FY06.
‡Co-ordinates and structure factors have been deposited in the Protein Data Bank as entries 3m1v (MCRHSCoM); 3m2r
(MCRCoB5SH); 3m2u (MCRCoB6SH); 3m2v (MCRCoB8SH); 3m30 (MCRCoB9SH).
*Address correspondence to: Carrie M. Wilmot, Tel: (612) 624-2406, wilmo004@umn.edu.
⊥Current addresses: Mishtu Dey, Department of Chemistry, Massachusetts Institute of Technology, 77 Massachusetts Ave.,
Cambridge, Massachusetts 02139; Arwen Pearson, Astbury Centre for Structural Molecular Biology, Institute for Molecular and
Cellular Biology, Astbury Building. Leeds, LS2 9JT, U.K.
#These authors contributed equally to this work.
Supporting Informational material is available free of charge via the Internet at http://pubs.acs.org and contains the following:
MATERIAL AND METHODS for single crystal UV-visible microspectrophotometry, X-ray photoreduction experiment and X-ray
crystallography of the MCR-heterodisulfide product complex (MCRCoBSH + methyl-SCoM); RESULTS AND DISCUSSION for
redox changes and MCRCoBSH + methyl-SCoM crystal structure; Table S1, X-ray Data Collection, Processing and Refinement
Statistics for MCRCoBSH + methyl-SCoM; Figure S1, use of Fo-Fc electron density in modelling MCRCoB8SH; Figure S2,
illustration of electron density quality of MCRCoB5SH; Figure S3, solution and single crystal UV-visible spectra; Figure S4,
modeling of CoB6SH and CoBSH into the electron density of MCRCoB6SH; Figure S5, partially occupied HSCoM in MCRCoB6SH;
Figure S6, the active site and substrate channel of MCRCoBSH + methyl-SCoM; Figure S7, alternative conformation of Valα482 in
MCRHSCoM; Figure S8, the two conformations of the gly-rich loop in MCRHSCoM; Figure S9, propagation of conformational
changes in MCRHSCoM; Figure S10, the two conformations of CoB9SH in MCRCoB9SH; Figure S11, EPR spectra of MCRred1
sample; Scheme S1, scheme of the characterized forms of MCR.
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This is consistent with the observation that CoB6SH is a slow substrate. A labile water in the
substrate channel was found to be a sensitive indicator for the presence of CoBSH and HSCoM.
The longer CoB8SH and CoB9SH analogues can be accommodated in the active site through
exclusion of this water. These analogues react with Ni(III)-methyl; a proposed MCR catalytic
intermediate of methanogenesis. The CoB8SH thiolate is 2.6 Å closer to the nickel than that of
CoBSH, but the additional carbon of CoB9SH only decreases the nickel thiolate distance a further
0.3 Å. Although the analogues did not induce any structural changes in the substrate channel, the
thiolates appeared to preferentially bind at two distinct positions in the channel; one being the
previously observed CoBSH thiolate position, and the other being at a hydrophobic annulus of
residues that lines the channel proximal to the nickel.
INTRODUCTION
Methanogenic archaea are organisms that under strict anaerobic conditions derive energy by
reducing compounds such as carbon dioxide, methylamine, acetate and methanol, to
methane (1, 2). The global production of methane by these organisms is estimated at one
billion tons annually. Microbially produced methane is not only a potential source of
renewable energy but also a potent greenhouse gas, and as such study of this process has
environmental ramifications. In these microorganisms, methyl-coenzyme M reductase
(MCR)1 is the enzyme that catalyzes the final step in methanogenesis, in which the
substrates methyl-coenzyme M (methyl-SCoM, 2-(methylthio)ethanesulfonate) and
coenzyme B (CoBSH, N-7-mercaptoheptanoylthreonine phosphate) are converted to
methane and a heterodisulfide (CoBS-SCoM) (Scheme 1) (3).
MCR is a 300 kDa protein with six subunits arranged in a α2β2γ2 oligomer (4). The known
crystal structures show that MCR has two active sites approximately 50 Å apart that are
deeply buried within the enzyme (5). The active site pocket is comprised of residues from
subunits α, α′, β and γ, with a 30 Å long substrate channel leading to the enzyme surface
(Figure 1). At the heart of the active site pocket is coenzyme F430, which is a highly reduced
nickel-containing tetrapyrrole (6–8). Currently sixteen distinct enzymatic and complexed
states of MCR have been spectroscopically characterized (Supporting Information, Scheme
S1) (9–31). In the resting active state of the enzyme, denoted MCRred1, the redox-active
nickel of F430 is present in the Ni(I) state (9, 16, 32). MCR is extremely oxygen sensitive
and upon oxygen exposure the enzyme enters an inactive Ni(II) state, denoted MCRred1-silent
(6). In this state it cannot be converted back to the active Ni(I) form by any known reducing
agent making this a challenging system to study. Additional complications involve the tight
association of coenzymes to purified MCR that are not easily displaced as demonstrated by
X-ray crystallographic and kinetic studies (5, 33–35).
Despite the fact that MCR has been studied for decades, no true catalytic intermediate has
been observed, and the actual mechanism remains elusive. Currently three general
mechanistic schemes for the enzymatic reaction have been proposed, each of which posit
different chemistry to initiate catalysis. Mechanism I involves Ni(I) acting as a nucleophile
in an SN2-type reaction that generates Ni(III)-methyl as an intermediate (Scheme 2A) (35–
38). Mechanism II starts with methyl-SCoM undergoing homolytic cleavage at the Ni(I) to
generate a methyl radical and a Ni(II)-SCoM species (Scheme 2B) (39–41). A more recently
proposed mechanism III suggests protonation of coenzyme F430 promotes reductive
cleavage of the methyl-SCoM thioether bond (42).
1Abbreviations: MCR, Methyl-coenzyme M reductase; methyl-SCoM, methyl-coenzyme M; CoBSH, coenzyme B; HSCoM,
coenzyme M; CoBS-SCoM, heterodisulfide of coenzyme B and coenzyme M; APS, Advanced Photon Source; ASU, asymmetric unit;
BPS, bromopropanesulfonate.
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Due to the stringent requirement to exclude O2, the available MCR crystal structures are all
in the inactive Ni(II) state. All but one contain CoBSH and HSCoM (demethylated methyl-
SCoM, an inhibitor and substrate analogue) in the active site (PDB codes 1hbn, 1hbo, 1hbu,
1e6y, 1e6v) (5, 33, 34). Another crystal structure has bound heterodisulfide product, CoBS-
SCoM (MCRsilent, PDB code 1hbm, Scheme 1 and Supporting Information, Scheme S1) (5,
33). All these structures reveal that both substrates access the active site through the same
channel (Figure 1). The binding site of HSCoM (and presumably methyl-SCoM) is more
deeply buried within the enzyme, and so it must enter prior to CoBSH for productive
chemistry to occur. As binding of CoBSH in the absence of co-substrate would be
inhibitory, it was suggested that a conformational change upon methyl-SCoM binding might
lower the Kd for CoBSH, and thus promote an ordered mechanism. Furthermore it has been
suggested that CoBSH binding induces a conformational change that brings the methyl-
SCoM substrate into closer proximity to the nickel, and this promotes C-S bond cleavage.
To investigate the proposed structural role of CoBSH in initiating catalysis, we have solved
the X-ray crystal structures of MCR in complex with four different CoBSH analogues.
CoBSH has a heptanoyl moiety linked to the thiol group, and the analogues are pentanoyl-,
hexanoyl-, octanoyl- or nonanoyl-containing derivatives of CoBSH (CoB5SH, CoB6SH,
CoB8SH and CoB9SH respectively; Figure 2) (3, 35, 43–47). In addition, we present a
structure in which the substrate channel predominantly lacks either CoBSH or
heterodisulfide product.
MATERIALS AND METHODS
Materials
The organism Methanothemobacter marburgensis (catalog OCM82) was obtained from the
Oregon Collection of Methanogens (Portland, OR). All buffers and media reagents were
obtained from Sigma-Aldrich (St. Louis, MO). The gases N2 (99.98%), H2/CO2 (80%/20%),
and ultra high purity H2 (99.999%) were obtained from Cryogenic Gases (Grand Rapids,
MI). Ti(III) citrate solutions were prepared from a stock solution of 246 mM Ti(III) citrate,
which was synthesized by adding sodium citrate to Ti(III) trichloride (30 wt % solution in 2
N hydrochloric acid) (Acros Organics, Morris Plains, NJ) under anaerobic conditions and
adjusting pH to 7.0 with sodium bicarbonate (48). The concentration of Ti(III) citrate was
determined by titrating against a solution of methyl viologen.
Synthesis of methyl-SCoM, CoB5SH, CoB6SH, CoBSH, CoB8SH, and CoB9SH
Methyl-SCoM was prepared from HSCoM and methyl iodide (49). The homodisulfides,
CoB5S-SCoB5, CoB6S-SCoB6, CoBS-SCoB, CoB8S-SCoB8, CoB9S-SCoB9 were prepared
as described from 5-bromovaleric acid, 6-bromohexanoic acid (Sigma-Aldrich, St. Louis,
MO), 7-bromoheptanoic acid (Karl Industries, Aurora, OH), 8-bromooctanoic acid, and 9-
bromononanoic acid (Matrix Scientific, Columbia, SC), respectively (43, 46). The free thiol
forms of CoB5SH, CoB6SH, CoBSH, CoB8SH, and CoB9SH were generated by the
reduction of the homodisulfides as previously described (45). The purity of the CoBSH
analogues was determined by 1H NMR spectroscopy. All compounds synthesized were
stored in a Vacuum Atmospheres chamber maintained at an oxygen level below 1 ppm, as
monitored continually with an oxygen analyzer (model 317, Teledyne Analytical
Instruments, City of Industry, CA) until use.
M. marburgensis Growth and MCRred1 Purification
Buffer preparations and all manipulations were performed under strict anaerobic conditions
in a Vacuum Atmospheres chamber maintained at an oxygen level below 1 ppm, as
monitored continually with an oxygen analyzer (model 317, Teledyne Analytical
Instruments, City of Industry, CA). MCRred1 was isolated from M. marburgensis cultured on
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H2/CO2 (80%/20%) at 65 °C in a 14-L fermentor (New Brunswick Scientific Co., Inc. New
Brunswick, NJ). Culture media were prepared as previously described (20, 24). MCRred1
was generated in vivo and purified as described previously (20). The purification procedure
routinely generates 65–75% MCRred1 as determined by UV-visible and EPR spectroscopy.
Spectroscopy of MCR
UV-visible spectra of the Ni(I)-containing MCRred1 were recorded in the anaerobic chamber
using a spectrophotometer (Model USB4000-UV-VIS, Ocean Optics, Dunedin, FL). EPR
spectra were recorded on a Bruker EMX spectrometer (Bruker Biospin Corp., Billerica,
MA), equipped with an Oxford ITC4 temperature controller, a Hewlett-Packard Model 5340
automatic frequency counter and Bruker gaussmeter. The EPR spectroscopic parameters
included: temperature, 70 K; microwave power, 10 mW; microwave frequency, 9.43 GHz;
receiver gain, 2 × 104; modulation amplitude, 10.0 G; modulation frequency, 100 kHz.
Double integrations of the EPR spectra were performed and referenced to a 1 mM copper
perchlorate standard. The NMR data were acquired at 298 K on a Bruker Avance DRX 500
MHz instrument equipped with a TXI cryoprobe.
Preparation of MCRred1 for Crystallization
All crystallization experiments were performed in the anaerobic chamber in which MCR
was purified unless otherwise noted. MCRred1 was prepared in 50 mM Tris, pH 7.6 and
excess HSCoM was removed by buffer exchange using an Amicon Ultra centrifuge filter
with a 50 kDa cut-off membrane (Millipore). Typically, 2–3 ml of MCRred1 was exchanged
with 10–15 ml of 50 mM Tris, pH 7.6. The enzyme was concentrated to 500–600 μl, and
this process was repeated three times. The fraction of MCRred1 in the purified MCR sample
was calculated from the UV-visible spectrum using extinction coefficients of 27.0
mM−1cm−1 at 385 nm for Ni(I)-MCRred1, and 9.15 mM−1cm−1 at 420 nm for Ni(II)-
MCRred1-silent (20). The amount of MCRred1 in samples used for crystallization was
determined to be ~80% and the concentration of total enzyme used was in the range of about
120–150 μM (~32–40 mg/ml). All crystallization experiments were performed anaerobically
by incubating 2.0 μl of enzyme solution in 50 mM Tris, pH 7.5 and 2.0 μl of reservoir
solution (100 mM Hepes-Na, pH 7.3/7.5/8.0; 150 mM magnesium acetate (Mg(CH3COO)2),
and 20/22% (w/v) PEG 400) in a sitting drop over 1 ml reservoir solution at 9 ºC. Triangular
and rectangular prismatic crystals with a bright yellowish-green color confirmed the
presence of nickel coenzyme F430. The crystals grew to a size of approximately 100–200 μm
in 4–5 days. CoBSH-depleted crystals were obtained by incubating 2 μl of a reaction
mixture containing 139 μM MCRred1 and 13 mM HSCoM with 2 μl of reservoir solution
(100 mM Hepes-Na pH 7.5, 150 mM magnesium acetate (Mg(CH3COO)2), 22% PEG 400).
Crystals of MCR complexed with the CoBSH analogues were grown by co-crystallization.
The CoB5SH co-crystals were obtained by incubating 2μl enzyme solution containing 124
μM MCRred1, 10 mM methyl-SCoM and 1 mM CoB5SH with 2μl of reservoir solution (100
mM Hepes-Na, pH 7.5, 150 mM (Mg(CH3COO)2), 22 % PEG 400). The crystals with
bound CoB6SH and CoB9SH were obtained by co-crystallization of 1 mM of analogue with
142 μM MCRred1 and equilibrated with 2μl of reservoir solution (100 mM Hepes-Na, pH
7.5, 150 mM Mg(CH3COO)2, 20 % PEG 400 for CoB6SH and 100 mM Hepes-Na, pH 7.3,
150 mM Mg(CH3COO)2), 22 % PEG 400 for CoB9SH). Crystals were cryoprotected in
reservoir solution containing 25 % (v/v) PEG 400 by soaking for 2–5 minutes before
cryocooling in liquid nitrogen in the anaerobic chamber. Crystals of CoB8SH bound to MCR
were obtained by incubating 2 μl of a mixture of 119 μM MCRred1 and 1 mM CoB8SH with
2 μl of reservoir solution (100 mM Hepes-Na, pH 7.3, 150 mM Mg(CH3COO)2), 20 % PEG
400). Before cryoprotection, the crystals were soaked for 5–10 minutes in a 100 mM
solution of methyl iodide2. The methyl iodide solution used for soaking was prepared by
adding a concentrated stock of methanolic solution of methyl iodide to the reservoir
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solution. Soaked crystals were quickly cryoprotected as described above and cryocooled in
the anaerobic chamber.
X-ray Diffraction Data Collection, Processing and Refinement
X-ray diffraction data were collected at 100 K on a ADSC Quantum-315 detector at the APS
Beamline 14-BM-C (BioCARS). The wavelength of X-rays was 0.979 Å. Data were
processed using HKL2000 (50). As in the previous X-ray crystallographic studies, the
crystals belong to the monoclinic space group P21 (a = 82 Å, b = 118 Å, c = 122 Å, β = 92°),
with one MCR molecule (two active sites) per asymmetric unit (5, 33). For refinement,
REFMAC in the Collaborative Computational Project Number 4 (CCP4) program suite was
used (51). A random sample of 5 % of the data across all resolution shells was chosen to
check refinement progress through calculation of an Rfree. The same reflections were used to
calculate Rfree for all structures, thus preventing bias due to high structural identity. The
remaining reflections were used in refinement (Rwork). Model building was done using the
Crystallographic Object-Oriented Toolkit (COOT) (52). The diffraction data and their
models are designated as MCRCoBXSH, where X is the number of carbons in the alkanoyl
portion of the analogue. Library files in CCP4 for F430 and CoBSH were incorrect, and these
were modified in Monomer Library Sketcher in the CCP4 program suite by comparison with
schematic drawings from Grabarse et al. (33). Co-ordinate and CCP4 library files for the
different CoBSH analogues were created in Monomer Library Sketcher. The general model
building and refinement strategy for all structures was as follows. It was clear from the
electron density in the substrate channel and at the active site that mixtures of species were
present in all datasets. These could be visualized with Fo-Fc and Fo-Fo difference electron
density maps (Supporting Information, Figure S1). The known positions of CoBSH and
HSCoM from the published Ni(II)-MCR crystal structures (PDB codes 1hbn, 1hbo, 1hbu
(33)) were used as guides to determine which species could be present in each dataset, and
these were then simultaneously modeled into the electron density. By alteration of their
relative occupancies (in 10% increments) followed by refinement, the ratio of occupancy
between different species was determined using the assumption that the average B-factors
for all molecular species bound should be similar to that of F430 and adjacent well-ordered
protein atoms within the active site and substrate channel. The combinations of modeled
ligands were constantly reassessed throughout refinement based on the remaining difference
electron density. This included test refinements of different ligand combinations during the
latter stages, thus using the optimized phases to check whether a different combination of
ligands could also explain the electron density. Sensible chemical structures and
interactions, along with keeping the combined occupancies of sterically mutually exclusive
species ≤ 100%, were maintained throughout refinement. The model was finally accepted
when the difference electron density map was minimal and the B-factors for the models
converged.
In practice the first structure refined was that of MCRCoB5SH. Initial phases were generated
by difference Fourier using a previously determined crystal structure (PDB code 1mro (5))
but with all non-bonded molecules, including water, removed from the model except F430.
Initial rigid body refinement followed by restrained refinement of MCRCoB5SH reduced the
Rwork to 26.5%. After model building and subsequent rounds of restrained refinement the
Rwork was 14.3% (Rfree 16.6%). Of the five structures only the CoB5SH analogue is
completely coincident with CoBSH, and so particular care had to be used in teasing apart the
ratios of the two species in modeling the MCRCoB5SH electron density. This was done by
2Methyl iodide was added with the intention of creating a Ni(III)-methyl species, which CoB8SH stabilizes. This was not achieved,
but the diffraction quality of this crystal was significantly better than crystals co-crystallized with CoB8SH alone, and so has been
included in this study.
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initially refining 100% CoBSH in the substrate channel. Positive Fo-Fc difference density
located at the carbon where the shorter CoB5SH thiol might be expected to be indicated the
presence of a more electron-rich species than carbon, which is consistent with the presence
of the CoB5SH sulfur. The refinement converged at a model containing a 50:50 mix of
CoBSH and the analogue. However, positive Fo-Fc difference density was still present at the
position of the CoBSH thiol, therefore a water molecule was added to the CoB5SH model at
50% occupancy and upon refinement this accounted for the electron density. An illustration
of the electron density quality from this structure is shown in Supporting Information,
Figure S2. An HSCoM-, CoBSH- and CoBSH analogue-free version of the refined
MCRCoB5SH structure was used as the starting model to generate initial phases for the four
other structures. After the initial round of restrained refinement the Rwork for these structures
were reduced to 14.5–15.6 %.
RESULTS AND DISCUSSION
Crystal Structures of MCR
Five crystal structures were determined, four of which are in complex with CoBSH
analogues differing in the number of carbons atoms in the alkanoyl portion of the molecule.
CoBSH is an N-7-mercaptoheptanoyl-containing molecule, whereas the four CoBSH
analogues contain N-5-mercaptopentanoyl-, N-6-mercaptohexanoyl-, N-8-mercaptooctanoyl-
or N-9-mercaptononanoyl-moieties (Figure 2). The corresponding crystal structures are
designated as MCRCoBXSH, where X is the number of carbons in the alkanoyl portion of the
analogue. The other crystal structure is of MCRred1c-silent (MCR in the Ni(II) state in
complex with HSCoM, designated here as MCRHSCoM) that is CoBSH-depleted. The
datasets have resolutions in the range from 1.35 – 1.8 Å. Although the crystallizations were
set up with the MCR solution predominantly in the Ni(I)-MCRred1 state, by the time X-ray
diffraction data were collected they had been oxidized to the Ni(II)-MCRred1-silent state
(Supporting Information). Following data collection there was no evidence for
photoreduction of the Ni(II) back to Ni(I) in any of the crystals, as assessed by single crystal
UV-visible microspectrophotometry (Supporting Information and Figure S3). Attempts to
photoreduce the crystals using different wavelengths and temperatures were unsuccessful
(Supporting Information).
Overall, the resulting structures are very similar to each other and to the previously
published structures of MCR, with differences mainly localized to the active site and
substrate channel. The two active sites in the ASU were refined independently. Unless
otherwise stated there was no difference between them. All five datasets contain a mixture
of species bound to the enzyme. There is always a background of CoBSH and HSCoM,
which co-purify with MCR and cannot be fully removed by extensive buffer exchange or by
the addition of a CoBSH analogue. HSCoM is added during purification of MCR, as it
stabilizes the resting active Ni(I) state (unpublished data), and this leads to HSCoM
occupancies between 50–100% amongst the structures (Table 1). In contrast CoBSH, which
is not added during purification, has occupancies ranging from 30–50%. As these
confounding species have all been described at high occupancy in other crystallographic
studies, the structural data of interest could be isolated (5, 33). In each case, the additional
electron density could be explained by inclusion of the appropriate CoBXSH model used in
that experiment at 50% or higher occupancy. The resulting models, along with 2Fo-Fc
electron density, are shown in Figure 3. The Rwork for the final structures range from 13.0 to
15.0 % (Rfree 15.5 to 19.5 %). The X-ray data collection, processing, refinement and model
building statistics are given in Table 1.
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Analogues shorter than CoBSH; CoB5SH and CoB6SH
CoB5SH is two methylene groups shorter than CoBSH, the MCR substrate. The
MCRCoB5SH structure is to 1.35 Å resolution. As expected the pentanoyl chain follows the
path of the CoBSH heptanoyl carbons down the substrate channel, and thus its thiol is
positioned in the same place as the second carbon preceding the CoBSH thiol (Figure 3A
and 4). There are no published MCR kinetic studies using CoB5SH, but as it binds in the
substrate channel, it is likely to be an inhibitor.
CoB6SH is one methylene shorter than CoBSH, and is a slow substrate of MCR. In this case
the 1.4 Å resolution electron density of MCRCoB6SH indicates that the analogue
unexpectedly binds in the substrate channel such that its thiol is virtually in the same
position as the thiol of CoBSH (Figure 3B and 4). The hexanoyl chain is oriented so that it
takes a shorter route down the substrate channel between carbons 2 and 5 (with the carbonyl
carbon labeled as carbon 1) than CoBSH (Figure 4, Supporting Information, Figure S4).
This short-cut is not seen in any of the other CoBXSH complex crystal structures, but
presumably arises because this CoB6SH binding conformer is energetically more favorable,
although it is not clear from the structure why this might be the case. CoB6SH binds very
tightly to MCR, with an apparent Ki value of 0.1 μM (3).
Water structure in the absence of HSCoM
The electron density for the MCRCoB6SH crystal structure only supported the modeling of 50
% bound HSCoM. In the fraction of MCR molecules where HSCoM is absent, the HSCoM
binding site is occupied by a network of four water molecules (Supporting Information,
Figure S5). Two waters are positioned close to the absent sulfonate oxygen positions of
HSCoM. Based on the presence of positive difference electron density, a third water was
modeled ligated to the Ni, and refined to a distance of ~2 Å (2.0 Å and 2.1 Å in the two
active sites of the ASU) with no distance restraint imposed between the Ni and water. This
water is in a similar position as the Ni coordinating sulfonate oxygen of the heterodisulfide
product in MCRsilent (Supporting Information, Figure S6 and PDB codes 3m32, 1hbm) (5,
33). The fourth water was in the vicinity of the expected position of a bridging water (W1)
seen in other structures (Figure 1, 3A and 3C).
Water structure in the absence of CoBSH
The 1.45 Å resolution electron density obtained for MCRHSCoM indicates that the substrate
channel contains only 30 % CoBSH. Nine ordered waters (W1–W9), along with an acetate
ion from the crystallization solution occupy the channel, with the acetate positioned where
the phosphothreonine linkage of CoBSH would be (Figure 3C). Presumably 1–2 further
waters would replace the acetate under physiological conditions. Other than W3 and W7, the
waters form hydrogen bonds with protein (Figure 5). One water (W2) occupies the same site
as the CoBSH thiol. Presumably due to the loss of favorable interactions that exist when
CoBSH is present, the hydrophobic side-chain of Valα482 adopts a second conformation
modeled at 60 % occupancy (Supporting Information, Figure S7).
Position of the “bridging” water, W1
The equivalent of W1 has previously been observed in MCRred1-silent and MCRox1-silent
crystal structures where, in the presence of CoBSH and HSCoM, it is sited equidistant (3.2
Å) between the two coenzyme thiols (PDB codes: 1hbn, 1hbo, 1hbu) and thus been termed
the “bridging water” (Figure 1) (5, 33). However, in the MCRHSCoM structure, due to the
presence of the more polarized W2 water, W1 is displaced away from HSCoM to maximize
the hydrogen bond interaction with W2 (2.9 Å to W2; 3.5 Å to HSCoM thiol, Figure 5). In
the MCRCoB5SH structure that also contained W2, the electron density indicated that this
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repositioning of W1 towards W2 also occurred. In contrast, the MCRCoB6SH structure
contained 100% thiol at the CoBSH position, but a partial occupancy of HSCoM (50%). In
this case the electron density for W1 indicated it had moved towards the nickel to form an
optimal hydrogen bond with a Ni-ligating water that was only present in the absence of
HSCoM (3.7 Å to CoB(6)SH thiol; 3.0 Å to Ni-ligating water, Supporting Information,
Figure S5). In all structures reported here, W1 (if present) appears to be a sensitive indicator
of the relative electronegativity of the Ni-ligated atom to that occupying the position of the
CoBSH thiol, and was a useful check in the crystallographic modeling and refinement
process.
Flexibility in the substrate channel: Alternative protein conformers
The binding site of HSCoM (and presumably methyl-SCoM) is more deeply buried within
the enzyme, and so it must enter prior to CoBSH for productive chemistry to occur. As
binding of CoBSH in the absence of co-substrate would be inhibitory, it was suggested that
a conformational change upon methyl-SCoM binding might lower the Kd for CoBSH, and
thus promote an ordered mechanism. Compared to the 1hbn MCRox1-silent and 1hbu
MCRred1-silent crystal structures, which both have full occupancy HSCoM, the lower
occupancy of HSCoM in the 1hbo MCRred1-silent structure was associated with significantly
greater flexibility within the channel, and the ability to model a second conformation of a
Gly-rich amino acid stretch that formed part of the CoBSH channel. This suggested that
methyl-SCoM binding might cause the channel to become more ordered, increasing the
affinity of MCR for CoBSH by conformational restriction rather than a switch mechanism
where the structure reorganizes from one well-defined conformer to another (33). In the
MCRHSCoM data containing 30 % CoBSH and 100% HSCoM, the Fo-Fc difference electron
density map at one of the two independent active sites in the ASU contained positive peaks
that suggested the presence of an alternate conformation also involving this part of the
polypeptide (Supporting Information, Figure S8). Using this as a guide, a similar second
conformation involving seven contiguous amino acid residues of the same Gly-rich amino
acid stretch (β366–372) could be modeled and refined at 20 % occupancy leaving no
residual difference density. Parts of the α′ subunit (α′111–129 and α′237–242) that are in
close proximity to this stretch of amino acids also exhibit second conformations, with the
main-chain carbonyl of α′243 in van der Waals contact with the B ring of F430 tetrapyrrole
(Supporting Information, Figure S9). Modeling these at 20% occupancy accounted for the
weak positive Fo-Fc difference electron density peaks observed in these areas. The evidence
of alternate conformers in these areas lends support to the proposal that increased flexibility
in the substrate channel propagates through the protein (33).
The MCRCoB6SH crystal structure contains 50 % CoB6SH, 50% CoBSH and 50% HSCoM.
In this case there is no evidence of an alternate loop conformation in either active site of the
ASU. However, as CoBSH and CoB6SH combined are at 100% occupancy, it is not
surprising their favorable interactions with the substrate channel would reduce
conformational disorder, despite the partial occupancy of HSCoM.
Analogues longer than CoBSH; CoB8SH and CoB9SH
Both analogues could be accommodated in the MCR substrate channel (Figures 3D and 3E).
The electron density supported final models containing 50% CoB8SH for MCRCoB8SH (1.8
Å resolution) and 60 % CoB9SH for MCRCoB9SH (1.45 Å resolution). The phosphate head-
groups are in identical positions to those of CoBSH, CoB5SH, CoB6SH (Figure 4) (5, 33).
Both analogues follow the crystallographically observed chain path of bound CoBSH, with
the extra atoms displacing the W1 water and placing the thiols closer to the nickel (Figure
6). CoB9SH does have a second conformer that deviates from the CoBSH path, but the thiol
position for this conformer and the CoBSH-tracking conformer are identical (Figure 3E and
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Supporting Information, Figure S10). Interestingly, the thiol of CoB8SH is not coincident
with the CoB9SH carbon that precedes the CoB9SH thiol (Figure 6). CoB8SH is an MCR
inhibitor with an apparent Ki of 15 μM (3). CoB9SH has never been tested for inhibition of
MCR-catalyzed methane formation, but it is reasonable to assume that it would be an
inhibitor.
CoBXSH thiol-to-nickel spatial relationship
The position of CoBSH in previous crystal structures poses a conundrum (5, 33). In all the
proposed catalytic mechanisms, CoBSH must interact with species generated at the nickel.
Perplexingly, the sulfur of the CoBSH substrate is 8.8 Å from the Ni(II) in the MCRox1-silent
and both MCRred1-silent crystal structures, and 6.4 Å from the thiol of the substrate analogue
HSCoM (Figure 1). Modeling studies demonstrated that the addition of a methyl group to
HSCoM did not bridge this gap (35, 45, 53). Therefore, a conformational change has been
postulated that would enable CoBSH to penetrate deeper into the substrate channel, and thus
approach closer to any nickelbound species. The heterodisulfide product in the MCRsilent
crystal structure has the CoBSH portion in virtually the same place as in MCRox1-silent,
giving no clue to possible structural changes that might occur to facilitate CoBSH reacting
with nickel-associated intermediates (5, 33).
Trigonometry suggests that if the alkanoyl chain of CoBSH or its analogue is in an extended
conformation, each additional unit in the chain would lead to the thiol moving ~1.2 Å
towards the Ni. Until this study there have been no crystal structures of CoBSH analogues in
complex with MCR, so mechanistic studies using different chain length analogues of
CoBSH assumed that shorter analogues would trace the observed path of CoBSH, and
longer analogues would penetrate about ~1.2 Å deeper per additional chain unit into the
channel. In the case of the shortest analogue CoB5SH, it does indeed follow the path of
CoBSH, with the thiol of CoB5SH being 2.8 Å away from the thiol position of CoBSH.
However, due to the conformation CoBSH adopts when bound in the substrate channel, the
difference in the S-Ni distance is small; the CoB5SH thiol being only 0.5 Å farther from the
Ni than CoBSH (8.8 Å for CoBSH vs. 9.3 Å for CoB5SH) (Table 2). This is due to the
alkanoyl chain of CoBSH not being in an extended conformation from carbons 4 to 6
(carbon 1 is the carbonyl carbon). CoB6SH on the other hand adopts a conformation that
places its thiol in virtually the same position as the thiol of CoBSH (Figure 4 and Table 2).
This is consistent with CoB6SH being a substrate. However, the kcat is 1000-fold lower than
for CoBSH (3, 35) although its Km value (180 μM, Dey & Ragsdale, in preparation) is
similar to that of CoBSH (Km = 75 μM (3)). The reason for this may be that the shorter
alkanoyl chain may not enable the analogue thiol to approach the nickel close enough for
efficient catalysis, and thus explain why CoB6SH is such a poor substrate.
In the case of the longer CoBXSH analogues, the sulfur of CoB8SH is 2.6 Å closer to the Ni
ion of F430 than that of CoBSH, and 2.5 Å closer to the thiol of HSCoM (Figure 6 and Table
2). The CoB9SH molecule follows the path of CoBSH, and reaches only a little further into
the substrate channel than CoB8SH, with the CoB9SH thiol positioned 2.9 Å closer to the Ni
than the thiol of CoBSH (Figure 6 and Table 2). This is only 0.3 Å closer than the distance
observed for the CoB8SH thiol, even though they are non-coincident. The distance to the
thiol of HSCoM is 2.6 Å closer than that of the substrate, CoBSH; only 0.1 Å closer than the
CoB8SH thiol. The two analogue thiols are above an annular hydrophobic aromatic
environment created by Pheα330, Tyrα333, Pheα443, Pheβ361 and Tyrβ367 that lies
between them and F430 (Figure 6). As a result, penetrating further into the channel may be
energetically unfavorable, consistent with the small difference in relative distances between
the CoB8SH/CoB9SH thiols and the HSCoM thiol/F430 nickel. The annulus is proposed to
be catalytically important in positioning methyl-SCoM and stabilizing the methane product,
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and the tyrosines have been proposed to be proton donors associated with mechanism II
(Scheme 2B) (5, 33).
Thus, there appear to be three preferential distances for thiols (including that of HSCoM)
within the MCR substrate channel; HSCoM at 2.4 Å, CoB6SH/CoBSH at 8.7 – 8.8 Å and
CoB8SH/CoB9SH at 5.9 – 6.2 Å from the nickel of F430 (Table 2).
Recent ENDOR, high-field continuous and pulse EPR work has identified changes in nickel
co-ordination when CoBSH is added to MCRred1c (active Ni(I)-MCRred1 + HSCoM) (14,
15, 18, 31). This generates up to 50% MCRred2 which is comprised of two distinct nickel co-
ordination geometries; an axial MCRred2a formally assigned as a Ni(III)-hydride, and a
rhombic MCRred2r in which the thiol of HSCoM is a Ni(I) ligand (Supporting Information,
Scheme S1). Formation of MCRred2 could also be induced by addition of CoBSH substrate
analogues CoBS-CH3 and CoBS-CF3, which have a chain length one unit longer than
substrate CoBSH (18, 53). The CoBS-CF3 enabled 19F-ENDOR studies to be performed,
and demonstrated that following CoBS-CF3 addition the remaining MCRred1 species had
Ni(I)-19F distances of 6.2 – 7.7 Å. This distance range agreed with a CoBS-CF3 model
created using the CoBSH position observed in the MCRox1-silent crystal structure (53).
However, in the MCRred2 species the Ni(I)-19F distances had shortened indicating a
movement of ~ 2 Å towards the nickel (MCRred2a Ni(I)-19 F, 4.0 – 5.5 Å; MCRred2r
Ni(I)-19F, 4.5 – 5.7 Å). In the case of the CoB8SH analogue (the closest equivalent to CoBS-
CF3 used in this study) the thiol to Ni(II) distance lies between the distance ranges observed
in the CoBS-CF3 studies, and so the fluorine(s) of the CoBS-CF3 in MCRred2 might
penetrate a little further into the hydrophobic annulus in the MCRred2 species. As the
alkanoyl chain of CoBSH is not fully extended it could easily undergo a similar
conformation change to that observed in the MCRred2 state.
Reaction of MCR Ni(III)-alkyl species’ with CoB8SH and CoB9SH
The two longer CoBXSH analogues have been shown to undergo alkylation when reacted
with MCRPS, a [Ni(III)-alkyl ↔ Ni(II)-alkylsulfonate radical] formed from reaction of
Ni(I)-MCRred1 with bromopropanesulfonate (BPS) (Supporting Information, Scheme S1)
(20, 23, 30, 45, 54). BPS is a substrate of MCRred1 in a reaction that involves a rapid
CoBSH-independent nucleophilic attack by Ni(I) on BPS to displace bromide and generate
MCRPS at a rate ~60-fold faster than generation of methane from CoBSH and methyl-
HSCoM (20, 45). Certain thiols can eliminate the propylsulfonate to yield a thioether
product and regenerate MCRred1, although at a rate 1000-fold slower than methane
formation (45). Both CoB8SH and CoB9SH can react with MCRPS to regenerate MCRred1,
but CoBSH cannot. The overall second-order rate constant for the reactivation of MCR by
CoB8SH is 160 M−1s−1, whereas for CoB9SH the reaction is more sluggish (12 M−1s−1).
CoB9SH might be expected to be closer to the proximal Ni ligand. It was therefore proposed
that this caused steric interference and explained why CoB9SH was a poorer reactivator of
MCR than CoB8SH. Our study has shown that the thiols of these two analogues are placed
such that they are approximately the same distance (~3.7 Å) from the thiol of HSCoM
ligated to the Ni atom (Table 2). The Ni(II)-HSCoM bond is 2.4 Å, whereas an Ni(III)-alkyl
bond is expected to be ~2 Å (24, 33), thus indicating that a conformational change is
required to effect the nucleophilic attack of the CoB8SH and CoB9SH thiols on an alkyl-
bound species. It would thus appear that a conformational change, such as observed in
MCRred2, is required for this chemistry also (53).
A Ni(III)-alkyl species is akin to the first intermediate in mechanism I of MCR-catalyzed
methane formation, Ni(III)-methyl (MCRMe, Supporting Information, Scheme S1, Scheme
2A) (11, 27). MCRMe has been shown to be capable of generating MCRred1 and methyl-
SCoM upon addition of HSCoM (which is the reverse of mechanism I, step 1, Scheme 2A);
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similar chemistry to the observed formation of a thioether product from the Ni(III)-alkyl.
Further addition of CoBSH following HSCoM treatment of MCRMe led to methane and
heterodisulfide formation, the natural products of methanogenesis. Although this lends
credence to mechanism I, it should be noted that like MCRPS, MCRMe in these experiments
was generated artificially. However, the MCRCoB9SH crystal structure demonstrates that the
two additional methylene units in the alkanoyl chain c.f. CoBSH, do not necessarily translate
into direct interaction of the thiol with the nickel proximal ligand. However, this could
represent the favorable position for a CoBSH thiol interacting with the methyl group of
methyl-SCoM. Just as the alkanoyl chain of CoB6SH has a more extended conformation
than CoBSH in the substrate channel, CoBSH could also adopt a more extended
conformation so that its thiol was in a similar position as the thiol of CoB8SH, priming it for
reaction with a nickel bound species.
If a significant conformational change is required early in MCR-catalyzed chemistry, which
would be a requirement of mechanism I, catalysis may well involve a rearrangement of the
aromatic amino acid annulus due to the presence of the methyl of methyl-SCoM, and this
might enable deeper penetration of CoBSH into MCR (Figure 6). All the crystal structures in
this study, and those solved previously, are of the inactive Ni(II)-MCR, which disfavors
close approach to the nickel in the absence of Ni(I)-bound methyl-SCoM, even in the case of
CoB9SH.
Conclusion
The goal of this study was to induce structural changes within the substrate channel and
active site of MCR using analogues of coenzyme CoBSH. It was hoped that this would shed
light on the nature of conformational changes that have been proposed to occur in MCR
catalysis. We have shown that that the CoBXSH analogues do not lead to any significant
conformational changes within the context of inactive Ni(II)-MCR. Therefore, it may be that
methyl-SCoM is the key coenzyme, in combination with a nickel oxidation state of 1 (and
3), that triggers a conformational change bringing the thiol of CoBSH closer to the nickel.
Thus, the crystal structure of the Ni(I)-methyl-SCoM/MCR complex may be required to
structurally define the conformational changes required for MCR-mediated chemistry.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
X-ray data were collected at beam-line 14-BM-C and photoreduction studies at 14-ID-B, BioCARS at the
Advanced Photon Source (APS), Argonne National Laboratory, Argonne, IL. We thank Vukica Srajer and Yu-
Sheng Chen for valuable assistance during data collection. Use of the Advanced Photon Source was supported by
the U.S. Department of Energy, Basic Energy Sciences, Office of Science under Contract No. DE-
AC02-06CH11357. Use of the BioCARS Sector 14 was supported by the National Institutes of Health, National
Center for Research Resources, under grant number RR007707. Computer resources were provided by the Basic
Sciences Computing Laboratory of the University of Minnesota Supercomputing Institute, and we thank Can
Ergenekan for his support. We also thank Ed Hoeffner at the Kahlert Structural Biology Laboratory (KSBL) at the
University of Minnesota. Use of the KSBL was supported by a Minnesota Partnership for Biotechnology and a
Medical Genomics Grant SPAP-05-0013-P-FY06.
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Figure 1.
The active site and substrate channel in the MCRox1-silent crystal structure (PDB code 1hbn)
(9). Coenzyme F430, CoBSH and HSCoM are drawn as stick colored by atom (carbon: dark
grey). The nickel is displayed as a green sphere, water as a red sphere. Interactions are
drawn as dashed lines, and the corresponding distance is indicted in Angstroms (Å). The
path of the substrate channel was defined in the absence of F430, CoBSH, HSCoM and
water, with the surface closest to the viewer cut away. The figure was generated using
PyMOL (http://www.pymol.org).
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Figure 2.
Drawing of CoBSH analogues; (A) N-5-mercaptopentanoylthreonine phosphate (CoB5SH);
(B) N-6-mercaptohexanoylthreonine phosphate (CoB6SH); (C) N-8-
mercaptooctanoylthreonine phosphate (CoB8SH); (D) N-9-mercaptononanoylthreonine
phosphate (CoB9SH).
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Figure 3.
The active sites and substrate channels of the MCR crystal structures.; (A) MCRCoB5SH; (B)
MCRCoB6SH; (C) MCRHSCoM; (D) MCRCoB8SH; (E) MCRCoB9SH. 2Fo-Fc electron density
map around the CoBXSH analogues, waters in the CoBSH binding part of the channel and
the acetate ion (contoured at 1σ) is shown as a blue mesh. The protein is drawn as cartoon.
CoBXSH and acetate are drawn as stick and colored by atom (carbon: CoB5SH orange;
CoB6SH pale yellow; CoB8SH light blue; CoB9SH magenta; acetate white). Coenzyme F430
and HSCoM are drawn as stick colored by atom (carbon: F430 dark grey; HSCoM medium
grey). The nickel is displayed as a green sphere, and waters as red spheres. The figure was
generated using PyMOL (http://www.pymol.org/).
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Figure 4.
Overlay of CoBSH (from PDB code 1hbn) and the different CoBSH analogues. CoBXSH are
drawn as stick with the thiol represented by a sphere, and colored CoB5SH orange; CoB6SH
pale yellow; CoBSH light green; CoB8SH light blue; CoB9SH magenta. The protein is
drawn as cartoon. Coenzyme F430 and HSCoM are drawn as stick colored by atom (carbon:
F430 dark grey; HSCoM medium grey). The nickel is displayed as a green sphere. The figure
was generated using PyMOL (http://www.pymol.org/).
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Figure 5.
Hydrogen bonding diagram for the water structure modeled in MCRHSCoM. The water
molecules are named as in Figure 3C (W1–W9); WA, WB and WC are water molecules that
are present in all structures (i.e. in concert with the substrate CoBSH and the CoBSH
analogues). Interactions between surrounding residues and the water molecules are drawn as
dashed lines, and the corresponding distance is indicated in Angstroms (Å).
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Figure 6.
Stereo image of the annulus of aromatic amino acids distal of coenzyme F430. The protein is
drawn as cartoon with the side-chains of the aromatic residues drawn as white stick.
CoBSH, (from PDB code 1hbn (9)), CoB8SH and CoB9SH are drawn as stick with the thiols
represented by spheres, and colored CoBSH light green; CoB8SH light blue; CoB9SH
magenta. Coenzyme F430 and HSCoM are drawn as stick colored by atom (carbon: F430
dark grey; HSCoM medium grey). The nickel is displayed as a green sphere. The figure was
generated using PyMOL (http://www.pymol.org/).
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Scheme 1.
Reaction catalyzed by methyl-coenzyme M reductase
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Scheme 2.
Two of the proposed catalytic mechanisms for methyl-coenzyme M reductase; (A)
mechanism I; (B) mechanism II.
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Table 1
X-ray Data Collection, Processing and Refinement Statistics
Data collection and processing statistics
Name of data set
MCRCoB5SH
MCRCoB6SH
MCRHSCoM
MCRCoB8SH
MCRCoB9SH
Measured reflections
1969388
2427498
1440665
1160543
1425506
Unique reflections
553755
446253
405349
211803
401701
Resolution (Å) a
50.0–1.30 (1.35–1.30)
50.0–1.40 (1.45–1.40)
50.0–1.45 (1.50–1.45)
50.0–1.80 (1.86–1.80)
50.0–1.45 (1.50–1.45)
Completeness (%) a
97.1 (78.1)
99.9 (100.0)
99.5 (99.7)
99.8 (100.0)
98.1 (95.4)
R-sym (%) a,b
5.5 (32.9)
7.3 (44.7)
6.2 (44.0)
8.4 (47.7)
5.6 (42.5)
I/σI a
22.3 (3.6)
20.4 (4.0)
20.2 (3.2)
21.8 (3.9)
24.3 (3.2)
Space group
P21
P21
P21
P21
P21
Refinement and model building statistics
Resolution (Å) a
20.49–1.30 (1.33–1.30)
19.89–1.40 (1.44–1.40)
20.15–1.45 (1.49–1.45)
19.93–1.80 (1.84–1.80)
20.07–1.45 (1.48–1.45)
No. of reflection in working set a
525817 (30239)
423854 (25833)
384868 (25791)
201128 (11193)
381474 (23611)
No. of reflection in test set a
27777 (1576)
22348 (1331)
20362 (1319)
10625 (557)
20163 (1210)
R-work (%) c
14.32
13.04
13.47
14.95
13.58
R-free (%) d
16.56
15.53
16.22
19.54
16.44
ESU (Å) R-work/R-free
0.044/0.046
0.049/0.051
0.056/0.059
0.121/0.119
0.057/0.060
No. protein atoms
20087
19960
20265
19750
20036
No. coenzyme atoms
218
220
180
224
272
No. ligand atoms
37
62
52
26
49
No. water molecules
2443
2352
2516
1893
2432
RMS
bond lengths (Å)
0.033
0.033
0.032
0.028
0.032
bond angles (deg.)
2.693
2.625
2.468
2.059
2.549
Ramachandran plot (%)
favored
97.8
97.5
97.6
97.2
97.7
allowed
2.1
2.4
2.3
2.7
2.1
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disallowed
0.1
0.1
0.1
0.1
0.1
Average B-factor (Å2)
protein
12.42
13.35
12.12
17.22
12.73
coenzymes
8.20
9.24
7.25
11.24
8.27
ligands
31.95
35.48
28.29
33.76
32.92
waters
22.95
24.89
23.85
26.79
24.09
over all
13.54
14.57
13.40
18.02
13.93
Occupancy of HSCoM per active site (%)e
90/90
50/50
100/100
90/90
90/85
Occupancy of CoBSH per active site (%) e
50/50
50/50
30/30
50/50
40/40
CoBSH analogue, occupancy per active site (%) e
CoB5SH, 50/50
CoB6SH, 50/50
CoB8SH, 50/50
CoB9SH, 60/60
Other molecule, occupancy per active site (%) e
Acetate, 70/70
aValues in brackets correspond to the highest resolution shell.
bR-sym = ΣhklΣNj=1|Ihkl−Ihkl(j)|/ΣhklN*Ihkl, sum over all reflections and all observations N, with Ihkl(j) intensity of the jth observation of reflection hkl and Ihkl mean intensity of the reflection hkl.
cR-work = Σ||Fo| − |Fc||/Σ|Fo|, where |Fo| = observed structure factor amplitude and |Fc| = calculated structure factor amplitude.
dR-free, R-factor based on 5% of the data excluded from refinement.
eOccupancy of model in each of the two crystallographically independent active sites in the ASU
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Table 2
Distances from analogue thiols.
CoBXS - SCoM distance (Å)
CoBXS - Ni distance (Å)
CoB5SH
7.11/7.11a
9.30/9.30
CoB6SH
6.26/6.26
8.70/8.70
CoB7SH (substrate) b
6.37/6.39
8.73/8.77
CoB8SH
3.75/3.78
6.16/6.17
CoB9SH
3.71/3.68
5.96/5.91
aDistances in the two crystallographically independent active sites in the ASU
bDistances in the 1.16 Å resolution MCRox1-silent structure (PDB code 1hbn)(33)
Biochemistry. Author manuscript; available in PMC 2011 September 7.
|
3M2V
|
Structural Insight into Methyl-Coenzyme M Reductase Chemistry using Coenzyme B Analogues
|
Structural Insight into Methyl-Coenzyme M Reductase Chemistry
using Coenzyme B Analogues,†,‡
Peder E. Cedervall§,#, Mishtu Dey||,#,⊥, Arwen R. Pearson§,⊥, Stephen W. Ragsdale||, and
Carrie M. Wilmot*,||,§
§ Department of Biochemistry, Molecular Biology and Biophysics, University of Minnesota,
Minneapolis, Minnesota 55455
|| Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109
Abstract
Methyl-coenzyme M reductase (MCR) catalyzes the final and rate-limiting step in methane
biogenesis; the reduction of methyl-coenzyme M (methyl-SCoM) by coenzyme B (CoBSH) to
methane and a heterodisulfide (CoBS-SCoM). Crystallographic studies show that the active site is
deeply buried within the enzyme, and contains a highly reduced nickel-tetrapyrrole, coenzyme
F430. Methyl-SCoM must enter the active site prior to CoBSH, as species derived from analogues
of methyl-SCoM are always observed bound to the F430 nickel in the deepest part of the 30 Å long
substrate channel that leads from the protein surface to the active site. The seven-carbon
mercaptoalkanoyl chain of CoBSH binds within a 16 Å predominantly hydrophobic part of the
channel close to F430, with the CoBSH thiolate lying closest to the nickel at a distance of 8.8 Å. It
has previously been suggested that binding of CoBSH initiates catalysis by inducing a
conformational change that moves methyl-SCoM closer to the nickel promoting cleavage of the C-
S bond of methyl-SCoM. In order to better understand the structural role of CoBSH early in the
MCR mechanism, we have determined crystal structures of MCR in complex with four different
CoBSH analogues; pentanoyl-, hexanoyl-, octanoyl- and nonanoyl- derivatives of CoBSH
(CoB5SH, CoB6SH, CoB8SH and CoB9SH respectively). The data presented here reveal that the
shorter CoB5SH mercaptoalkanoyl chain overlays with that of CoBSH, but terminates two units
short of the CoBSH thiolate position. In contrast, the mercaptoalkanoyl chain of CoB6SH adopts a
different conformation, such that its thiolate is coincident with the position of the CoBSH thiolate.
†This work was supported by a Department of Energy grant DE-FG02-08ER15931 to S.W.R and supplement to C.M.W. and a
Minnesota Partnership for Biotechnology and Medical Genomics grant SPAP-05-0013-P-FY06.
‡Co-ordinates and structure factors have been deposited in the Protein Data Bank as entries 3m1v (MCRHSCoM); 3m2r
(MCRCoB5SH); 3m2u (MCRCoB6SH); 3m2v (MCRCoB8SH); 3m30 (MCRCoB9SH).
*Address correspondence to: Carrie M. Wilmot, Tel: (612) 624-2406, wilmo004@umn.edu.
⊥Current addresses: Mishtu Dey, Department of Chemistry, Massachusetts Institute of Technology, 77 Massachusetts Ave.,
Cambridge, Massachusetts 02139; Arwen Pearson, Astbury Centre for Structural Molecular Biology, Institute for Molecular and
Cellular Biology, Astbury Building. Leeds, LS2 9JT, U.K.
#These authors contributed equally to this work.
Supporting Informational material is available free of charge via the Internet at http://pubs.acs.org and contains the following:
MATERIAL AND METHODS for single crystal UV-visible microspectrophotometry, X-ray photoreduction experiment and X-ray
crystallography of the MCR-heterodisulfide product complex (MCRCoBSH + methyl-SCoM); RESULTS AND DISCUSSION for
redox changes and MCRCoBSH + methyl-SCoM crystal structure; Table S1, X-ray Data Collection, Processing and Refinement
Statistics for MCRCoBSH + methyl-SCoM; Figure S1, use of Fo-Fc electron density in modelling MCRCoB8SH; Figure S2,
illustration of electron density quality of MCRCoB5SH; Figure S3, solution and single crystal UV-visible spectra; Figure S4,
modeling of CoB6SH and CoBSH into the electron density of MCRCoB6SH; Figure S5, partially occupied HSCoM in MCRCoB6SH;
Figure S6, the active site and substrate channel of MCRCoBSH + methyl-SCoM; Figure S7, alternative conformation of Valα482 in
MCRHSCoM; Figure S8, the two conformations of the gly-rich loop in MCRHSCoM; Figure S9, propagation of conformational
changes in MCRHSCoM; Figure S10, the two conformations of CoB9SH in MCRCoB9SH; Figure S11, EPR spectra of MCRred1
sample; Scheme S1, scheme of the characterized forms of MCR.
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Published in final edited form as:
Biochemistry. 2010 September 7; 49(35): 7683–7693. doi:10.1021/bi100458d.
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This is consistent with the observation that CoB6SH is a slow substrate. A labile water in the
substrate channel was found to be a sensitive indicator for the presence of CoBSH and HSCoM.
The longer CoB8SH and CoB9SH analogues can be accommodated in the active site through
exclusion of this water. These analogues react with Ni(III)-methyl; a proposed MCR catalytic
intermediate of methanogenesis. The CoB8SH thiolate is 2.6 Å closer to the nickel than that of
CoBSH, but the additional carbon of CoB9SH only decreases the nickel thiolate distance a further
0.3 Å. Although the analogues did not induce any structural changes in the substrate channel, the
thiolates appeared to preferentially bind at two distinct positions in the channel; one being the
previously observed CoBSH thiolate position, and the other being at a hydrophobic annulus of
residues that lines the channel proximal to the nickel.
INTRODUCTION
Methanogenic archaea are organisms that under strict anaerobic conditions derive energy by
reducing compounds such as carbon dioxide, methylamine, acetate and methanol, to
methane (1, 2). The global production of methane by these organisms is estimated at one
billion tons annually. Microbially produced methane is not only a potential source of
renewable energy but also a potent greenhouse gas, and as such study of this process has
environmental ramifications. In these microorganisms, methyl-coenzyme M reductase
(MCR)1 is the enzyme that catalyzes the final step in methanogenesis, in which the
substrates methyl-coenzyme M (methyl-SCoM, 2-(methylthio)ethanesulfonate) and
coenzyme B (CoBSH, N-7-mercaptoheptanoylthreonine phosphate) are converted to
methane and a heterodisulfide (CoBS-SCoM) (Scheme 1) (3).
MCR is a 300 kDa protein with six subunits arranged in a α2β2γ2 oligomer (4). The known
crystal structures show that MCR has two active sites approximately 50 Å apart that are
deeply buried within the enzyme (5). The active site pocket is comprised of residues from
subunits α, α′, β and γ, with a 30 Å long substrate channel leading to the enzyme surface
(Figure 1). At the heart of the active site pocket is coenzyme F430, which is a highly reduced
nickel-containing tetrapyrrole (6–8). Currently sixteen distinct enzymatic and complexed
states of MCR have been spectroscopically characterized (Supporting Information, Scheme
S1) (9–31). In the resting active state of the enzyme, denoted MCRred1, the redox-active
nickel of F430 is present in the Ni(I) state (9, 16, 32). MCR is extremely oxygen sensitive
and upon oxygen exposure the enzyme enters an inactive Ni(II) state, denoted MCRred1-silent
(6). In this state it cannot be converted back to the active Ni(I) form by any known reducing
agent making this a challenging system to study. Additional complications involve the tight
association of coenzymes to purified MCR that are not easily displaced as demonstrated by
X-ray crystallographic and kinetic studies (5, 33–35).
Despite the fact that MCR has been studied for decades, no true catalytic intermediate has
been observed, and the actual mechanism remains elusive. Currently three general
mechanistic schemes for the enzymatic reaction have been proposed, each of which posit
different chemistry to initiate catalysis. Mechanism I involves Ni(I) acting as a nucleophile
in an SN2-type reaction that generates Ni(III)-methyl as an intermediate (Scheme 2A) (35–
38). Mechanism II starts with methyl-SCoM undergoing homolytic cleavage at the Ni(I) to
generate a methyl radical and a Ni(II)-SCoM species (Scheme 2B) (39–41). A more recently
proposed mechanism III suggests protonation of coenzyme F430 promotes reductive
cleavage of the methyl-SCoM thioether bond (42).
1Abbreviations: MCR, Methyl-coenzyme M reductase; methyl-SCoM, methyl-coenzyme M; CoBSH, coenzyme B; HSCoM,
coenzyme M; CoBS-SCoM, heterodisulfide of coenzyme B and coenzyme M; APS, Advanced Photon Source; ASU, asymmetric unit;
BPS, bromopropanesulfonate.
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Due to the stringent requirement to exclude O2, the available MCR crystal structures are all
in the inactive Ni(II) state. All but one contain CoBSH and HSCoM (demethylated methyl-
SCoM, an inhibitor and substrate analogue) in the active site (PDB codes 1hbn, 1hbo, 1hbu,
1e6y, 1e6v) (5, 33, 34). Another crystal structure has bound heterodisulfide product, CoBS-
SCoM (MCRsilent, PDB code 1hbm, Scheme 1 and Supporting Information, Scheme S1) (5,
33). All these structures reveal that both substrates access the active site through the same
channel (Figure 1). The binding site of HSCoM (and presumably methyl-SCoM) is more
deeply buried within the enzyme, and so it must enter prior to CoBSH for productive
chemistry to occur. As binding of CoBSH in the absence of co-substrate would be
inhibitory, it was suggested that a conformational change upon methyl-SCoM binding might
lower the Kd for CoBSH, and thus promote an ordered mechanism. Furthermore it has been
suggested that CoBSH binding induces a conformational change that brings the methyl-
SCoM substrate into closer proximity to the nickel, and this promotes C-S bond cleavage.
To investigate the proposed structural role of CoBSH in initiating catalysis, we have solved
the X-ray crystal structures of MCR in complex with four different CoBSH analogues.
CoBSH has a heptanoyl moiety linked to the thiol group, and the analogues are pentanoyl-,
hexanoyl-, octanoyl- or nonanoyl-containing derivatives of CoBSH (CoB5SH, CoB6SH,
CoB8SH and CoB9SH respectively; Figure 2) (3, 35, 43–47). In addition, we present a
structure in which the substrate channel predominantly lacks either CoBSH or
heterodisulfide product.
MATERIALS AND METHODS
Materials
The organism Methanothemobacter marburgensis (catalog OCM82) was obtained from the
Oregon Collection of Methanogens (Portland, OR). All buffers and media reagents were
obtained from Sigma-Aldrich (St. Louis, MO). The gases N2 (99.98%), H2/CO2 (80%/20%),
and ultra high purity H2 (99.999%) were obtained from Cryogenic Gases (Grand Rapids,
MI). Ti(III) citrate solutions were prepared from a stock solution of 246 mM Ti(III) citrate,
which was synthesized by adding sodium citrate to Ti(III) trichloride (30 wt % solution in 2
N hydrochloric acid) (Acros Organics, Morris Plains, NJ) under anaerobic conditions and
adjusting pH to 7.0 with sodium bicarbonate (48). The concentration of Ti(III) citrate was
determined by titrating against a solution of methyl viologen.
Synthesis of methyl-SCoM, CoB5SH, CoB6SH, CoBSH, CoB8SH, and CoB9SH
Methyl-SCoM was prepared from HSCoM and methyl iodide (49). The homodisulfides,
CoB5S-SCoB5, CoB6S-SCoB6, CoBS-SCoB, CoB8S-SCoB8, CoB9S-SCoB9 were prepared
as described from 5-bromovaleric acid, 6-bromohexanoic acid (Sigma-Aldrich, St. Louis,
MO), 7-bromoheptanoic acid (Karl Industries, Aurora, OH), 8-bromooctanoic acid, and 9-
bromononanoic acid (Matrix Scientific, Columbia, SC), respectively (43, 46). The free thiol
forms of CoB5SH, CoB6SH, CoBSH, CoB8SH, and CoB9SH were generated by the
reduction of the homodisulfides as previously described (45). The purity of the CoBSH
analogues was determined by 1H NMR spectroscopy. All compounds synthesized were
stored in a Vacuum Atmospheres chamber maintained at an oxygen level below 1 ppm, as
monitored continually with an oxygen analyzer (model 317, Teledyne Analytical
Instruments, City of Industry, CA) until use.
M. marburgensis Growth and MCRred1 Purification
Buffer preparations and all manipulations were performed under strict anaerobic conditions
in a Vacuum Atmospheres chamber maintained at an oxygen level below 1 ppm, as
monitored continually with an oxygen analyzer (model 317, Teledyne Analytical
Instruments, City of Industry, CA). MCRred1 was isolated from M. marburgensis cultured on
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H2/CO2 (80%/20%) at 65 °C in a 14-L fermentor (New Brunswick Scientific Co., Inc. New
Brunswick, NJ). Culture media were prepared as previously described (20, 24). MCRred1
was generated in vivo and purified as described previously (20). The purification procedure
routinely generates 65–75% MCRred1 as determined by UV-visible and EPR spectroscopy.
Spectroscopy of MCR
UV-visible spectra of the Ni(I)-containing MCRred1 were recorded in the anaerobic chamber
using a spectrophotometer (Model USB4000-UV-VIS, Ocean Optics, Dunedin, FL). EPR
spectra were recorded on a Bruker EMX spectrometer (Bruker Biospin Corp., Billerica,
MA), equipped with an Oxford ITC4 temperature controller, a Hewlett-Packard Model 5340
automatic frequency counter and Bruker gaussmeter. The EPR spectroscopic parameters
included: temperature, 70 K; microwave power, 10 mW; microwave frequency, 9.43 GHz;
receiver gain, 2 × 104; modulation amplitude, 10.0 G; modulation frequency, 100 kHz.
Double integrations of the EPR spectra were performed and referenced to a 1 mM copper
perchlorate standard. The NMR data were acquired at 298 K on a Bruker Avance DRX 500
MHz instrument equipped with a TXI cryoprobe.
Preparation of MCRred1 for Crystallization
All crystallization experiments were performed in the anaerobic chamber in which MCR
was purified unless otherwise noted. MCRred1 was prepared in 50 mM Tris, pH 7.6 and
excess HSCoM was removed by buffer exchange using an Amicon Ultra centrifuge filter
with a 50 kDa cut-off membrane (Millipore). Typically, 2–3 ml of MCRred1 was exchanged
with 10–15 ml of 50 mM Tris, pH 7.6. The enzyme was concentrated to 500–600 μl, and
this process was repeated three times. The fraction of MCRred1 in the purified MCR sample
was calculated from the UV-visible spectrum using extinction coefficients of 27.0
mM−1cm−1 at 385 nm for Ni(I)-MCRred1, and 9.15 mM−1cm−1 at 420 nm for Ni(II)-
MCRred1-silent (20). The amount of MCRred1 in samples used for crystallization was
determined to be ~80% and the concentration of total enzyme used was in the range of about
120–150 μM (~32–40 mg/ml). All crystallization experiments were performed anaerobically
by incubating 2.0 μl of enzyme solution in 50 mM Tris, pH 7.5 and 2.0 μl of reservoir
solution (100 mM Hepes-Na, pH 7.3/7.5/8.0; 150 mM magnesium acetate (Mg(CH3COO)2),
and 20/22% (w/v) PEG 400) in a sitting drop over 1 ml reservoir solution at 9 ºC. Triangular
and rectangular prismatic crystals with a bright yellowish-green color confirmed the
presence of nickel coenzyme F430. The crystals grew to a size of approximately 100–200 μm
in 4–5 days. CoBSH-depleted crystals were obtained by incubating 2 μl of a reaction
mixture containing 139 μM MCRred1 and 13 mM HSCoM with 2 μl of reservoir solution
(100 mM Hepes-Na pH 7.5, 150 mM magnesium acetate (Mg(CH3COO)2), 22% PEG 400).
Crystals of MCR complexed with the CoBSH analogues were grown by co-crystallization.
The CoB5SH co-crystals were obtained by incubating 2μl enzyme solution containing 124
μM MCRred1, 10 mM methyl-SCoM and 1 mM CoB5SH with 2μl of reservoir solution (100
mM Hepes-Na, pH 7.5, 150 mM (Mg(CH3COO)2), 22 % PEG 400). The crystals with
bound CoB6SH and CoB9SH were obtained by co-crystallization of 1 mM of analogue with
142 μM MCRred1 and equilibrated with 2μl of reservoir solution (100 mM Hepes-Na, pH
7.5, 150 mM Mg(CH3COO)2, 20 % PEG 400 for CoB6SH and 100 mM Hepes-Na, pH 7.3,
150 mM Mg(CH3COO)2), 22 % PEG 400 for CoB9SH). Crystals were cryoprotected in
reservoir solution containing 25 % (v/v) PEG 400 by soaking for 2–5 minutes before
cryocooling in liquid nitrogen in the anaerobic chamber. Crystals of CoB8SH bound to MCR
were obtained by incubating 2 μl of a mixture of 119 μM MCRred1 and 1 mM CoB8SH with
2 μl of reservoir solution (100 mM Hepes-Na, pH 7.3, 150 mM Mg(CH3COO)2), 20 % PEG
400). Before cryoprotection, the crystals were soaked for 5–10 minutes in a 100 mM
solution of methyl iodide2. The methyl iodide solution used for soaking was prepared by
adding a concentrated stock of methanolic solution of methyl iodide to the reservoir
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solution. Soaked crystals were quickly cryoprotected as described above and cryocooled in
the anaerobic chamber.
X-ray Diffraction Data Collection, Processing and Refinement
X-ray diffraction data were collected at 100 K on a ADSC Quantum-315 detector at the APS
Beamline 14-BM-C (BioCARS). The wavelength of X-rays was 0.979 Å. Data were
processed using HKL2000 (50). As in the previous X-ray crystallographic studies, the
crystals belong to the monoclinic space group P21 (a = 82 Å, b = 118 Å, c = 122 Å, β = 92°),
with one MCR molecule (two active sites) per asymmetric unit (5, 33). For refinement,
REFMAC in the Collaborative Computational Project Number 4 (CCP4) program suite was
used (51). A random sample of 5 % of the data across all resolution shells was chosen to
check refinement progress through calculation of an Rfree. The same reflections were used to
calculate Rfree for all structures, thus preventing bias due to high structural identity. The
remaining reflections were used in refinement (Rwork). Model building was done using the
Crystallographic Object-Oriented Toolkit (COOT) (52). The diffraction data and their
models are designated as MCRCoBXSH, where X is the number of carbons in the alkanoyl
portion of the analogue. Library files in CCP4 for F430 and CoBSH were incorrect, and these
were modified in Monomer Library Sketcher in the CCP4 program suite by comparison with
schematic drawings from Grabarse et al. (33). Co-ordinate and CCP4 library files for the
different CoBSH analogues were created in Monomer Library Sketcher. The general model
building and refinement strategy for all structures was as follows. It was clear from the
electron density in the substrate channel and at the active site that mixtures of species were
present in all datasets. These could be visualized with Fo-Fc and Fo-Fo difference electron
density maps (Supporting Information, Figure S1). The known positions of CoBSH and
HSCoM from the published Ni(II)-MCR crystal structures (PDB codes 1hbn, 1hbo, 1hbu
(33)) were used as guides to determine which species could be present in each dataset, and
these were then simultaneously modeled into the electron density. By alteration of their
relative occupancies (in 10% increments) followed by refinement, the ratio of occupancy
between different species was determined using the assumption that the average B-factors
for all molecular species bound should be similar to that of F430 and adjacent well-ordered
protein atoms within the active site and substrate channel. The combinations of modeled
ligands were constantly reassessed throughout refinement based on the remaining difference
electron density. This included test refinements of different ligand combinations during the
latter stages, thus using the optimized phases to check whether a different combination of
ligands could also explain the electron density. Sensible chemical structures and
interactions, along with keeping the combined occupancies of sterically mutually exclusive
species ≤ 100%, were maintained throughout refinement. The model was finally accepted
when the difference electron density map was minimal and the B-factors for the models
converged.
In practice the first structure refined was that of MCRCoB5SH. Initial phases were generated
by difference Fourier using a previously determined crystal structure (PDB code 1mro (5))
but with all non-bonded molecules, including water, removed from the model except F430.
Initial rigid body refinement followed by restrained refinement of MCRCoB5SH reduced the
Rwork to 26.5%. After model building and subsequent rounds of restrained refinement the
Rwork was 14.3% (Rfree 16.6%). Of the five structures only the CoB5SH analogue is
completely coincident with CoBSH, and so particular care had to be used in teasing apart the
ratios of the two species in modeling the MCRCoB5SH electron density. This was done by
2Methyl iodide was added with the intention of creating a Ni(III)-methyl species, which CoB8SH stabilizes. This was not achieved,
but the diffraction quality of this crystal was significantly better than crystals co-crystallized with CoB8SH alone, and so has been
included in this study.
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initially refining 100% CoBSH in the substrate channel. Positive Fo-Fc difference density
located at the carbon where the shorter CoB5SH thiol might be expected to be indicated the
presence of a more electron-rich species than carbon, which is consistent with the presence
of the CoB5SH sulfur. The refinement converged at a model containing a 50:50 mix of
CoBSH and the analogue. However, positive Fo-Fc difference density was still present at the
position of the CoBSH thiol, therefore a water molecule was added to the CoB5SH model at
50% occupancy and upon refinement this accounted for the electron density. An illustration
of the electron density quality from this structure is shown in Supporting Information,
Figure S2. An HSCoM-, CoBSH- and CoBSH analogue-free version of the refined
MCRCoB5SH structure was used as the starting model to generate initial phases for the four
other structures. After the initial round of restrained refinement the Rwork for these structures
were reduced to 14.5–15.6 %.
RESULTS AND DISCUSSION
Crystal Structures of MCR
Five crystal structures were determined, four of which are in complex with CoBSH
analogues differing in the number of carbons atoms in the alkanoyl portion of the molecule.
CoBSH is an N-7-mercaptoheptanoyl-containing molecule, whereas the four CoBSH
analogues contain N-5-mercaptopentanoyl-, N-6-mercaptohexanoyl-, N-8-mercaptooctanoyl-
or N-9-mercaptononanoyl-moieties (Figure 2). The corresponding crystal structures are
designated as MCRCoBXSH, where X is the number of carbons in the alkanoyl portion of the
analogue. The other crystal structure is of MCRred1c-silent (MCR in the Ni(II) state in
complex with HSCoM, designated here as MCRHSCoM) that is CoBSH-depleted. The
datasets have resolutions in the range from 1.35 – 1.8 Å. Although the crystallizations were
set up with the MCR solution predominantly in the Ni(I)-MCRred1 state, by the time X-ray
diffraction data were collected they had been oxidized to the Ni(II)-MCRred1-silent state
(Supporting Information). Following data collection there was no evidence for
photoreduction of the Ni(II) back to Ni(I) in any of the crystals, as assessed by single crystal
UV-visible microspectrophotometry (Supporting Information and Figure S3). Attempts to
photoreduce the crystals using different wavelengths and temperatures were unsuccessful
(Supporting Information).
Overall, the resulting structures are very similar to each other and to the previously
published structures of MCR, with differences mainly localized to the active site and
substrate channel. The two active sites in the ASU were refined independently. Unless
otherwise stated there was no difference between them. All five datasets contain a mixture
of species bound to the enzyme. There is always a background of CoBSH and HSCoM,
which co-purify with MCR and cannot be fully removed by extensive buffer exchange or by
the addition of a CoBSH analogue. HSCoM is added during purification of MCR, as it
stabilizes the resting active Ni(I) state (unpublished data), and this leads to HSCoM
occupancies between 50–100% amongst the structures (Table 1). In contrast CoBSH, which
is not added during purification, has occupancies ranging from 30–50%. As these
confounding species have all been described at high occupancy in other crystallographic
studies, the structural data of interest could be isolated (5, 33). In each case, the additional
electron density could be explained by inclusion of the appropriate CoBXSH model used in
that experiment at 50% or higher occupancy. The resulting models, along with 2Fo-Fc
electron density, are shown in Figure 3. The Rwork for the final structures range from 13.0 to
15.0 % (Rfree 15.5 to 19.5 %). The X-ray data collection, processing, refinement and model
building statistics are given in Table 1.
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Analogues shorter than CoBSH; CoB5SH and CoB6SH
CoB5SH is two methylene groups shorter than CoBSH, the MCR substrate. The
MCRCoB5SH structure is to 1.35 Å resolution. As expected the pentanoyl chain follows the
path of the CoBSH heptanoyl carbons down the substrate channel, and thus its thiol is
positioned in the same place as the second carbon preceding the CoBSH thiol (Figure 3A
and 4). There are no published MCR kinetic studies using CoB5SH, but as it binds in the
substrate channel, it is likely to be an inhibitor.
CoB6SH is one methylene shorter than CoBSH, and is a slow substrate of MCR. In this case
the 1.4 Å resolution electron density of MCRCoB6SH indicates that the analogue
unexpectedly binds in the substrate channel such that its thiol is virtually in the same
position as the thiol of CoBSH (Figure 3B and 4). The hexanoyl chain is oriented so that it
takes a shorter route down the substrate channel between carbons 2 and 5 (with the carbonyl
carbon labeled as carbon 1) than CoBSH (Figure 4, Supporting Information, Figure S4).
This short-cut is not seen in any of the other CoBXSH complex crystal structures, but
presumably arises because this CoB6SH binding conformer is energetically more favorable,
although it is not clear from the structure why this might be the case. CoB6SH binds very
tightly to MCR, with an apparent Ki value of 0.1 μM (3).
Water structure in the absence of HSCoM
The electron density for the MCRCoB6SH crystal structure only supported the modeling of 50
% bound HSCoM. In the fraction of MCR molecules where HSCoM is absent, the HSCoM
binding site is occupied by a network of four water molecules (Supporting Information,
Figure S5). Two waters are positioned close to the absent sulfonate oxygen positions of
HSCoM. Based on the presence of positive difference electron density, a third water was
modeled ligated to the Ni, and refined to a distance of ~2 Å (2.0 Å and 2.1 Å in the two
active sites of the ASU) with no distance restraint imposed between the Ni and water. This
water is in a similar position as the Ni coordinating sulfonate oxygen of the heterodisulfide
product in MCRsilent (Supporting Information, Figure S6 and PDB codes 3m32, 1hbm) (5,
33). The fourth water was in the vicinity of the expected position of a bridging water (W1)
seen in other structures (Figure 1, 3A and 3C).
Water structure in the absence of CoBSH
The 1.45 Å resolution electron density obtained for MCRHSCoM indicates that the substrate
channel contains only 30 % CoBSH. Nine ordered waters (W1–W9), along with an acetate
ion from the crystallization solution occupy the channel, with the acetate positioned where
the phosphothreonine linkage of CoBSH would be (Figure 3C). Presumably 1–2 further
waters would replace the acetate under physiological conditions. Other than W3 and W7, the
waters form hydrogen bonds with protein (Figure 5). One water (W2) occupies the same site
as the CoBSH thiol. Presumably due to the loss of favorable interactions that exist when
CoBSH is present, the hydrophobic side-chain of Valα482 adopts a second conformation
modeled at 60 % occupancy (Supporting Information, Figure S7).
Position of the “bridging” water, W1
The equivalent of W1 has previously been observed in MCRred1-silent and MCRox1-silent
crystal structures where, in the presence of CoBSH and HSCoM, it is sited equidistant (3.2
Å) between the two coenzyme thiols (PDB codes: 1hbn, 1hbo, 1hbu) and thus been termed
the “bridging water” (Figure 1) (5, 33). However, in the MCRHSCoM structure, due to the
presence of the more polarized W2 water, W1 is displaced away from HSCoM to maximize
the hydrogen bond interaction with W2 (2.9 Å to W2; 3.5 Å to HSCoM thiol, Figure 5). In
the MCRCoB5SH structure that also contained W2, the electron density indicated that this
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repositioning of W1 towards W2 also occurred. In contrast, the MCRCoB6SH structure
contained 100% thiol at the CoBSH position, but a partial occupancy of HSCoM (50%). In
this case the electron density for W1 indicated it had moved towards the nickel to form an
optimal hydrogen bond with a Ni-ligating water that was only present in the absence of
HSCoM (3.7 Å to CoB(6)SH thiol; 3.0 Å to Ni-ligating water, Supporting Information,
Figure S5). In all structures reported here, W1 (if present) appears to be a sensitive indicator
of the relative electronegativity of the Ni-ligated atom to that occupying the position of the
CoBSH thiol, and was a useful check in the crystallographic modeling and refinement
process.
Flexibility in the substrate channel: Alternative protein conformers
The binding site of HSCoM (and presumably methyl-SCoM) is more deeply buried within
the enzyme, and so it must enter prior to CoBSH for productive chemistry to occur. As
binding of CoBSH in the absence of co-substrate would be inhibitory, it was suggested that
a conformational change upon methyl-SCoM binding might lower the Kd for CoBSH, and
thus promote an ordered mechanism. Compared to the 1hbn MCRox1-silent and 1hbu
MCRred1-silent crystal structures, which both have full occupancy HSCoM, the lower
occupancy of HSCoM in the 1hbo MCRred1-silent structure was associated with significantly
greater flexibility within the channel, and the ability to model a second conformation of a
Gly-rich amino acid stretch that formed part of the CoBSH channel. This suggested that
methyl-SCoM binding might cause the channel to become more ordered, increasing the
affinity of MCR for CoBSH by conformational restriction rather than a switch mechanism
where the structure reorganizes from one well-defined conformer to another (33). In the
MCRHSCoM data containing 30 % CoBSH and 100% HSCoM, the Fo-Fc difference electron
density map at one of the two independent active sites in the ASU contained positive peaks
that suggested the presence of an alternate conformation also involving this part of the
polypeptide (Supporting Information, Figure S8). Using this as a guide, a similar second
conformation involving seven contiguous amino acid residues of the same Gly-rich amino
acid stretch (β366–372) could be modeled and refined at 20 % occupancy leaving no
residual difference density. Parts of the α′ subunit (α′111–129 and α′237–242) that are in
close proximity to this stretch of amino acids also exhibit second conformations, with the
main-chain carbonyl of α′243 in van der Waals contact with the B ring of F430 tetrapyrrole
(Supporting Information, Figure S9). Modeling these at 20% occupancy accounted for the
weak positive Fo-Fc difference electron density peaks observed in these areas. The evidence
of alternate conformers in these areas lends support to the proposal that increased flexibility
in the substrate channel propagates through the protein (33).
The MCRCoB6SH crystal structure contains 50 % CoB6SH, 50% CoBSH and 50% HSCoM.
In this case there is no evidence of an alternate loop conformation in either active site of the
ASU. However, as CoBSH and CoB6SH combined are at 100% occupancy, it is not
surprising their favorable interactions with the substrate channel would reduce
conformational disorder, despite the partial occupancy of HSCoM.
Analogues longer than CoBSH; CoB8SH and CoB9SH
Both analogues could be accommodated in the MCR substrate channel (Figures 3D and 3E).
The electron density supported final models containing 50% CoB8SH for MCRCoB8SH (1.8
Å resolution) and 60 % CoB9SH for MCRCoB9SH (1.45 Å resolution). The phosphate head-
groups are in identical positions to those of CoBSH, CoB5SH, CoB6SH (Figure 4) (5, 33).
Both analogues follow the crystallographically observed chain path of bound CoBSH, with
the extra atoms displacing the W1 water and placing the thiols closer to the nickel (Figure
6). CoB9SH does have a second conformer that deviates from the CoBSH path, but the thiol
position for this conformer and the CoBSH-tracking conformer are identical (Figure 3E and
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Supporting Information, Figure S10). Interestingly, the thiol of CoB8SH is not coincident
with the CoB9SH carbon that precedes the CoB9SH thiol (Figure 6). CoB8SH is an MCR
inhibitor with an apparent Ki of 15 μM (3). CoB9SH has never been tested for inhibition of
MCR-catalyzed methane formation, but it is reasonable to assume that it would be an
inhibitor.
CoBXSH thiol-to-nickel spatial relationship
The position of CoBSH in previous crystal structures poses a conundrum (5, 33). In all the
proposed catalytic mechanisms, CoBSH must interact with species generated at the nickel.
Perplexingly, the sulfur of the CoBSH substrate is 8.8 Å from the Ni(II) in the MCRox1-silent
and both MCRred1-silent crystal structures, and 6.4 Å from the thiol of the substrate analogue
HSCoM (Figure 1). Modeling studies demonstrated that the addition of a methyl group to
HSCoM did not bridge this gap (35, 45, 53). Therefore, a conformational change has been
postulated that would enable CoBSH to penetrate deeper into the substrate channel, and thus
approach closer to any nickelbound species. The heterodisulfide product in the MCRsilent
crystal structure has the CoBSH portion in virtually the same place as in MCRox1-silent,
giving no clue to possible structural changes that might occur to facilitate CoBSH reacting
with nickel-associated intermediates (5, 33).
Trigonometry suggests that if the alkanoyl chain of CoBSH or its analogue is in an extended
conformation, each additional unit in the chain would lead to the thiol moving ~1.2 Å
towards the Ni. Until this study there have been no crystal structures of CoBSH analogues in
complex with MCR, so mechanistic studies using different chain length analogues of
CoBSH assumed that shorter analogues would trace the observed path of CoBSH, and
longer analogues would penetrate about ~1.2 Å deeper per additional chain unit into the
channel. In the case of the shortest analogue CoB5SH, it does indeed follow the path of
CoBSH, with the thiol of CoB5SH being 2.8 Å away from the thiol position of CoBSH.
However, due to the conformation CoBSH adopts when bound in the substrate channel, the
difference in the S-Ni distance is small; the CoB5SH thiol being only 0.5 Å farther from the
Ni than CoBSH (8.8 Å for CoBSH vs. 9.3 Å for CoB5SH) (Table 2). This is due to the
alkanoyl chain of CoBSH not being in an extended conformation from carbons 4 to 6
(carbon 1 is the carbonyl carbon). CoB6SH on the other hand adopts a conformation that
places its thiol in virtually the same position as the thiol of CoBSH (Figure 4 and Table 2).
This is consistent with CoB6SH being a substrate. However, the kcat is 1000-fold lower than
for CoBSH (3, 35) although its Km value (180 μM, Dey & Ragsdale, in preparation) is
similar to that of CoBSH (Km = 75 μM (3)). The reason for this may be that the shorter
alkanoyl chain may not enable the analogue thiol to approach the nickel close enough for
efficient catalysis, and thus explain why CoB6SH is such a poor substrate.
In the case of the longer CoBXSH analogues, the sulfur of CoB8SH is 2.6 Å closer to the Ni
ion of F430 than that of CoBSH, and 2.5 Å closer to the thiol of HSCoM (Figure 6 and Table
2). The CoB9SH molecule follows the path of CoBSH, and reaches only a little further into
the substrate channel than CoB8SH, with the CoB9SH thiol positioned 2.9 Å closer to the Ni
than the thiol of CoBSH (Figure 6 and Table 2). This is only 0.3 Å closer than the distance
observed for the CoB8SH thiol, even though they are non-coincident. The distance to the
thiol of HSCoM is 2.6 Å closer than that of the substrate, CoBSH; only 0.1 Å closer than the
CoB8SH thiol. The two analogue thiols are above an annular hydrophobic aromatic
environment created by Pheα330, Tyrα333, Pheα443, Pheβ361 and Tyrβ367 that lies
between them and F430 (Figure 6). As a result, penetrating further into the channel may be
energetically unfavorable, consistent with the small difference in relative distances between
the CoB8SH/CoB9SH thiols and the HSCoM thiol/F430 nickel. The annulus is proposed to
be catalytically important in positioning methyl-SCoM and stabilizing the methane product,
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and the tyrosines have been proposed to be proton donors associated with mechanism II
(Scheme 2B) (5, 33).
Thus, there appear to be three preferential distances for thiols (including that of HSCoM)
within the MCR substrate channel; HSCoM at 2.4 Å, CoB6SH/CoBSH at 8.7 – 8.8 Å and
CoB8SH/CoB9SH at 5.9 – 6.2 Å from the nickel of F430 (Table 2).
Recent ENDOR, high-field continuous and pulse EPR work has identified changes in nickel
co-ordination when CoBSH is added to MCRred1c (active Ni(I)-MCRred1 + HSCoM) (14,
15, 18, 31). This generates up to 50% MCRred2 which is comprised of two distinct nickel co-
ordination geometries; an axial MCRred2a formally assigned as a Ni(III)-hydride, and a
rhombic MCRred2r in which the thiol of HSCoM is a Ni(I) ligand (Supporting Information,
Scheme S1). Formation of MCRred2 could also be induced by addition of CoBSH substrate
analogues CoBS-CH3 and CoBS-CF3, which have a chain length one unit longer than
substrate CoBSH (18, 53). The CoBS-CF3 enabled 19F-ENDOR studies to be performed,
and demonstrated that following CoBS-CF3 addition the remaining MCRred1 species had
Ni(I)-19F distances of 6.2 – 7.7 Å. This distance range agreed with a CoBS-CF3 model
created using the CoBSH position observed in the MCRox1-silent crystal structure (53).
However, in the MCRred2 species the Ni(I)-19F distances had shortened indicating a
movement of ~ 2 Å towards the nickel (MCRred2a Ni(I)-19 F, 4.0 – 5.5 Å; MCRred2r
Ni(I)-19F, 4.5 – 5.7 Å). In the case of the CoB8SH analogue (the closest equivalent to CoBS-
CF3 used in this study) the thiol to Ni(II) distance lies between the distance ranges observed
in the CoBS-CF3 studies, and so the fluorine(s) of the CoBS-CF3 in MCRred2 might
penetrate a little further into the hydrophobic annulus in the MCRred2 species. As the
alkanoyl chain of CoBSH is not fully extended it could easily undergo a similar
conformation change to that observed in the MCRred2 state.
Reaction of MCR Ni(III)-alkyl species’ with CoB8SH and CoB9SH
The two longer CoBXSH analogues have been shown to undergo alkylation when reacted
with MCRPS, a [Ni(III)-alkyl ↔ Ni(II)-alkylsulfonate radical] formed from reaction of
Ni(I)-MCRred1 with bromopropanesulfonate (BPS) (Supporting Information, Scheme S1)
(20, 23, 30, 45, 54). BPS is a substrate of MCRred1 in a reaction that involves a rapid
CoBSH-independent nucleophilic attack by Ni(I) on BPS to displace bromide and generate
MCRPS at a rate ~60-fold faster than generation of methane from CoBSH and methyl-
HSCoM (20, 45). Certain thiols can eliminate the propylsulfonate to yield a thioether
product and regenerate MCRred1, although at a rate 1000-fold slower than methane
formation (45). Both CoB8SH and CoB9SH can react with MCRPS to regenerate MCRred1,
but CoBSH cannot. The overall second-order rate constant for the reactivation of MCR by
CoB8SH is 160 M−1s−1, whereas for CoB9SH the reaction is more sluggish (12 M−1s−1).
CoB9SH might be expected to be closer to the proximal Ni ligand. It was therefore proposed
that this caused steric interference and explained why CoB9SH was a poorer reactivator of
MCR than CoB8SH. Our study has shown that the thiols of these two analogues are placed
such that they are approximately the same distance (~3.7 Å) from the thiol of HSCoM
ligated to the Ni atom (Table 2). The Ni(II)-HSCoM bond is 2.4 Å, whereas an Ni(III)-alkyl
bond is expected to be ~2 Å (24, 33), thus indicating that a conformational change is
required to effect the nucleophilic attack of the CoB8SH and CoB9SH thiols on an alkyl-
bound species. It would thus appear that a conformational change, such as observed in
MCRred2, is required for this chemistry also (53).
A Ni(III)-alkyl species is akin to the first intermediate in mechanism I of MCR-catalyzed
methane formation, Ni(III)-methyl (MCRMe, Supporting Information, Scheme S1, Scheme
2A) (11, 27). MCRMe has been shown to be capable of generating MCRred1 and methyl-
SCoM upon addition of HSCoM (which is the reverse of mechanism I, step 1, Scheme 2A);
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similar chemistry to the observed formation of a thioether product from the Ni(III)-alkyl.
Further addition of CoBSH following HSCoM treatment of MCRMe led to methane and
heterodisulfide formation, the natural products of methanogenesis. Although this lends
credence to mechanism I, it should be noted that like MCRPS, MCRMe in these experiments
was generated artificially. However, the MCRCoB9SH crystal structure demonstrates that the
two additional methylene units in the alkanoyl chain c.f. CoBSH, do not necessarily translate
into direct interaction of the thiol with the nickel proximal ligand. However, this could
represent the favorable position for a CoBSH thiol interacting with the methyl group of
methyl-SCoM. Just as the alkanoyl chain of CoB6SH has a more extended conformation
than CoBSH in the substrate channel, CoBSH could also adopt a more extended
conformation so that its thiol was in a similar position as the thiol of CoB8SH, priming it for
reaction with a nickel bound species.
If a significant conformational change is required early in MCR-catalyzed chemistry, which
would be a requirement of mechanism I, catalysis may well involve a rearrangement of the
aromatic amino acid annulus due to the presence of the methyl of methyl-SCoM, and this
might enable deeper penetration of CoBSH into MCR (Figure 6). All the crystal structures in
this study, and those solved previously, are of the inactive Ni(II)-MCR, which disfavors
close approach to the nickel in the absence of Ni(I)-bound methyl-SCoM, even in the case of
CoB9SH.
Conclusion
The goal of this study was to induce structural changes within the substrate channel and
active site of MCR using analogues of coenzyme CoBSH. It was hoped that this would shed
light on the nature of conformational changes that have been proposed to occur in MCR
catalysis. We have shown that that the CoBXSH analogues do not lead to any significant
conformational changes within the context of inactive Ni(II)-MCR. Therefore, it may be that
methyl-SCoM is the key coenzyme, in combination with a nickel oxidation state of 1 (and
3), that triggers a conformational change bringing the thiol of CoBSH closer to the nickel.
Thus, the crystal structure of the Ni(I)-methyl-SCoM/MCR complex may be required to
structurally define the conformational changes required for MCR-mediated chemistry.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
X-ray data were collected at beam-line 14-BM-C and photoreduction studies at 14-ID-B, BioCARS at the
Advanced Photon Source (APS), Argonne National Laboratory, Argonne, IL. We thank Vukica Srajer and Yu-
Sheng Chen for valuable assistance during data collection. Use of the Advanced Photon Source was supported by
the U.S. Department of Energy, Basic Energy Sciences, Office of Science under Contract No. DE-
AC02-06CH11357. Use of the BioCARS Sector 14 was supported by the National Institutes of Health, National
Center for Research Resources, under grant number RR007707. Computer resources were provided by the Basic
Sciences Computing Laboratory of the University of Minnesota Supercomputing Institute, and we thank Can
Ergenekan for his support. We also thank Ed Hoeffner at the Kahlert Structural Biology Laboratory (KSBL) at the
University of Minnesota. Use of the KSBL was supported by a Minnesota Partnership for Biotechnology and a
Medical Genomics Grant SPAP-05-0013-P-FY06.
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Crystallogr. 2004; 60:2126–2132. [PubMed: 15572765]
53. Ebner S, Jaun B, Goenrich M, Thauer RK, Harmer J. Binding of coenzyme B induces a major
conformational change in the active site of methyl-coenzyme M reductase. J Am Chem Soc. 2010;
132:567–575. [PubMed: 20014831]
54. Goenrich M, Mahlert F, Duin EC, Bauer C, Jaun B, Thauer RK. Probing the reactivity of Ni in the
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Figure 1.
The active site and substrate channel in the MCRox1-silent crystal structure (PDB code 1hbn)
(9). Coenzyme F430, CoBSH and HSCoM are drawn as stick colored by atom (carbon: dark
grey). The nickel is displayed as a green sphere, water as a red sphere. Interactions are
drawn as dashed lines, and the corresponding distance is indicted in Angstroms (Å). The
path of the substrate channel was defined in the absence of F430, CoBSH, HSCoM and
water, with the surface closest to the viewer cut away. The figure was generated using
PyMOL (http://www.pymol.org).
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Figure 2.
Drawing of CoBSH analogues; (A) N-5-mercaptopentanoylthreonine phosphate (CoB5SH);
(B) N-6-mercaptohexanoylthreonine phosphate (CoB6SH); (C) N-8-
mercaptooctanoylthreonine phosphate (CoB8SH); (D) N-9-mercaptononanoylthreonine
phosphate (CoB9SH).
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Figure 3.
The active sites and substrate channels of the MCR crystal structures.; (A) MCRCoB5SH; (B)
MCRCoB6SH; (C) MCRHSCoM; (D) MCRCoB8SH; (E) MCRCoB9SH. 2Fo-Fc electron density
map around the CoBXSH analogues, waters in the CoBSH binding part of the channel and
the acetate ion (contoured at 1σ) is shown as a blue mesh. The protein is drawn as cartoon.
CoBXSH and acetate are drawn as stick and colored by atom (carbon: CoB5SH orange;
CoB6SH pale yellow; CoB8SH light blue; CoB9SH magenta; acetate white). Coenzyme F430
and HSCoM are drawn as stick colored by atom (carbon: F430 dark grey; HSCoM medium
grey). The nickel is displayed as a green sphere, and waters as red spheres. The figure was
generated using PyMOL (http://www.pymol.org/).
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Figure 4.
Overlay of CoBSH (from PDB code 1hbn) and the different CoBSH analogues. CoBXSH are
drawn as stick with the thiol represented by a sphere, and colored CoB5SH orange; CoB6SH
pale yellow; CoBSH light green; CoB8SH light blue; CoB9SH magenta. The protein is
drawn as cartoon. Coenzyme F430 and HSCoM are drawn as stick colored by atom (carbon:
F430 dark grey; HSCoM medium grey). The nickel is displayed as a green sphere. The figure
was generated using PyMOL (http://www.pymol.org/).
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Figure 5.
Hydrogen bonding diagram for the water structure modeled in MCRHSCoM. The water
molecules are named as in Figure 3C (W1–W9); WA, WB and WC are water molecules that
are present in all structures (i.e. in concert with the substrate CoBSH and the CoBSH
analogues). Interactions between surrounding residues and the water molecules are drawn as
dashed lines, and the corresponding distance is indicated in Angstroms (Å).
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Figure 6.
Stereo image of the annulus of aromatic amino acids distal of coenzyme F430. The protein is
drawn as cartoon with the side-chains of the aromatic residues drawn as white stick.
CoBSH, (from PDB code 1hbn (9)), CoB8SH and CoB9SH are drawn as stick with the thiols
represented by spheres, and colored CoBSH light green; CoB8SH light blue; CoB9SH
magenta. Coenzyme F430 and HSCoM are drawn as stick colored by atom (carbon: F430
dark grey; HSCoM medium grey). The nickel is displayed as a green sphere. The figure was
generated using PyMOL (http://www.pymol.org/).
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Scheme 1.
Reaction catalyzed by methyl-coenzyme M reductase
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Scheme 2.
Two of the proposed catalytic mechanisms for methyl-coenzyme M reductase; (A)
mechanism I; (B) mechanism II.
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Table 1
X-ray Data Collection, Processing and Refinement Statistics
Data collection and processing statistics
Name of data set
MCRCoB5SH
MCRCoB6SH
MCRHSCoM
MCRCoB8SH
MCRCoB9SH
Measured reflections
1969388
2427498
1440665
1160543
1425506
Unique reflections
553755
446253
405349
211803
401701
Resolution (Å) a
50.0–1.30 (1.35–1.30)
50.0–1.40 (1.45–1.40)
50.0–1.45 (1.50–1.45)
50.0–1.80 (1.86–1.80)
50.0–1.45 (1.50–1.45)
Completeness (%) a
97.1 (78.1)
99.9 (100.0)
99.5 (99.7)
99.8 (100.0)
98.1 (95.4)
R-sym (%) a,b
5.5 (32.9)
7.3 (44.7)
6.2 (44.0)
8.4 (47.7)
5.6 (42.5)
I/σI a
22.3 (3.6)
20.4 (4.0)
20.2 (3.2)
21.8 (3.9)
24.3 (3.2)
Space group
P21
P21
P21
P21
P21
Refinement and model building statistics
Resolution (Å) a
20.49–1.30 (1.33–1.30)
19.89–1.40 (1.44–1.40)
20.15–1.45 (1.49–1.45)
19.93–1.80 (1.84–1.80)
20.07–1.45 (1.48–1.45)
No. of reflection in working set a
525817 (30239)
423854 (25833)
384868 (25791)
201128 (11193)
381474 (23611)
No. of reflection in test set a
27777 (1576)
22348 (1331)
20362 (1319)
10625 (557)
20163 (1210)
R-work (%) c
14.32
13.04
13.47
14.95
13.58
R-free (%) d
16.56
15.53
16.22
19.54
16.44
ESU (Å) R-work/R-free
0.044/0.046
0.049/0.051
0.056/0.059
0.121/0.119
0.057/0.060
No. protein atoms
20087
19960
20265
19750
20036
No. coenzyme atoms
218
220
180
224
272
No. ligand atoms
37
62
52
26
49
No. water molecules
2443
2352
2516
1893
2432
RMS
bond lengths (Å)
0.033
0.033
0.032
0.028
0.032
bond angles (deg.)
2.693
2.625
2.468
2.059
2.549
Ramachandran plot (%)
favored
97.8
97.5
97.6
97.2
97.7
allowed
2.1
2.4
2.3
2.7
2.1
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disallowed
0.1
0.1
0.1
0.1
0.1
Average B-factor (Å2)
protein
12.42
13.35
12.12
17.22
12.73
coenzymes
8.20
9.24
7.25
11.24
8.27
ligands
31.95
35.48
28.29
33.76
32.92
waters
22.95
24.89
23.85
26.79
24.09
over all
13.54
14.57
13.40
18.02
13.93
Occupancy of HSCoM per active site (%)e
90/90
50/50
100/100
90/90
90/85
Occupancy of CoBSH per active site (%) e
50/50
50/50
30/30
50/50
40/40
CoBSH analogue, occupancy per active site (%) e
CoB5SH, 50/50
CoB6SH, 50/50
CoB8SH, 50/50
CoB9SH, 60/60
Other molecule, occupancy per active site (%) e
Acetate, 70/70
aValues in brackets correspond to the highest resolution shell.
bR-sym = ΣhklΣNj=1|Ihkl−Ihkl(j)|/ΣhklN*Ihkl, sum over all reflections and all observations N, with Ihkl(j) intensity of the jth observation of reflection hkl and Ihkl mean intensity of the reflection hkl.
cR-work = Σ||Fo| − |Fc||/Σ|Fo|, where |Fo| = observed structure factor amplitude and |Fc| = calculated structure factor amplitude.
dR-free, R-factor based on 5% of the data excluded from refinement.
eOccupancy of model in each of the two crystallographically independent active sites in the ASU
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Table 2
Distances from analogue thiols.
CoBXS - SCoM distance (Å)
CoBXS - Ni distance (Å)
CoB5SH
7.11/7.11a
9.30/9.30
CoB6SH
6.26/6.26
8.70/8.70
CoB7SH (substrate) b
6.37/6.39
8.73/8.77
CoB8SH
3.75/3.78
6.16/6.17
CoB9SH
3.71/3.68
5.96/5.91
aDistances in the two crystallographically independent active sites in the ASU
bDistances in the 1.16 Å resolution MCRox1-silent structure (PDB code 1hbn)(33)
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|
3M30
|
Structural Insight into Methyl-Coenzyme M Reductase Chemistry using Coenzyme B Analogues
|
Structural Insight into Methyl-Coenzyme M Reductase Chemistry
using Coenzyme B Analogues,†,‡
Peder E. Cedervall§,#, Mishtu Dey||,#,⊥, Arwen R. Pearson§,⊥, Stephen W. Ragsdale||, and
Carrie M. Wilmot*,||,§
§ Department of Biochemistry, Molecular Biology and Biophysics, University of Minnesota,
Minneapolis, Minnesota 55455
|| Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109
Abstract
Methyl-coenzyme M reductase (MCR) catalyzes the final and rate-limiting step in methane
biogenesis; the reduction of methyl-coenzyme M (methyl-SCoM) by coenzyme B (CoBSH) to
methane and a heterodisulfide (CoBS-SCoM). Crystallographic studies show that the active site is
deeply buried within the enzyme, and contains a highly reduced nickel-tetrapyrrole, coenzyme
F430. Methyl-SCoM must enter the active site prior to CoBSH, as species derived from analogues
of methyl-SCoM are always observed bound to the F430 nickel in the deepest part of the 30 Å long
substrate channel that leads from the protein surface to the active site. The seven-carbon
mercaptoalkanoyl chain of CoBSH binds within a 16 Å predominantly hydrophobic part of the
channel close to F430, with the CoBSH thiolate lying closest to the nickel at a distance of 8.8 Å. It
has previously been suggested that binding of CoBSH initiates catalysis by inducing a
conformational change that moves methyl-SCoM closer to the nickel promoting cleavage of the C-
S bond of methyl-SCoM. In order to better understand the structural role of CoBSH early in the
MCR mechanism, we have determined crystal structures of MCR in complex with four different
CoBSH analogues; pentanoyl-, hexanoyl-, octanoyl- and nonanoyl- derivatives of CoBSH
(CoB5SH, CoB6SH, CoB8SH and CoB9SH respectively). The data presented here reveal that the
shorter CoB5SH mercaptoalkanoyl chain overlays with that of CoBSH, but terminates two units
short of the CoBSH thiolate position. In contrast, the mercaptoalkanoyl chain of CoB6SH adopts a
different conformation, such that its thiolate is coincident with the position of the CoBSH thiolate.
†This work was supported by a Department of Energy grant DE-FG02-08ER15931 to S.W.R and supplement to C.M.W. and a
Minnesota Partnership for Biotechnology and Medical Genomics grant SPAP-05-0013-P-FY06.
‡Co-ordinates and structure factors have been deposited in the Protein Data Bank as entries 3m1v (MCRHSCoM); 3m2r
(MCRCoB5SH); 3m2u (MCRCoB6SH); 3m2v (MCRCoB8SH); 3m30 (MCRCoB9SH).
*Address correspondence to: Carrie M. Wilmot, Tel: (612) 624-2406, wilmo004@umn.edu.
⊥Current addresses: Mishtu Dey, Department of Chemistry, Massachusetts Institute of Technology, 77 Massachusetts Ave.,
Cambridge, Massachusetts 02139; Arwen Pearson, Astbury Centre for Structural Molecular Biology, Institute for Molecular and
Cellular Biology, Astbury Building. Leeds, LS2 9JT, U.K.
#These authors contributed equally to this work.
Supporting Informational material is available free of charge via the Internet at http://pubs.acs.org and contains the following:
MATERIAL AND METHODS for single crystal UV-visible microspectrophotometry, X-ray photoreduction experiment and X-ray
crystallography of the MCR-heterodisulfide product complex (MCRCoBSH + methyl-SCoM); RESULTS AND DISCUSSION for
redox changes and MCRCoBSH + methyl-SCoM crystal structure; Table S1, X-ray Data Collection, Processing and Refinement
Statistics for MCRCoBSH + methyl-SCoM; Figure S1, use of Fo-Fc electron density in modelling MCRCoB8SH; Figure S2,
illustration of electron density quality of MCRCoB5SH; Figure S3, solution and single crystal UV-visible spectra; Figure S4,
modeling of CoB6SH and CoBSH into the electron density of MCRCoB6SH; Figure S5, partially occupied HSCoM in MCRCoB6SH;
Figure S6, the active site and substrate channel of MCRCoBSH + methyl-SCoM; Figure S7, alternative conformation of Valα482 in
MCRHSCoM; Figure S8, the two conformations of the gly-rich loop in MCRHSCoM; Figure S9, propagation of conformational
changes in MCRHSCoM; Figure S10, the two conformations of CoB9SH in MCRCoB9SH; Figure S11, EPR spectra of MCRred1
sample; Scheme S1, scheme of the characterized forms of MCR.
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This is consistent with the observation that CoB6SH is a slow substrate. A labile water in the
substrate channel was found to be a sensitive indicator for the presence of CoBSH and HSCoM.
The longer CoB8SH and CoB9SH analogues can be accommodated in the active site through
exclusion of this water. These analogues react with Ni(III)-methyl; a proposed MCR catalytic
intermediate of methanogenesis. The CoB8SH thiolate is 2.6 Å closer to the nickel than that of
CoBSH, but the additional carbon of CoB9SH only decreases the nickel thiolate distance a further
0.3 Å. Although the analogues did not induce any structural changes in the substrate channel, the
thiolates appeared to preferentially bind at two distinct positions in the channel; one being the
previously observed CoBSH thiolate position, and the other being at a hydrophobic annulus of
residues that lines the channel proximal to the nickel.
INTRODUCTION
Methanogenic archaea are organisms that under strict anaerobic conditions derive energy by
reducing compounds such as carbon dioxide, methylamine, acetate and methanol, to
methane (1, 2). The global production of methane by these organisms is estimated at one
billion tons annually. Microbially produced methane is not only a potential source of
renewable energy but also a potent greenhouse gas, and as such study of this process has
environmental ramifications. In these microorganisms, methyl-coenzyme M reductase
(MCR)1 is the enzyme that catalyzes the final step in methanogenesis, in which the
substrates methyl-coenzyme M (methyl-SCoM, 2-(methylthio)ethanesulfonate) and
coenzyme B (CoBSH, N-7-mercaptoheptanoylthreonine phosphate) are converted to
methane and a heterodisulfide (CoBS-SCoM) (Scheme 1) (3).
MCR is a 300 kDa protein with six subunits arranged in a α2β2γ2 oligomer (4). The known
crystal structures show that MCR has two active sites approximately 50 Å apart that are
deeply buried within the enzyme (5). The active site pocket is comprised of residues from
subunits α, α′, β and γ, with a 30 Å long substrate channel leading to the enzyme surface
(Figure 1). At the heart of the active site pocket is coenzyme F430, which is a highly reduced
nickel-containing tetrapyrrole (6–8). Currently sixteen distinct enzymatic and complexed
states of MCR have been spectroscopically characterized (Supporting Information, Scheme
S1) (9–31). In the resting active state of the enzyme, denoted MCRred1, the redox-active
nickel of F430 is present in the Ni(I) state (9, 16, 32). MCR is extremely oxygen sensitive
and upon oxygen exposure the enzyme enters an inactive Ni(II) state, denoted MCRred1-silent
(6). In this state it cannot be converted back to the active Ni(I) form by any known reducing
agent making this a challenging system to study. Additional complications involve the tight
association of coenzymes to purified MCR that are not easily displaced as demonstrated by
X-ray crystallographic and kinetic studies (5, 33–35).
Despite the fact that MCR has been studied for decades, no true catalytic intermediate has
been observed, and the actual mechanism remains elusive. Currently three general
mechanistic schemes for the enzymatic reaction have been proposed, each of which posit
different chemistry to initiate catalysis. Mechanism I involves Ni(I) acting as a nucleophile
in an SN2-type reaction that generates Ni(III)-methyl as an intermediate (Scheme 2A) (35–
38). Mechanism II starts with methyl-SCoM undergoing homolytic cleavage at the Ni(I) to
generate a methyl radical and a Ni(II)-SCoM species (Scheme 2B) (39–41). A more recently
proposed mechanism III suggests protonation of coenzyme F430 promotes reductive
cleavage of the methyl-SCoM thioether bond (42).
1Abbreviations: MCR, Methyl-coenzyme M reductase; methyl-SCoM, methyl-coenzyme M; CoBSH, coenzyme B; HSCoM,
coenzyme M; CoBS-SCoM, heterodisulfide of coenzyme B and coenzyme M; APS, Advanced Photon Source; ASU, asymmetric unit;
BPS, bromopropanesulfonate.
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Due to the stringent requirement to exclude O2, the available MCR crystal structures are all
in the inactive Ni(II) state. All but one contain CoBSH and HSCoM (demethylated methyl-
SCoM, an inhibitor and substrate analogue) in the active site (PDB codes 1hbn, 1hbo, 1hbu,
1e6y, 1e6v) (5, 33, 34). Another crystal structure has bound heterodisulfide product, CoBS-
SCoM (MCRsilent, PDB code 1hbm, Scheme 1 and Supporting Information, Scheme S1) (5,
33). All these structures reveal that both substrates access the active site through the same
channel (Figure 1). The binding site of HSCoM (and presumably methyl-SCoM) is more
deeply buried within the enzyme, and so it must enter prior to CoBSH for productive
chemistry to occur. As binding of CoBSH in the absence of co-substrate would be
inhibitory, it was suggested that a conformational change upon methyl-SCoM binding might
lower the Kd for CoBSH, and thus promote an ordered mechanism. Furthermore it has been
suggested that CoBSH binding induces a conformational change that brings the methyl-
SCoM substrate into closer proximity to the nickel, and this promotes C-S bond cleavage.
To investigate the proposed structural role of CoBSH in initiating catalysis, we have solved
the X-ray crystal structures of MCR in complex with four different CoBSH analogues.
CoBSH has a heptanoyl moiety linked to the thiol group, and the analogues are pentanoyl-,
hexanoyl-, octanoyl- or nonanoyl-containing derivatives of CoBSH (CoB5SH, CoB6SH,
CoB8SH and CoB9SH respectively; Figure 2) (3, 35, 43–47). In addition, we present a
structure in which the substrate channel predominantly lacks either CoBSH or
heterodisulfide product.
MATERIALS AND METHODS
Materials
The organism Methanothemobacter marburgensis (catalog OCM82) was obtained from the
Oregon Collection of Methanogens (Portland, OR). All buffers and media reagents were
obtained from Sigma-Aldrich (St. Louis, MO). The gases N2 (99.98%), H2/CO2 (80%/20%),
and ultra high purity H2 (99.999%) were obtained from Cryogenic Gases (Grand Rapids,
MI). Ti(III) citrate solutions were prepared from a stock solution of 246 mM Ti(III) citrate,
which was synthesized by adding sodium citrate to Ti(III) trichloride (30 wt % solution in 2
N hydrochloric acid) (Acros Organics, Morris Plains, NJ) under anaerobic conditions and
adjusting pH to 7.0 with sodium bicarbonate (48). The concentration of Ti(III) citrate was
determined by titrating against a solution of methyl viologen.
Synthesis of methyl-SCoM, CoB5SH, CoB6SH, CoBSH, CoB8SH, and CoB9SH
Methyl-SCoM was prepared from HSCoM and methyl iodide (49). The homodisulfides,
CoB5S-SCoB5, CoB6S-SCoB6, CoBS-SCoB, CoB8S-SCoB8, CoB9S-SCoB9 were prepared
as described from 5-bromovaleric acid, 6-bromohexanoic acid (Sigma-Aldrich, St. Louis,
MO), 7-bromoheptanoic acid (Karl Industries, Aurora, OH), 8-bromooctanoic acid, and 9-
bromononanoic acid (Matrix Scientific, Columbia, SC), respectively (43, 46). The free thiol
forms of CoB5SH, CoB6SH, CoBSH, CoB8SH, and CoB9SH were generated by the
reduction of the homodisulfides as previously described (45). The purity of the CoBSH
analogues was determined by 1H NMR spectroscopy. All compounds synthesized were
stored in a Vacuum Atmospheres chamber maintained at an oxygen level below 1 ppm, as
monitored continually with an oxygen analyzer (model 317, Teledyne Analytical
Instruments, City of Industry, CA) until use.
M. marburgensis Growth and MCRred1 Purification
Buffer preparations and all manipulations were performed under strict anaerobic conditions
in a Vacuum Atmospheres chamber maintained at an oxygen level below 1 ppm, as
monitored continually with an oxygen analyzer (model 317, Teledyne Analytical
Instruments, City of Industry, CA). MCRred1 was isolated from M. marburgensis cultured on
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H2/CO2 (80%/20%) at 65 °C in a 14-L fermentor (New Brunswick Scientific Co., Inc. New
Brunswick, NJ). Culture media were prepared as previously described (20, 24). MCRred1
was generated in vivo and purified as described previously (20). The purification procedure
routinely generates 65–75% MCRred1 as determined by UV-visible and EPR spectroscopy.
Spectroscopy of MCR
UV-visible spectra of the Ni(I)-containing MCRred1 were recorded in the anaerobic chamber
using a spectrophotometer (Model USB4000-UV-VIS, Ocean Optics, Dunedin, FL). EPR
spectra were recorded on a Bruker EMX spectrometer (Bruker Biospin Corp., Billerica,
MA), equipped with an Oxford ITC4 temperature controller, a Hewlett-Packard Model 5340
automatic frequency counter and Bruker gaussmeter. The EPR spectroscopic parameters
included: temperature, 70 K; microwave power, 10 mW; microwave frequency, 9.43 GHz;
receiver gain, 2 × 104; modulation amplitude, 10.0 G; modulation frequency, 100 kHz.
Double integrations of the EPR spectra were performed and referenced to a 1 mM copper
perchlorate standard. The NMR data were acquired at 298 K on a Bruker Avance DRX 500
MHz instrument equipped with a TXI cryoprobe.
Preparation of MCRred1 for Crystallization
All crystallization experiments were performed in the anaerobic chamber in which MCR
was purified unless otherwise noted. MCRred1 was prepared in 50 mM Tris, pH 7.6 and
excess HSCoM was removed by buffer exchange using an Amicon Ultra centrifuge filter
with a 50 kDa cut-off membrane (Millipore). Typically, 2–3 ml of MCRred1 was exchanged
with 10–15 ml of 50 mM Tris, pH 7.6. The enzyme was concentrated to 500–600 μl, and
this process was repeated three times. The fraction of MCRred1 in the purified MCR sample
was calculated from the UV-visible spectrum using extinction coefficients of 27.0
mM−1cm−1 at 385 nm for Ni(I)-MCRred1, and 9.15 mM−1cm−1 at 420 nm for Ni(II)-
MCRred1-silent (20). The amount of MCRred1 in samples used for crystallization was
determined to be ~80% and the concentration of total enzyme used was in the range of about
120–150 μM (~32–40 mg/ml). All crystallization experiments were performed anaerobically
by incubating 2.0 μl of enzyme solution in 50 mM Tris, pH 7.5 and 2.0 μl of reservoir
solution (100 mM Hepes-Na, pH 7.3/7.5/8.0; 150 mM magnesium acetate (Mg(CH3COO)2),
and 20/22% (w/v) PEG 400) in a sitting drop over 1 ml reservoir solution at 9 ºC. Triangular
and rectangular prismatic crystals with a bright yellowish-green color confirmed the
presence of nickel coenzyme F430. The crystals grew to a size of approximately 100–200 μm
in 4–5 days. CoBSH-depleted crystals were obtained by incubating 2 μl of a reaction
mixture containing 139 μM MCRred1 and 13 mM HSCoM with 2 μl of reservoir solution
(100 mM Hepes-Na pH 7.5, 150 mM magnesium acetate (Mg(CH3COO)2), 22% PEG 400).
Crystals of MCR complexed with the CoBSH analogues were grown by co-crystallization.
The CoB5SH co-crystals were obtained by incubating 2μl enzyme solution containing 124
μM MCRred1, 10 mM methyl-SCoM and 1 mM CoB5SH with 2μl of reservoir solution (100
mM Hepes-Na, pH 7.5, 150 mM (Mg(CH3COO)2), 22 % PEG 400). The crystals with
bound CoB6SH and CoB9SH were obtained by co-crystallization of 1 mM of analogue with
142 μM MCRred1 and equilibrated with 2μl of reservoir solution (100 mM Hepes-Na, pH
7.5, 150 mM Mg(CH3COO)2, 20 % PEG 400 for CoB6SH and 100 mM Hepes-Na, pH 7.3,
150 mM Mg(CH3COO)2), 22 % PEG 400 for CoB9SH). Crystals were cryoprotected in
reservoir solution containing 25 % (v/v) PEG 400 by soaking for 2–5 minutes before
cryocooling in liquid nitrogen in the anaerobic chamber. Crystals of CoB8SH bound to MCR
were obtained by incubating 2 μl of a mixture of 119 μM MCRred1 and 1 mM CoB8SH with
2 μl of reservoir solution (100 mM Hepes-Na, pH 7.3, 150 mM Mg(CH3COO)2), 20 % PEG
400). Before cryoprotection, the crystals were soaked for 5–10 minutes in a 100 mM
solution of methyl iodide2. The methyl iodide solution used for soaking was prepared by
adding a concentrated stock of methanolic solution of methyl iodide to the reservoir
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solution. Soaked crystals were quickly cryoprotected as described above and cryocooled in
the anaerobic chamber.
X-ray Diffraction Data Collection, Processing and Refinement
X-ray diffraction data were collected at 100 K on a ADSC Quantum-315 detector at the APS
Beamline 14-BM-C (BioCARS). The wavelength of X-rays was 0.979 Å. Data were
processed using HKL2000 (50). As in the previous X-ray crystallographic studies, the
crystals belong to the monoclinic space group P21 (a = 82 Å, b = 118 Å, c = 122 Å, β = 92°),
with one MCR molecule (two active sites) per asymmetric unit (5, 33). For refinement,
REFMAC in the Collaborative Computational Project Number 4 (CCP4) program suite was
used (51). A random sample of 5 % of the data across all resolution shells was chosen to
check refinement progress through calculation of an Rfree. The same reflections were used to
calculate Rfree for all structures, thus preventing bias due to high structural identity. The
remaining reflections were used in refinement (Rwork). Model building was done using the
Crystallographic Object-Oriented Toolkit (COOT) (52). The diffraction data and their
models are designated as MCRCoBXSH, where X is the number of carbons in the alkanoyl
portion of the analogue. Library files in CCP4 for F430 and CoBSH were incorrect, and these
were modified in Monomer Library Sketcher in the CCP4 program suite by comparison with
schematic drawings from Grabarse et al. (33). Co-ordinate and CCP4 library files for the
different CoBSH analogues were created in Monomer Library Sketcher. The general model
building and refinement strategy for all structures was as follows. It was clear from the
electron density in the substrate channel and at the active site that mixtures of species were
present in all datasets. These could be visualized with Fo-Fc and Fo-Fo difference electron
density maps (Supporting Information, Figure S1). The known positions of CoBSH and
HSCoM from the published Ni(II)-MCR crystal structures (PDB codes 1hbn, 1hbo, 1hbu
(33)) were used as guides to determine which species could be present in each dataset, and
these were then simultaneously modeled into the electron density. By alteration of their
relative occupancies (in 10% increments) followed by refinement, the ratio of occupancy
between different species was determined using the assumption that the average B-factors
for all molecular species bound should be similar to that of F430 and adjacent well-ordered
protein atoms within the active site and substrate channel. The combinations of modeled
ligands were constantly reassessed throughout refinement based on the remaining difference
electron density. This included test refinements of different ligand combinations during the
latter stages, thus using the optimized phases to check whether a different combination of
ligands could also explain the electron density. Sensible chemical structures and
interactions, along with keeping the combined occupancies of sterically mutually exclusive
species ≤ 100%, were maintained throughout refinement. The model was finally accepted
when the difference electron density map was minimal and the B-factors for the models
converged.
In practice the first structure refined was that of MCRCoB5SH. Initial phases were generated
by difference Fourier using a previously determined crystal structure (PDB code 1mro (5))
but with all non-bonded molecules, including water, removed from the model except F430.
Initial rigid body refinement followed by restrained refinement of MCRCoB5SH reduced the
Rwork to 26.5%. After model building and subsequent rounds of restrained refinement the
Rwork was 14.3% (Rfree 16.6%). Of the five structures only the CoB5SH analogue is
completely coincident with CoBSH, and so particular care had to be used in teasing apart the
ratios of the two species in modeling the MCRCoB5SH electron density. This was done by
2Methyl iodide was added with the intention of creating a Ni(III)-methyl species, which CoB8SH stabilizes. This was not achieved,
but the diffraction quality of this crystal was significantly better than crystals co-crystallized with CoB8SH alone, and so has been
included in this study.
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initially refining 100% CoBSH in the substrate channel. Positive Fo-Fc difference density
located at the carbon where the shorter CoB5SH thiol might be expected to be indicated the
presence of a more electron-rich species than carbon, which is consistent with the presence
of the CoB5SH sulfur. The refinement converged at a model containing a 50:50 mix of
CoBSH and the analogue. However, positive Fo-Fc difference density was still present at the
position of the CoBSH thiol, therefore a water molecule was added to the CoB5SH model at
50% occupancy and upon refinement this accounted for the electron density. An illustration
of the electron density quality from this structure is shown in Supporting Information,
Figure S2. An HSCoM-, CoBSH- and CoBSH analogue-free version of the refined
MCRCoB5SH structure was used as the starting model to generate initial phases for the four
other structures. After the initial round of restrained refinement the Rwork for these structures
were reduced to 14.5–15.6 %.
RESULTS AND DISCUSSION
Crystal Structures of MCR
Five crystal structures were determined, four of which are in complex with CoBSH
analogues differing in the number of carbons atoms in the alkanoyl portion of the molecule.
CoBSH is an N-7-mercaptoheptanoyl-containing molecule, whereas the four CoBSH
analogues contain N-5-mercaptopentanoyl-, N-6-mercaptohexanoyl-, N-8-mercaptooctanoyl-
or N-9-mercaptononanoyl-moieties (Figure 2). The corresponding crystal structures are
designated as MCRCoBXSH, where X is the number of carbons in the alkanoyl portion of the
analogue. The other crystal structure is of MCRred1c-silent (MCR in the Ni(II) state in
complex with HSCoM, designated here as MCRHSCoM) that is CoBSH-depleted. The
datasets have resolutions in the range from 1.35 – 1.8 Å. Although the crystallizations were
set up with the MCR solution predominantly in the Ni(I)-MCRred1 state, by the time X-ray
diffraction data were collected they had been oxidized to the Ni(II)-MCRred1-silent state
(Supporting Information). Following data collection there was no evidence for
photoreduction of the Ni(II) back to Ni(I) in any of the crystals, as assessed by single crystal
UV-visible microspectrophotometry (Supporting Information and Figure S3). Attempts to
photoreduce the crystals using different wavelengths and temperatures were unsuccessful
(Supporting Information).
Overall, the resulting structures are very similar to each other and to the previously
published structures of MCR, with differences mainly localized to the active site and
substrate channel. The two active sites in the ASU were refined independently. Unless
otherwise stated there was no difference between them. All five datasets contain a mixture
of species bound to the enzyme. There is always a background of CoBSH and HSCoM,
which co-purify with MCR and cannot be fully removed by extensive buffer exchange or by
the addition of a CoBSH analogue. HSCoM is added during purification of MCR, as it
stabilizes the resting active Ni(I) state (unpublished data), and this leads to HSCoM
occupancies between 50–100% amongst the structures (Table 1). In contrast CoBSH, which
is not added during purification, has occupancies ranging from 30–50%. As these
confounding species have all been described at high occupancy in other crystallographic
studies, the structural data of interest could be isolated (5, 33). In each case, the additional
electron density could be explained by inclusion of the appropriate CoBXSH model used in
that experiment at 50% or higher occupancy. The resulting models, along with 2Fo-Fc
electron density, are shown in Figure 3. The Rwork for the final structures range from 13.0 to
15.0 % (Rfree 15.5 to 19.5 %). The X-ray data collection, processing, refinement and model
building statistics are given in Table 1.
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Analogues shorter than CoBSH; CoB5SH and CoB6SH
CoB5SH is two methylene groups shorter than CoBSH, the MCR substrate. The
MCRCoB5SH structure is to 1.35 Å resolution. As expected the pentanoyl chain follows the
path of the CoBSH heptanoyl carbons down the substrate channel, and thus its thiol is
positioned in the same place as the second carbon preceding the CoBSH thiol (Figure 3A
and 4). There are no published MCR kinetic studies using CoB5SH, but as it binds in the
substrate channel, it is likely to be an inhibitor.
CoB6SH is one methylene shorter than CoBSH, and is a slow substrate of MCR. In this case
the 1.4 Å resolution electron density of MCRCoB6SH indicates that the analogue
unexpectedly binds in the substrate channel such that its thiol is virtually in the same
position as the thiol of CoBSH (Figure 3B and 4). The hexanoyl chain is oriented so that it
takes a shorter route down the substrate channel between carbons 2 and 5 (with the carbonyl
carbon labeled as carbon 1) than CoBSH (Figure 4, Supporting Information, Figure S4).
This short-cut is not seen in any of the other CoBXSH complex crystal structures, but
presumably arises because this CoB6SH binding conformer is energetically more favorable,
although it is not clear from the structure why this might be the case. CoB6SH binds very
tightly to MCR, with an apparent Ki value of 0.1 μM (3).
Water structure in the absence of HSCoM
The electron density for the MCRCoB6SH crystal structure only supported the modeling of 50
% bound HSCoM. In the fraction of MCR molecules where HSCoM is absent, the HSCoM
binding site is occupied by a network of four water molecules (Supporting Information,
Figure S5). Two waters are positioned close to the absent sulfonate oxygen positions of
HSCoM. Based on the presence of positive difference electron density, a third water was
modeled ligated to the Ni, and refined to a distance of ~2 Å (2.0 Å and 2.1 Å in the two
active sites of the ASU) with no distance restraint imposed between the Ni and water. This
water is in a similar position as the Ni coordinating sulfonate oxygen of the heterodisulfide
product in MCRsilent (Supporting Information, Figure S6 and PDB codes 3m32, 1hbm) (5,
33). The fourth water was in the vicinity of the expected position of a bridging water (W1)
seen in other structures (Figure 1, 3A and 3C).
Water structure in the absence of CoBSH
The 1.45 Å resolution electron density obtained for MCRHSCoM indicates that the substrate
channel contains only 30 % CoBSH. Nine ordered waters (W1–W9), along with an acetate
ion from the crystallization solution occupy the channel, with the acetate positioned where
the phosphothreonine linkage of CoBSH would be (Figure 3C). Presumably 1–2 further
waters would replace the acetate under physiological conditions. Other than W3 and W7, the
waters form hydrogen bonds with protein (Figure 5). One water (W2) occupies the same site
as the CoBSH thiol. Presumably due to the loss of favorable interactions that exist when
CoBSH is present, the hydrophobic side-chain of Valα482 adopts a second conformation
modeled at 60 % occupancy (Supporting Information, Figure S7).
Position of the “bridging” water, W1
The equivalent of W1 has previously been observed in MCRred1-silent and MCRox1-silent
crystal structures where, in the presence of CoBSH and HSCoM, it is sited equidistant (3.2
Å) between the two coenzyme thiols (PDB codes: 1hbn, 1hbo, 1hbu) and thus been termed
the “bridging water” (Figure 1) (5, 33). However, in the MCRHSCoM structure, due to the
presence of the more polarized W2 water, W1 is displaced away from HSCoM to maximize
the hydrogen bond interaction with W2 (2.9 Å to W2; 3.5 Å to HSCoM thiol, Figure 5). In
the MCRCoB5SH structure that also contained W2, the electron density indicated that this
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repositioning of W1 towards W2 also occurred. In contrast, the MCRCoB6SH structure
contained 100% thiol at the CoBSH position, but a partial occupancy of HSCoM (50%). In
this case the electron density for W1 indicated it had moved towards the nickel to form an
optimal hydrogen bond with a Ni-ligating water that was only present in the absence of
HSCoM (3.7 Å to CoB(6)SH thiol; 3.0 Å to Ni-ligating water, Supporting Information,
Figure S5). In all structures reported here, W1 (if present) appears to be a sensitive indicator
of the relative electronegativity of the Ni-ligated atom to that occupying the position of the
CoBSH thiol, and was a useful check in the crystallographic modeling and refinement
process.
Flexibility in the substrate channel: Alternative protein conformers
The binding site of HSCoM (and presumably methyl-SCoM) is more deeply buried within
the enzyme, and so it must enter prior to CoBSH for productive chemistry to occur. As
binding of CoBSH in the absence of co-substrate would be inhibitory, it was suggested that
a conformational change upon methyl-SCoM binding might lower the Kd for CoBSH, and
thus promote an ordered mechanism. Compared to the 1hbn MCRox1-silent and 1hbu
MCRred1-silent crystal structures, which both have full occupancy HSCoM, the lower
occupancy of HSCoM in the 1hbo MCRred1-silent structure was associated with significantly
greater flexibility within the channel, and the ability to model a second conformation of a
Gly-rich amino acid stretch that formed part of the CoBSH channel. This suggested that
methyl-SCoM binding might cause the channel to become more ordered, increasing the
affinity of MCR for CoBSH by conformational restriction rather than a switch mechanism
where the structure reorganizes from one well-defined conformer to another (33). In the
MCRHSCoM data containing 30 % CoBSH and 100% HSCoM, the Fo-Fc difference electron
density map at one of the two independent active sites in the ASU contained positive peaks
that suggested the presence of an alternate conformation also involving this part of the
polypeptide (Supporting Information, Figure S8). Using this as a guide, a similar second
conformation involving seven contiguous amino acid residues of the same Gly-rich amino
acid stretch (β366–372) could be modeled and refined at 20 % occupancy leaving no
residual difference density. Parts of the α′ subunit (α′111–129 and α′237–242) that are in
close proximity to this stretch of amino acids also exhibit second conformations, with the
main-chain carbonyl of α′243 in van der Waals contact with the B ring of F430 tetrapyrrole
(Supporting Information, Figure S9). Modeling these at 20% occupancy accounted for the
weak positive Fo-Fc difference electron density peaks observed in these areas. The evidence
of alternate conformers in these areas lends support to the proposal that increased flexibility
in the substrate channel propagates through the protein (33).
The MCRCoB6SH crystal structure contains 50 % CoB6SH, 50% CoBSH and 50% HSCoM.
In this case there is no evidence of an alternate loop conformation in either active site of the
ASU. However, as CoBSH and CoB6SH combined are at 100% occupancy, it is not
surprising their favorable interactions with the substrate channel would reduce
conformational disorder, despite the partial occupancy of HSCoM.
Analogues longer than CoBSH; CoB8SH and CoB9SH
Both analogues could be accommodated in the MCR substrate channel (Figures 3D and 3E).
The electron density supported final models containing 50% CoB8SH for MCRCoB8SH (1.8
Å resolution) and 60 % CoB9SH for MCRCoB9SH (1.45 Å resolution). The phosphate head-
groups are in identical positions to those of CoBSH, CoB5SH, CoB6SH (Figure 4) (5, 33).
Both analogues follow the crystallographically observed chain path of bound CoBSH, with
the extra atoms displacing the W1 water and placing the thiols closer to the nickel (Figure
6). CoB9SH does have a second conformer that deviates from the CoBSH path, but the thiol
position for this conformer and the CoBSH-tracking conformer are identical (Figure 3E and
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Supporting Information, Figure S10). Interestingly, the thiol of CoB8SH is not coincident
with the CoB9SH carbon that precedes the CoB9SH thiol (Figure 6). CoB8SH is an MCR
inhibitor with an apparent Ki of 15 μM (3). CoB9SH has never been tested for inhibition of
MCR-catalyzed methane formation, but it is reasonable to assume that it would be an
inhibitor.
CoBXSH thiol-to-nickel spatial relationship
The position of CoBSH in previous crystal structures poses a conundrum (5, 33). In all the
proposed catalytic mechanisms, CoBSH must interact with species generated at the nickel.
Perplexingly, the sulfur of the CoBSH substrate is 8.8 Å from the Ni(II) in the MCRox1-silent
and both MCRred1-silent crystal structures, and 6.4 Å from the thiol of the substrate analogue
HSCoM (Figure 1). Modeling studies demonstrated that the addition of a methyl group to
HSCoM did not bridge this gap (35, 45, 53). Therefore, a conformational change has been
postulated that would enable CoBSH to penetrate deeper into the substrate channel, and thus
approach closer to any nickelbound species. The heterodisulfide product in the MCRsilent
crystal structure has the CoBSH portion in virtually the same place as in MCRox1-silent,
giving no clue to possible structural changes that might occur to facilitate CoBSH reacting
with nickel-associated intermediates (5, 33).
Trigonometry suggests that if the alkanoyl chain of CoBSH or its analogue is in an extended
conformation, each additional unit in the chain would lead to the thiol moving ~1.2 Å
towards the Ni. Until this study there have been no crystal structures of CoBSH analogues in
complex with MCR, so mechanistic studies using different chain length analogues of
CoBSH assumed that shorter analogues would trace the observed path of CoBSH, and
longer analogues would penetrate about ~1.2 Å deeper per additional chain unit into the
channel. In the case of the shortest analogue CoB5SH, it does indeed follow the path of
CoBSH, with the thiol of CoB5SH being 2.8 Å away from the thiol position of CoBSH.
However, due to the conformation CoBSH adopts when bound in the substrate channel, the
difference in the S-Ni distance is small; the CoB5SH thiol being only 0.5 Å farther from the
Ni than CoBSH (8.8 Å for CoBSH vs. 9.3 Å for CoB5SH) (Table 2). This is due to the
alkanoyl chain of CoBSH not being in an extended conformation from carbons 4 to 6
(carbon 1 is the carbonyl carbon). CoB6SH on the other hand adopts a conformation that
places its thiol in virtually the same position as the thiol of CoBSH (Figure 4 and Table 2).
This is consistent with CoB6SH being a substrate. However, the kcat is 1000-fold lower than
for CoBSH (3, 35) although its Km value (180 μM, Dey & Ragsdale, in preparation) is
similar to that of CoBSH (Km = 75 μM (3)). The reason for this may be that the shorter
alkanoyl chain may not enable the analogue thiol to approach the nickel close enough for
efficient catalysis, and thus explain why CoB6SH is such a poor substrate.
In the case of the longer CoBXSH analogues, the sulfur of CoB8SH is 2.6 Å closer to the Ni
ion of F430 than that of CoBSH, and 2.5 Å closer to the thiol of HSCoM (Figure 6 and Table
2). The CoB9SH molecule follows the path of CoBSH, and reaches only a little further into
the substrate channel than CoB8SH, with the CoB9SH thiol positioned 2.9 Å closer to the Ni
than the thiol of CoBSH (Figure 6 and Table 2). This is only 0.3 Å closer than the distance
observed for the CoB8SH thiol, even though they are non-coincident. The distance to the
thiol of HSCoM is 2.6 Å closer than that of the substrate, CoBSH; only 0.1 Å closer than the
CoB8SH thiol. The two analogue thiols are above an annular hydrophobic aromatic
environment created by Pheα330, Tyrα333, Pheα443, Pheβ361 and Tyrβ367 that lies
between them and F430 (Figure 6). As a result, penetrating further into the channel may be
energetically unfavorable, consistent with the small difference in relative distances between
the CoB8SH/CoB9SH thiols and the HSCoM thiol/F430 nickel. The annulus is proposed to
be catalytically important in positioning methyl-SCoM and stabilizing the methane product,
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and the tyrosines have been proposed to be proton donors associated with mechanism II
(Scheme 2B) (5, 33).
Thus, there appear to be three preferential distances for thiols (including that of HSCoM)
within the MCR substrate channel; HSCoM at 2.4 Å, CoB6SH/CoBSH at 8.7 – 8.8 Å and
CoB8SH/CoB9SH at 5.9 – 6.2 Å from the nickel of F430 (Table 2).
Recent ENDOR, high-field continuous and pulse EPR work has identified changes in nickel
co-ordination when CoBSH is added to MCRred1c (active Ni(I)-MCRred1 + HSCoM) (14,
15, 18, 31). This generates up to 50% MCRred2 which is comprised of two distinct nickel co-
ordination geometries; an axial MCRred2a formally assigned as a Ni(III)-hydride, and a
rhombic MCRred2r in which the thiol of HSCoM is a Ni(I) ligand (Supporting Information,
Scheme S1). Formation of MCRred2 could also be induced by addition of CoBSH substrate
analogues CoBS-CH3 and CoBS-CF3, which have a chain length one unit longer than
substrate CoBSH (18, 53). The CoBS-CF3 enabled 19F-ENDOR studies to be performed,
and demonstrated that following CoBS-CF3 addition the remaining MCRred1 species had
Ni(I)-19F distances of 6.2 – 7.7 Å. This distance range agreed with a CoBS-CF3 model
created using the CoBSH position observed in the MCRox1-silent crystal structure (53).
However, in the MCRred2 species the Ni(I)-19F distances had shortened indicating a
movement of ~ 2 Å towards the nickel (MCRred2a Ni(I)-19 F, 4.0 – 5.5 Å; MCRred2r
Ni(I)-19F, 4.5 – 5.7 Å). In the case of the CoB8SH analogue (the closest equivalent to CoBS-
CF3 used in this study) the thiol to Ni(II) distance lies between the distance ranges observed
in the CoBS-CF3 studies, and so the fluorine(s) of the CoBS-CF3 in MCRred2 might
penetrate a little further into the hydrophobic annulus in the MCRred2 species. As the
alkanoyl chain of CoBSH is not fully extended it could easily undergo a similar
conformation change to that observed in the MCRred2 state.
Reaction of MCR Ni(III)-alkyl species’ with CoB8SH and CoB9SH
The two longer CoBXSH analogues have been shown to undergo alkylation when reacted
with MCRPS, a [Ni(III)-alkyl ↔ Ni(II)-alkylsulfonate radical] formed from reaction of
Ni(I)-MCRred1 with bromopropanesulfonate (BPS) (Supporting Information, Scheme S1)
(20, 23, 30, 45, 54). BPS is a substrate of MCRred1 in a reaction that involves a rapid
CoBSH-independent nucleophilic attack by Ni(I) on BPS to displace bromide and generate
MCRPS at a rate ~60-fold faster than generation of methane from CoBSH and methyl-
HSCoM (20, 45). Certain thiols can eliminate the propylsulfonate to yield a thioether
product and regenerate MCRred1, although at a rate 1000-fold slower than methane
formation (45). Both CoB8SH and CoB9SH can react with MCRPS to regenerate MCRred1,
but CoBSH cannot. The overall second-order rate constant for the reactivation of MCR by
CoB8SH is 160 M−1s−1, whereas for CoB9SH the reaction is more sluggish (12 M−1s−1).
CoB9SH might be expected to be closer to the proximal Ni ligand. It was therefore proposed
that this caused steric interference and explained why CoB9SH was a poorer reactivator of
MCR than CoB8SH. Our study has shown that the thiols of these two analogues are placed
such that they are approximately the same distance (~3.7 Å) from the thiol of HSCoM
ligated to the Ni atom (Table 2). The Ni(II)-HSCoM bond is 2.4 Å, whereas an Ni(III)-alkyl
bond is expected to be ~2 Å (24, 33), thus indicating that a conformational change is
required to effect the nucleophilic attack of the CoB8SH and CoB9SH thiols on an alkyl-
bound species. It would thus appear that a conformational change, such as observed in
MCRred2, is required for this chemistry also (53).
A Ni(III)-alkyl species is akin to the first intermediate in mechanism I of MCR-catalyzed
methane formation, Ni(III)-methyl (MCRMe, Supporting Information, Scheme S1, Scheme
2A) (11, 27). MCRMe has been shown to be capable of generating MCRred1 and methyl-
SCoM upon addition of HSCoM (which is the reverse of mechanism I, step 1, Scheme 2A);
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similar chemistry to the observed formation of a thioether product from the Ni(III)-alkyl.
Further addition of CoBSH following HSCoM treatment of MCRMe led to methane and
heterodisulfide formation, the natural products of methanogenesis. Although this lends
credence to mechanism I, it should be noted that like MCRPS, MCRMe in these experiments
was generated artificially. However, the MCRCoB9SH crystal structure demonstrates that the
two additional methylene units in the alkanoyl chain c.f. CoBSH, do not necessarily translate
into direct interaction of the thiol with the nickel proximal ligand. However, this could
represent the favorable position for a CoBSH thiol interacting with the methyl group of
methyl-SCoM. Just as the alkanoyl chain of CoB6SH has a more extended conformation
than CoBSH in the substrate channel, CoBSH could also adopt a more extended
conformation so that its thiol was in a similar position as the thiol of CoB8SH, priming it for
reaction with a nickel bound species.
If a significant conformational change is required early in MCR-catalyzed chemistry, which
would be a requirement of mechanism I, catalysis may well involve a rearrangement of the
aromatic amino acid annulus due to the presence of the methyl of methyl-SCoM, and this
might enable deeper penetration of CoBSH into MCR (Figure 6). All the crystal structures in
this study, and those solved previously, are of the inactive Ni(II)-MCR, which disfavors
close approach to the nickel in the absence of Ni(I)-bound methyl-SCoM, even in the case of
CoB9SH.
Conclusion
The goal of this study was to induce structural changes within the substrate channel and
active site of MCR using analogues of coenzyme CoBSH. It was hoped that this would shed
light on the nature of conformational changes that have been proposed to occur in MCR
catalysis. We have shown that that the CoBXSH analogues do not lead to any significant
conformational changes within the context of inactive Ni(II)-MCR. Therefore, it may be that
methyl-SCoM is the key coenzyme, in combination with a nickel oxidation state of 1 (and
3), that triggers a conformational change bringing the thiol of CoBSH closer to the nickel.
Thus, the crystal structure of the Ni(I)-methyl-SCoM/MCR complex may be required to
structurally define the conformational changes required for MCR-mediated chemistry.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
X-ray data were collected at beam-line 14-BM-C and photoreduction studies at 14-ID-B, BioCARS at the
Advanced Photon Source (APS), Argonne National Laboratory, Argonne, IL. We thank Vukica Srajer and Yu-
Sheng Chen for valuable assistance during data collection. Use of the Advanced Photon Source was supported by
the U.S. Department of Energy, Basic Energy Sciences, Office of Science under Contract No. DE-
AC02-06CH11357. Use of the BioCARS Sector 14 was supported by the National Institutes of Health, National
Center for Research Resources, under grant number RR007707. Computer resources were provided by the Basic
Sciences Computing Laboratory of the University of Minnesota Supercomputing Institute, and we thank Can
Ergenekan for his support. We also thank Ed Hoeffner at the Kahlert Structural Biology Laboratory (KSBL) at the
University of Minnesota. Use of the KSBL was supported by a Minnesota Partnership for Biotechnology and a
Medical Genomics Grant SPAP-05-0013-P-FY06.
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Figure 1.
The active site and substrate channel in the MCRox1-silent crystal structure (PDB code 1hbn)
(9). Coenzyme F430, CoBSH and HSCoM are drawn as stick colored by atom (carbon: dark
grey). The nickel is displayed as a green sphere, water as a red sphere. Interactions are
drawn as dashed lines, and the corresponding distance is indicted in Angstroms (Å). The
path of the substrate channel was defined in the absence of F430, CoBSH, HSCoM and
water, with the surface closest to the viewer cut away. The figure was generated using
PyMOL (http://www.pymol.org).
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Figure 2.
Drawing of CoBSH analogues; (A) N-5-mercaptopentanoylthreonine phosphate (CoB5SH);
(B) N-6-mercaptohexanoylthreonine phosphate (CoB6SH); (C) N-8-
mercaptooctanoylthreonine phosphate (CoB8SH); (D) N-9-mercaptononanoylthreonine
phosphate (CoB9SH).
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Figure 3.
The active sites and substrate channels of the MCR crystal structures.; (A) MCRCoB5SH; (B)
MCRCoB6SH; (C) MCRHSCoM; (D) MCRCoB8SH; (E) MCRCoB9SH. 2Fo-Fc electron density
map around the CoBXSH analogues, waters in the CoBSH binding part of the channel and
the acetate ion (contoured at 1σ) is shown as a blue mesh. The protein is drawn as cartoon.
CoBXSH and acetate are drawn as stick and colored by atom (carbon: CoB5SH orange;
CoB6SH pale yellow; CoB8SH light blue; CoB9SH magenta; acetate white). Coenzyme F430
and HSCoM are drawn as stick colored by atom (carbon: F430 dark grey; HSCoM medium
grey). The nickel is displayed as a green sphere, and waters as red spheres. The figure was
generated using PyMOL (http://www.pymol.org/).
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Figure 4.
Overlay of CoBSH (from PDB code 1hbn) and the different CoBSH analogues. CoBXSH are
drawn as stick with the thiol represented by a sphere, and colored CoB5SH orange; CoB6SH
pale yellow; CoBSH light green; CoB8SH light blue; CoB9SH magenta. The protein is
drawn as cartoon. Coenzyme F430 and HSCoM are drawn as stick colored by atom (carbon:
F430 dark grey; HSCoM medium grey). The nickel is displayed as a green sphere. The figure
was generated using PyMOL (http://www.pymol.org/).
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Figure 5.
Hydrogen bonding diagram for the water structure modeled in MCRHSCoM. The water
molecules are named as in Figure 3C (W1–W9); WA, WB and WC are water molecules that
are present in all structures (i.e. in concert with the substrate CoBSH and the CoBSH
analogues). Interactions between surrounding residues and the water molecules are drawn as
dashed lines, and the corresponding distance is indicated in Angstroms (Å).
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Figure 6.
Stereo image of the annulus of aromatic amino acids distal of coenzyme F430. The protein is
drawn as cartoon with the side-chains of the aromatic residues drawn as white stick.
CoBSH, (from PDB code 1hbn (9)), CoB8SH and CoB9SH are drawn as stick with the thiols
represented by spheres, and colored CoBSH light green; CoB8SH light blue; CoB9SH
magenta. Coenzyme F430 and HSCoM are drawn as stick colored by atom (carbon: F430
dark grey; HSCoM medium grey). The nickel is displayed as a green sphere. The figure was
generated using PyMOL (http://www.pymol.org/).
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Scheme 1.
Reaction catalyzed by methyl-coenzyme M reductase
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Scheme 2.
Two of the proposed catalytic mechanisms for methyl-coenzyme M reductase; (A)
mechanism I; (B) mechanism II.
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Table 1
X-ray Data Collection, Processing and Refinement Statistics
Data collection and processing statistics
Name of data set
MCRCoB5SH
MCRCoB6SH
MCRHSCoM
MCRCoB8SH
MCRCoB9SH
Measured reflections
1969388
2427498
1440665
1160543
1425506
Unique reflections
553755
446253
405349
211803
401701
Resolution (Å) a
50.0–1.30 (1.35–1.30)
50.0–1.40 (1.45–1.40)
50.0–1.45 (1.50–1.45)
50.0–1.80 (1.86–1.80)
50.0–1.45 (1.50–1.45)
Completeness (%) a
97.1 (78.1)
99.9 (100.0)
99.5 (99.7)
99.8 (100.0)
98.1 (95.4)
R-sym (%) a,b
5.5 (32.9)
7.3 (44.7)
6.2 (44.0)
8.4 (47.7)
5.6 (42.5)
I/σI a
22.3 (3.6)
20.4 (4.0)
20.2 (3.2)
21.8 (3.9)
24.3 (3.2)
Space group
P21
P21
P21
P21
P21
Refinement and model building statistics
Resolution (Å) a
20.49–1.30 (1.33–1.30)
19.89–1.40 (1.44–1.40)
20.15–1.45 (1.49–1.45)
19.93–1.80 (1.84–1.80)
20.07–1.45 (1.48–1.45)
No. of reflection in working set a
525817 (30239)
423854 (25833)
384868 (25791)
201128 (11193)
381474 (23611)
No. of reflection in test set a
27777 (1576)
22348 (1331)
20362 (1319)
10625 (557)
20163 (1210)
R-work (%) c
14.32
13.04
13.47
14.95
13.58
R-free (%) d
16.56
15.53
16.22
19.54
16.44
ESU (Å) R-work/R-free
0.044/0.046
0.049/0.051
0.056/0.059
0.121/0.119
0.057/0.060
No. protein atoms
20087
19960
20265
19750
20036
No. coenzyme atoms
218
220
180
224
272
No. ligand atoms
37
62
52
26
49
No. water molecules
2443
2352
2516
1893
2432
RMS
bond lengths (Å)
0.033
0.033
0.032
0.028
0.032
bond angles (deg.)
2.693
2.625
2.468
2.059
2.549
Ramachandran plot (%)
favored
97.8
97.5
97.6
97.2
97.7
allowed
2.1
2.4
2.3
2.7
2.1
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disallowed
0.1
0.1
0.1
0.1
0.1
Average B-factor (Å2)
protein
12.42
13.35
12.12
17.22
12.73
coenzymes
8.20
9.24
7.25
11.24
8.27
ligands
31.95
35.48
28.29
33.76
32.92
waters
22.95
24.89
23.85
26.79
24.09
over all
13.54
14.57
13.40
18.02
13.93
Occupancy of HSCoM per active site (%)e
90/90
50/50
100/100
90/90
90/85
Occupancy of CoBSH per active site (%) e
50/50
50/50
30/30
50/50
40/40
CoBSH analogue, occupancy per active site (%) e
CoB5SH, 50/50
CoB6SH, 50/50
CoB8SH, 50/50
CoB9SH, 60/60
Other molecule, occupancy per active site (%) e
Acetate, 70/70
aValues in brackets correspond to the highest resolution shell.
bR-sym = ΣhklΣNj=1|Ihkl−Ihkl(j)|/ΣhklN*Ihkl, sum over all reflections and all observations N, with Ihkl(j) intensity of the jth observation of reflection hkl and Ihkl mean intensity of the reflection hkl.
cR-work = Σ||Fo| − |Fc||/Σ|Fo|, where |Fo| = observed structure factor amplitude and |Fc| = calculated structure factor amplitude.
dR-free, R-factor based on 5% of the data excluded from refinement.
eOccupancy of model in each of the two crystallographically independent active sites in the ASU
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Table 2
Distances from analogue thiols.
CoBXS - SCoM distance (Å)
CoBXS - Ni distance (Å)
CoB5SH
7.11/7.11a
9.30/9.30
CoB6SH
6.26/6.26
8.70/8.70
CoB7SH (substrate) b
6.37/6.39
8.73/8.77
CoB8SH
3.75/3.78
6.16/6.17
CoB9SH
3.71/3.68
5.96/5.91
aDistances in the two crystallographically independent active sites in the ASU
bDistances in the 1.16 Å resolution MCRox1-silent structure (PDB code 1hbn)(33)
Biochemistry. Author manuscript; available in PMC 2011 September 7.
|
3M31
|
Structure of the C150A/C295A mutant of S. cerevisiae Ero1p
|
Steps in reductive activation of the
disulfide-generating enzyme Ero1p
Nimrod Heldman,1 Ohad Vonshak,1 Carolyn S. Sevier,2 Elvira Vitu,1
Tevie Mehlman,3 and Deborah Fass1*
1Department of Structural Biology, Weizmann Institute of Science, Rehovot 76100, Israel
2Department of Biology, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139
3Department of Biological Research Support, Weizmann Institute of Science, Rehovot 76100, Israel
Received 8 March 2010; Accepted 9 July 2010
DOI: 10.1002/pro.473
Published online 28 July 2010 proteinscience.org
Abstract: Ero1p is the primary catalyst of disulfide bond formation in the yeast endoplasmic
reticulum (ER). Ero1p contains a pair of essential disulfide bonds that participate directly in the
electron transfer pathway from substrate thiol groups to oxygen. Remarkably, elimination of
certain other Ero1p disulfides by mutation enhances enzyme activity. In particular, the C150A/
C295A Ero1p mutant exhibits increased thiol oxidation in vitro and in vivo and interferes with redox
homeostasis in yeast cells by hyperoxidizing the ER. Inhibitory disulfides of Ero1p are thus
important for enzyme regulation. To visualize the differences between de-regulated and wild-type
Ero1p, we determined the crystal structure of Ero1p C150A/C295A. The structure revealed local
changes compared to the wild-type enzyme around the sites of mutation, but no conformational
transitions within 25 A˚ of the active site were observed. To determine how the C150AC295
disulfide nonetheless participates in redox regulation of Ero1p, we analyzed using mass
spectrometry the changes in Ero1p disulfide connectivity as a function of time after encounter with
reducing substrates. We found that the C150AC295 disulfide sets a physiologically appropriate
threshold for enzyme activation by guarding a key neighboring disulfide from reduction. This study
illustrates the diverse and interconnected roles that disulfides can play in redox regulation of
protein activity.
Keywords: disulfide bond formation; enzyme activation; flavin adenine dinucleotide; lag phase
Introduction
A fundamental question in cell biology is how a bal-
ance between thiols and disulfides is maintained in
the endoplasmic reticulum (ER) to promote efficient
oxidation of proteins while preventing irreversible
mispairing of disulfides.1 The ER sulfhydryl oxidase
Ero1 catalyzes formation of disulfide bonds2,3 and
may also serve as a redox sensor, tailoring its activ-
ity according to the thiol/disulfide balance or the
presence of specific reduced substrates in the com-
partment.4,5 Although Ero1 has certain structural
and mechanistic features in common with other sulf-
hydryl oxidases,6 Ero1 exhibits complex kinetics not
observed in other enzyme families catalyzing the
same chemical reaction. This unique behavior of
Ero1,
which
includes
a
pronounced
lag
phase
observed in assays of catalytic activity on model sub-
strates performed in vitro,4,7 may be a manifestation
of the regulatory feedback mechanism to prevent
over-oxidation of the ER thiol pool.4 Therefore, a
thorough analysis of Ero1 kinetics and their bio-
chemical
and
structural
bases
is
essential
for
Abbreviations: DTT, dithiothreitol; ER, endoplasmic reticulum;
FAD, flavin adenine dinucleotide; PDI, protein disulfide isomer-
ase; Pdi1pred, reduced yeast PDI; PEG, polyethylene glycol; Trx,
E. coli thioredoxin I; Trxred, reduced Trx.
Additional Supporting Information may be found in the online
version of this article.
Grant
sponsors: U.S.-Israel
Binational
Science Foundation;
Kimmelman Center for Macromolecular Assemblies.
Nimrod Heldman and Ohad Vonshak contributed equally to this
work.
*Correspondence to: Deborah Fass, Department of Structural
Biology, Weizmann Institute of Science, Rehovot 76100, Israel.
E-mail: deborah.fass@weizmann.ac.il
Published by Wiley-Blackwell. V
C 2010 The Protein Society
PROTEIN SCIENCE 2010 VOL 19:1863—1876
1863
understanding the origin of the redox balance in the
ER. A previous study presented the finding that
mutation of certain cysteine residues in yeast Ero1
(Ero1p) increases enzyme activity.4 We now address
the
mechanism
by
which
noncatalytic
disulfides
tune the response of the enzyme to thiol species in
the environment.
Yeast Ero1p has 14 cysteine residues and a bound
flavin adenine dinucleotide (FAD) cofactor [Fig. 1(A)].
Two pairs of Ero1p cysteines are on the direct electron-
transfer pathway from substrate to flavin [Fig. 1(B)].
One of these pairs, a Cys-X-X-Cys motif that forms the
active-site disulfide (C352AC355), abuts the isoalloxa-
zine of the FAD and most likely transfers electrons
directly to the cofactor.6,9 The second pair (C100A
C105), the ‘‘shuttle’’ disulfide, is a Cys-X4-Cys motif on a
flexible loop near the active site and appears to mediate
transfer of electrons from substrate proteins to the
active-site disulfide.6,9,10 A third conserved disulfide
(C90AC349) is in the vicinity of the active site but is
not essential for Ero1p activity in vitro or under condi-
tions that have been examined in vivo.4 Its precise role
and the reason for conservation are unclear. Other
disulfides, more distant from the active site, are also
dispensable for activity, but they may nevertheless con-
tribute to regulation of the enzyme. We reported that a
yeast Ero1p mutant lacking the C150AC295 disulfide,
which is 35 A˚ from the FAD isoalloxazine, has a short-
ened lag phase in enzyme assays on model substrates
in vitro, increases the intracellular ratio of oxidized to
reduced glutathione, and decreases viability of yeast.4
These observations demonstrate an important role for
Ero1p redox centers off the catalytic electron transfer
pathway and distant from the active site.
A
remaining
question
is
how
loss
of
the
C150AC295 disulfide in Ero1p has such profound
effects on enzyme activity and on redox homeostasis in
cells, and whether the C150A/C295A mutant can pro-
vide insight into the autoregulatory mechanism of the
wild-type enzyme. The structure of Ero1p, obtained pre-
viously from two crystal forms (hexagonal at 2.8 A˚ reso-
lution and centered orthorhombic at 2.2 A˚ resolution),6
suggested the possibility of redox-dependent conforma-
tional changes. The temperature factor distribution
[Fig. 1(A)] and a comparison of the structures from the
two crystal forms highlighted the flexibility of loops in
what is designated the ‘‘top’’ of the Ero1p structure. A
congregation of disulfides in the loop-rich region and at
the interface with the 10-helix core of the enzyme, to-
gether with numerous exposed hydrophobic residues
and a richly featured surface containing hydrophobic
pockets and grooves [Fig. 1(C)], suggests that reduction
or elimination of disulfides may enable conformational
rearrangement of the top portion of the enzyme, with
consequent effects on enzyme activity. Conformational
changes upon disulfide reduction may occur also in
mammalian Ero1a, as supported by changes in intrinsic
tryptophan/tyrosine fluorescence upon reduction or
mutation of certain disulfides.11
To test the hypothesis that elimination of the
C150AC295 disulfide in Ero1p causes conformational
Figure 1. A: Ribbon diagram of wild-type Ero1p (Protein Data
Bank code: 1RP4) colored according to temperature factor, with
red corresponding to high temperature factor regions and blue
to low. The FAD is shown in sticks. Disulfide bonds and the
unpaired C208 are illustrated in ball-and-stick representation
and labeled. Eleven of the 14 cysteines in full-length yeast Ero1p
are present in the truncated protein that was crystallized. B: The
two-electron transfer events involved in oxidation of Pdi1p by
Ero1p are illustrated as paired arrows.
C: The molecular surface of Ero1p is shown from two angles
with cysteine sulfur atoms colored yellow and surface-exposed
hydrophobic side-chains colored green. Aromatic side chains
are in a darker shade. The table lists the solvent-accessible
surface areas (SAS) of the side chains of the indicated cysteines
as determined using the program areaimol (CCP4 package).8
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PROTEINSCIENCE.ORG
Activation of Sulfhydryl Oxidase Ero1p
changes that increase enzyme activity, we deter-
mined the structure of the C150A/C295A de-regu-
lated Ero1p mutant by X-ray crystallography. Inter-
pretation
of
this
structure
was
facilitated
by
enzymatic assays of wild-type and mutant Ero1p oxi-
dizing either native or model substrates. Tandem
mass spectrometry (LC-MS/MS) analysis of Ero1p di-
sulfide connectivity during activation and turnover
revealed the basis for the hyperactivity of the C150A/
C295A mutant and provided insight into the natural
activation mechanism of the enzyme.
Results
Electron transfer kinetics of the C150A/C295A
Ero1p mutant
Increased turnover rates and a shortened lag phase
for the C150A/C295A Ero1p mutant were previously
reported based on oxygen consumption assays.4 To
obtain greater precision at early time points, we per-
formed similar reactions using stopped-flow fluorim-
etry to monitor oxidation of reduced thioredoxin
(Trxred) by Ero1p [Fig. 2(A)]. Free FAD was used as
the electron acceptor for Ero1p instead of oxygen,
since reduction of the flavin is detectable by a
decrease in fluorescence. Two recombinant wild-type
Ero1p
constructs,
spanning
amino
acid
residues
10–424 or 56–424,4,6,7 were examined. These two
constructs differ by approximately twofold in their
activities, but their progress curves are identical in
shape, as can be seen by scaling the time axes
[Fig. 2(B)]. Thus, the lag phase observed during
oxidation of Trxred is an inherent property of the
wild-type enzyme on this substrate regardless of
absolute activity. The lag phase is also independent
of whether molecular oxygen or FAD is used as the
electron acceptor. In contrast, the C150A/C295A mu-
tant has a progress curve qualitatively different
from either wild-type version of Ero1p [Fig. 2(B)].
Ero1p C150A/C295A structure
The dramatic impact on enzyme kinetics of the
C150A/C295A
double
mutation
prompted
us
to
investigate the structure of this Ero1p variant. We
sought to reveal any conformational differences com-
pared to wild type that may explain the differences
in activity and pre-steady state kinetics. Mutant pro-
tein was produced in E. coli with approximately one-
third the yields obtained for wild type and was iso-
lated using a comparable protocol.6 As for wild-type
Ero1p,
crystals
of
Ero1p
C150A/C295A
were
obtained in both hexagonal and centered orthorhom-
bic forms. The orthorhombic form was chosen for
further study because it diffracted to higher resolu-
tion. The Ero1p C150A/C295A structure was deter-
mined using phases calculated from a partial model
of wild-type Ero1p (see Materials and Methods sec-
tion). The structure was refined using diffraction
data to 1.85 A˚ resolution (Table I).
Conformational differences compared to wild-
type Ero1p were observed in the region of the miss-
ing C150AC295 disulfide (Fig. 3, left panel). These
differences occur primarily in the a-helix down-
stream of C295 (designated helix a6).6 The a6 helix
is five residues longer at the amino terminus in the
C150A/C295A
mutant,
extending
from
L297
as
opposed to I302, allowing D296 to cap the helix. In
the wild-type structure, participation of C295 in the
disulfide with C150 prevents D296 from capping the
a6 helix, and the shorter version of the helix is
instead partially capped in trans by the side chain of
S293. The two Cb atoms of A150 and A295 are 9.2 A˚
apart in Ero1p C150A/C295A, in contrast to the 3.8
A˚ distance between the analogous cysteine Cb atoms
of the wild-type enzyme. In summary, the region in
the vicinity of residues 150 and 295 apparently
Figure 2. Enzyme progress curves of various Ero1p
constructs. Residue number boundaries appear in subscript
in the label of each curve. A: Ero1p activity was monitored
by stopped-flow fluorimetry. Ero1p, FAD, and Trxred were
mixed under anaerobic conditions at final concentrations of
1, 100, and 50 lM, respectively. The exogenous FAD
served as the electron acceptor for Ero1p, and
fluorescence decay of FAD upon reduction reported the
progress of the reaction. B: Superposition of fluorescence
data by rescaling Ero110–424 and Ero156–424 C150A/C295A
data along the time axis provides evidence for altered
regulation of Ero156–424 C150A/C295A activity.
Heldman et al.
PROTEIN SCIENCE VOL 19:1863—1876
1865
relaxes in the double cysteine-to-alanine mutant to a
conformation incompatible with the disulfide but
containing more regular and extensive secondary
structure.
Despite these differences, most of the Ero1p
C150A/C295A structure, including the active-site
region (Fig. 3, right panel), is similar to that previ-
ously observed for wild-type Ero1p. The resolution of
the diffraction data was significantly better than
obtained for wild type, but regions that previously
gave poor electron density (i.e., the 155–165 and
108–116 loops) were also apparently flexible in the
Ero1p C150A/C295A crystals. No major differences
were seen in the solvent exposure of other disulfide
bonds in the structure. The relatively minor differ-
ences in the C150A/C295A Ero1p structure do not
rule out the possibility of global changes in protein
dynamics, but any putative changes in dynamics did
not prohibit crystallization. The absence of a clear
structural explanation for the increased activity of
the C150A/C295A mutant led us to initiate a more
detailed biochemical analysis of the reductive activa-
tion process.
Gel electrophoretic analysis of disulfide
reduction during Ero1p activation
Reduction of a series of disulfide bonds in Ero1p
upon encounter with reducing substrate has been
observed by changes in migration rate of the enzyme
on denaturing gels.4 These experiments are per-
formed by incubating Ero1p with substrate and
blocking the reaction after various times using rapid
alkylating agents to modify free thiols. For our cur-
rent studies, we compared reactions blocked with
the small alkylating agent N-ethyl maleimide (NEM)
versus the larger maleimide derivative 4-acetamido-
40-maleimidylstilbene-2,20-disulfonic acid (AMS) and
analyzed the samples on both reducing and nonre-
ducing denaturing gels. These experiments extend
our
previous
work
with
AMS-modified
samples
resolved under nonreducing conditions4 and allow
for additional insight into the reductive activation
process of Ero1p.
The thiol trapping experiments provide informa-
tion on the number of free cysteines that become
reduced and the nature of any remaining disulfides
in Ero1p at a given time after substrate addition.
When samples are resolved under reducing condi-
tions, the migration rate of Ero1p reflects the abso-
lute mass of the enzyme, which depends on both the
number of cysteines modified and the type of alkyl-
ating agent used. AMS contributes 510 D per cys-
teine modified, whereas NEM contributes 125 D.
The small change in mass due to NEM alkylation of
Table I. Summary of Data Collection and
Refinement Statistics
Ero1p mutant
C150A/C295A
C143A/C166A
Space group
C2221
P62
Unit cell
parameters (A˚ )
73.4 132.8
102.7
107.2 107.2
124.2
Resolution (A˚ )
500.0–1.85
50.0–3.2
Completeness (%)
99.0
99.9
Redundancy
4.9
12.4
Rsym
a
0.037
0.236
hI/rIi
15.4
5.7
Total reflections/test set
42,641/2864
13,131/672
Rwork/Rfree
b
0.209/0.238
0.244/0.293
Rms deviations from ideality
Bonds (A˚ )
0.005
0.008
Angles ()
1.314
1.847
Number of atoms
Protein
2899
2842
Water
211
12
FAD
53
53
NEM
9
9
Cd2þ
1
1
a Rsym
¼
RhklRi|Ii(hkl)
hI(hkl)i|/RhklRiIi(hkl),
where
Ii(hkl) is the observed intensity and hI(hkl)i is the average
intensity for i observations.
b Rwork, Rfree ¼ R||Fobs| |Fcalc||/R|Fobs|, where Fobs and
Fcalc are the observed and calculated structure factors,
respectively. A set of reflections (6.7%) were excluded from
refinement and used to calculate Rfree.
Figure 3. Superposition of the structures of wild-type and C150A/C295A Ero1p. In the central panel, both structures are shown
in beige. In the zoom views to either side, the Ero1p C150A/C295A structure is shown in dark red. The left zoom window is
rotated relative to the central view to show structural differences with greater clarity. The dotted line indicates a region of poor
electron density that could not be modeled in the mutant structure and was backbone traced in the wild-type structure. The right
zoom window shows the close correspondence of the wild-type and mutant enzymes in the region of the active site.
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PROTEINSCIENCE.ORG
Activation of Sulfhydryl Oxidase Ero1p
free thiols does not result in an observable shift in
migration of Ero1p on a reducing, denaturing gel. In
contrast, under the same gel conditions, Ero1p with
AMS-modified cysteines can be distinguished from
unmodified
Ero1p.
When
samples
are
analyzed
under nonreducing conditions, the migration rate of
Ero1p is influenced not only by the change in mass
from modified cysteines but also by the presence or
absence of disulfide bonds between cysteines distant
from one another in amino acid sequence, which
affects the hydrodynamic radius of the SDS-bound,
denatured protein. For AMS-modified samples, the
mobility of Ero1p will be influenced both by the con-
tribution of any disulfides to the hydrodynamic ra-
dius and by the 510 D mass added per modified
thiol. When NEM is used, the mass change due to
thiol derivation is negligible, and the apparent mo-
bility under nonreducing conditions will primarily
reflect the increase in hydrodynamic radius upon
reduction of disulfides between cysteines distant
from one another in amino acid sequence.
In experiments performed using Trxred as a sub-
strate, Ero1p was converted stepwise to a set of
more slowly migrating species (Fig. 4), as reported
previously.4 Appearance of these species correlates
with increased enzyme activity.4 At later times in
the reaction, when most of the substrate had been
oxidized, Ero1p became re-oxidized to its initial state
of low activity. For the Ero1p C150A/C295A mutant,
the more slowly migrating species appeared earlier
in the reaction and disappeared earlier as well (Fig.
4). Furthermore, the most slowly migrating band
was never the most populated species in the Ero1p
C150A/C295A reaction, whereas it was the dominant
species at a certain point (1 min) in the reaction
with the wild-type enzyme.
As
explained
above,
the
different
migration
rates of Ero1p species reflect different disulfide
Figure 4. Disulfide reduction during catalysis by Ero1p and the de-regulated mutant C150A/C295A. Enzyme at a final
concentration of 2 lM was mixed with 100 lM Trxred. Aliquots were removed from the reactions at the indicated time points,
and disulfide status was preserved by blocking with maleimide reagents. The migration of Ero1p on SDS-PAGE at the various
time points was examined and compared with wild-type Ero1p (WT), the indicated Ero1p mutants, and DTT-treated Ero1p
(WT reduced). (A) The Trxred oxidation reaction was blocked with AMS and separated by SDS-PAGE under nonreducing
conditions. The asterisk (*) indicates a species that migrates slower than Ero1p C90A/C349A when blocked with AMS, but
faster than this disulfide mutant when blocked with NEM (compare with panel C). (B) The same experiment as in (A) was
performed, but the samples were reduced with DTT after AMS treatment before separation by SDS-PAGE. (C) Reactions
were blocked with NEM and applied to the gel under nonreducing conditions.
Heldman et al.
PROTEIN SCIENCE VOL 19:1863—1876
1867
connectivity. A C90A/C349A mutant, which lacks the
disulfide that closes the largest loop in the protein,
shows a major shift in mobility relative to wild-type
Ero1p; this mutant produces the largest increase in
hydrodynamic radius of any single disulfide mutant
relative to wild type (Fig. 4). Notably, when the reac-
tion of wild-type Ero1p with substrate was blocked
with AMS, the most slowly migrating species [aster-
isk in left panel of Fig. 4(A)] ran above the position
of the C90A/C349A mutant. However, the same reac-
tion blocked with NEM resulted only in species that
migrated more quickly than the C90A/C349A mu-
tant [asterisk in left panel of Fig. 4(C)]. The lack of
a maximal change in the hydrodynamic radius of
NEM-treated Ero1p implies that the C90AC349 di-
sulfide may be intact under these reaction condi-
tions. For Ero1p with an intact C90AC349 bond, the
shift observed with AMS likely reflects the combined
effect of the increase in mass due to thiol modifica-
tion and the lesser hydrodynamic changes observed
during reduction of other disulfides less distant in
sequence than C90AC349. Alternatively, the slowest
migrating Ero1p species could lack C90AC349, and
the moderate migration shift may reflect the pres-
ence of an alternate long-range disulfide formed by
thiol/disulfide exchange. A catalytic intermediate
with a disulfide between C105 and C352 has been
previously trapped in vivo,10 and the presence of
this long-range disulfide could perhaps account for
faster mobility even if C90AC349 were absent.
At early time points (<1 min), Ero1p blocked
with NEM did not exhibit a significant shift in mo-
bility on the gel [Fig. 4(C), left panel]. In contrast,
Ero1p blocked with AMS did display species with
lower electrophoretic mobility at these times on both
nonreducing and reducing gels [Fig. 4(A,B)], sug-
gesting reduction of one or more disulfides that do
not greatly affect the protein hydrodynamic radius
under denaturing conditions. What appeared as a
single species in the NEM-blocked reaction was
resolved into two distinct species by AMS treatment,
which is most readily observed under nonreducing
conditions but is also apparent as a closely migrat-
ing
pair
of
bands
under
reducing
conditions.
Between 0.1 and 0.5 min, which is the time-frame
corresponding to enzyme activation (Fig. 2), the ratio
of the upper to lower bands of the 40 kD doublet
for the AMS-blocked reaction increased [Fig. 4(A)].
For early time points in reactions using the
Ero1p C150A/C295A mutant, the single major band
seen in the NEM-blocked reactions was similarly
resolved into two species by blocking with AMS.
However, in this case, the mutant Ero1p appeared
maximally reduced at 0.1 min, and the ratio of the
upper
to
lower
bands
of
the
47
kD
doublet
decreased between 0.1 and 0.5 min as the Ero1p mu-
tant protein was rapidly re-oxidized. Notably, signifi-
cant retardation of mobility was seen for the Ero1p
C150A/C295A mutant at the shortest time point.
However, the shifted species migrate faster than
fully reduced Ero1p or the C90A/C349A mutant and
thus
are
not
likely
to
reflect
reduction
of
the
C90AC349 disulfide.
Mass spectrometric analysis of Ero1p activation
To directly map the Ero1p disulfides that were
reduced in each gel-shifted species described above,
we subjected to in-gel proteolysis and tandem mass
spectrometry bands from AMS-blocked reactions run
under
nonreducing
conditions
(Fig.
5).
Earlier
attempts at disulfide mapping of Ero1p using ma-
trix-assisted laser desorption/ionization time of flight
(MALDI-TOF)
mass
spectrometry
suggested
the
presence of disulfides inconsistent with the X-ray
crystal structure (data not shown), revealing the
risk of over-interpreting data consisting of only par-
ent peptide masses. LC-MS/MS data, in contrast,
enable
more
reliable
assignments
of
peptide
identities.
The first LC-MS/MS experiment [Fig. 5(B)] was
performed without in-gel reduction and alkylation,
such that cysteine connectivity in disulfides was pre-
served. The first and most rapidly migrating par-
tially reduced species observed showed C100 in
AMS-modified
form
and
the
C143AC166
and
C150AC295 disulfides intact. The next shifted spe-
cies showed C166 alkylated and C150AC295 in di-
sulfide form. The most slowly migrating species
showed the active-site disulfide (C352–355) to be
intact and C150, C166, and C349 to be AMS-modi-
fied. Again, it should be noted that this species,
when blocked with NEM rather than AMS, migrates
faster than the C90A/C349A mutant [Fig. 4(C), as-
terisk], suggesting that it nevertheless contains a
long-range disulfide. In this set of experiments, we
did not observe any non-native disulfides, that is,
those not present in the Ero1p crystal structure.
To complement this information, we did a sec-
ond LC-MS/MS experiment, in which Ero1p species
in gel slices from the same AMS-blocked experiment
were reduced with dithiothreitol (DTT) and alky-
lated with iodoacetamide before in-gel proteolysis.
This experiment revealed which cysteines were par-
ticipating in disulfides (carbamidomethylated) and
which were reduced (AMS-modified) at each time
point [Fig. 5(C)]. Disulfide connectivity information
was lost, but peptide recovery could in principle be
altered or improved by reduction of the disulfide
bonded peptides. This experiment was performed
both on the wild-type enzyme and on Ero1p C150A/
C295A. For both versions, the first Ero1p thiol to be
detected in AMS-modified form was C143. The spe-
cies showing reduced and modified C143 increased
in intensity between 0.1 and 0.5 min for wild-type
Ero1p
but
was
present
as
the
dominant
band
already at the 0.1 min time point of the Ero1p
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PROTEINSCIENCE.ORG
Activation of Sulfhydryl Oxidase Ero1p
C150A/C295A reaction. Analysis of the highest band
in the wild-type reaction revealed populations of spe-
cies with C150 and C349 reduced as well. The high-
est band of the C150A/C295A mutant also showed
reduction of C349. Overall, the mass spectrometry
data revealed a progressive loss of disulfides in wild-
type Ero1p in the order C100AC105, C143AC166,
C150AC295,
and
perhaps
C90AC349.
For
the
C150A/C295A mutant, the most striking and consist-
ent observation was the appearance of C143 as an
AMS-modified thiol at early timepoints.
The above mass spectrometry analyses were
conducted on Ero1p after reaction with a model sub-
strate. To determine how a native substrate affects
noncatalytic disulfides of Ero1p, we performed com-
parable experiments using yeast Protein Disulfide
Isomerase (Pdi1p). Ero1p was mixed with reduced
Pdi1p (Pdi1pred), and the reaction was blocked with
either NEM or AMS. Because Pdi1p migrates to a
similar position on electrophoretic gels as the more
reduced species of Ero1p observed in the experi-
ments with Trxred, Ero1p was visualized with a fluo-
rescent stain specific to poly-histidine tags, and an
untagged
version
of
Pdi1p
was
used.
Wild-type
Ero1p blocked with NEM at various points in the
reaction showed no species with retarded migration
[Fig. 6(A), left panel], whereas blocking with AMS
revealed a minor population of modified Ero1p [Fig.
6(B), left panel]. In contrast, a significant population
of Ero1p C150A/C295A was shifted upon addition of
Pdi1pred, after blocking with either NEM or AMS
[Fig. 6(A,B), right panels]. This observation suggests
that a disulfide between nonvicinal cysteines is
reduced in the slowly migrating fraction. Disulfide
rearrangement is also a formal possibility. In these
studies, no Ero1p species similar to the most slowly
migrating species seen in the reactions with Trxred
were observed, though Ero1p and Ero1p C150A/
C295A did oxidize Pdi1pred under the conditions of
the experiment [Fig. 6(C)]. Therefore, Ero1p is func-
tional on Pdi1p without quantitative reduction of
multiple regulatory disulfides.
After establishing that the partially reduced
species of Ero1p or Ero1p C150A/C295A obtained
upon reaction with Pdi1pred do not comigrate with
Pdi1p itself (55 kD), we aimed to identify the disul-
fide that becomes reduced in the C150A/C295A mu-
tant. We incubated Ero1p C150A/C295A for short
times with Pdi1pred, blocked the reaction with either
Figure 5. Mass spectrometric identification of partially
reduced Ero1p species. A: The upper panels of Figure 4
(reaction time-courses of wild-type and C150A/C295A Ero1p
blocked with AMS) are reproduced, with the bands that were
subjected to mass spectrometry indicated by ovals.
B: A summary of the mass spectrometry results from each
band is indicated on a map of the Ero1p primary structure. All
disulfides observed in the Ero1p crystal structures were found
in the time zero band (i.e., in the absence of substrate), as
indicated by the linked open circles, and no non-native
disulfides were detected. The homogeneity of the time zero
band, and the fact that this species was obtained from pure
protein stock rather than from aliquots removed from enzymatic
reactions, may explain the improved peptide coverage
compared to the other bands analyzed. Reduction of
C143AC166, indicated by unpaired dark circles, was seen in
bands that appeared earlier than those showing reduction of
C150AC295. It should also be noted that reduction of
C143AC166 does not result in a mobility change in the
wild-type enzyme blocked with NEM, as can be seen by
comparing Figures 4(C) and 5(A). Reduction of this disulfide
does, however, cause a shift when it occurs in the C150A/
C295A background. C: The indicated bands were subject to
reduction and alkylation with iodoacetamide before proteolytic
cleavage and mass spectrometry analysis. Open circles indicate
cysteines that were detected as modified by iodoacetamide,
whereas dark circles indicate cysteines that were detected as
being modified by AMS, and thus had been reduced during the
reaction with Trxred. Half-filled circles represent cysteines
detected in both states in the same experiment.
Heldman et al.
PROTEIN SCIENCE VOL 19:1863—1876
1869
NEM or AMS, separated the proteins by SDS-PAGE,
and stained with Coomassie (gel not shown). In the
AMS-treated sample, the higher band derived from
the Ero1p mutant comigrated with a Pdi1p contami-
nant or degradation product and was not analyzed.
In contrast, in the NEM-treated sample, the band
apparently corresponding to that labeled with an as-
terisk in the right panel of Figure 6(A) could be
identified and excised. Tandem mass spectrometry
analysis of this band showed both C143 and C166
in
NEM-modified
form,
demonstrating
that
C143AC166 had been reduced by Pdi1pred. A single
six-amino acid peptide containing reduced and modi-
fied C90 was also observed, although the species an-
alyzed migrated much more quickly in the gel than
the C90A/C349A mutant and is thus likely to con-
tain a long-range disulfide. Displacement of C90 by
attack of C143, or more likely C166 (see below), on
C349 might occur in a fraction of the Ero1p mole-
cules, yielding a species with similar hydrodynamic
radius to that having C143 and C166 reduced and
the remaining disulfides intact.
Ero1p C143A/C166A structure
The identification of C143AC166 as the disulfide
that is reduced much more rapidly in the hyperac-
tive C150A/C295A Ero1p mutant prompted us to
investigate the structural changes that occur in
Ero1p upon removal of this disulfide. The Ero1p mu-
tant C143A/C166A was produced in E. coli with
20% the yields obtained for wild type. Crystals
were obtained in the primitive hexagonal form with
unit cell dimensions similar to those of crystals
obtained with wild-type Ero1p in this space group
(Table I). The Ero1p C143A/C166A structure was
refined using diffraction data to 3.2 A˚ resolution.
Figure 6. The reaction between 2 lM Ero1p or Ero1p C150A/C295A and 75 lM Pdi1pred was blocked with (A) NEM or (B)
AMS after various times and subjected to SDS-PAGE under nonreducing conditions. The band indicated by an open
arrowhead in the wild-type Ero1p time course is a contaminant from the Pdi1p preparation, as it appears also in the Pdi1p
control lane. A significant fraction of Ero1p C150A/C295A shows retarded mobility at early time points when blocked with
either NEM or AMS (asterisks). A band corresponding to that marked with an asterisk in NEM-blocked Ero1p C150A/C295A
was analyzed by LC-MS/MS, and the C143AC166 disulfide was found to be reduced. (C) Oxidation of Pdi1p by Ero1p and
Ero1p C150A/C295A under similar conditions as the experiments in (A) and (B). Pdi1pred was mixed with wild-type or mutant
Ero1p. Aliquots were removed at various times and reacted with PEG-maleimide of 5 kD. Mal-PEG modified Pdi1p species
are indicated in the top portion of the gel. The band labeled Pdi1pox has both active sites oxidized and was thus resistant to
PEGylation.
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Activation of Sulfhydryl Oxidase Ero1p
As for the C150A/C295A mutant, local rear-
rangements of the polypeptide chain were seen in
the C143A/C166A around the mutation site. How-
ever, in contrast to the C150A/C295A mutant, in
which both polypeptide segments containing the cys-
teine-to-alanine mutations acquired more regular
secondary
structure
upon
mutation,
the
C143A/
C166A double mutant is more disordered than wild
type [Fig. 7(A)]. A cluster of acidic residues (D137,
D138, D140, D141, and E142) immediately upstream
of C143A assumes an alternate conformation [Fig.
7(B)], without an increase in secondary structure.
The region surrounding C166A loses its connection
to the helical core of the enzyme, and the poly-
peptide chain between residues 154 and 175 cannot
be traced in electron density maps due to poor elec-
tron density [Fig. 7(C)]. In the wild-type Ero1p
structures and in C150A/C295A, electron density
corresponding to the region around residues 155–
165 was also poor or uninterpretable. However, the
disordered region is significantly extended, in the
direction of the active site, in the C143A/C166A
mutant.
Kinetics studies of single and double mutants
of the C143AC166 disulfide
The structural changes seen in the Ero1p C143A/
C166A mutant are not sufficient to activate the
enzyme; this mutant, when examined previously,
was found not to be hyperactive but rather slightly
slower than wild-type Ero1p.4 To determine whether
the thiol form of either cysteine participating in this
bond may have a direct role in activation, we con-
structed
the
single
cysteine-to-alanine
mutants
C143A and C166A. The C166A mutant behaved sim-
ilarly to the C143A/C166A double mutant. However,
the rate of oxidation of Trxred by the C143A mutant
was found to be indistinguishable from the rate of
oxidation by the de-regulated C150A/C295A mutant
in a gel-based assay [Fig. 7(D)]. This observation
Figure 7. C143 and C166 play different roles in the Ero1p structure. A: Stereo image of a superposition of the structures of
wild-type (colored beige) and C143A/C166A Ero1p (dark red). Disulfides and the carboxy terminus are labeled. The endpoints
of a large missing loop are indicated by circles and labeled with the number of the last modeled residue in each case.
Mutation of C143 and C166 increases the length of the disordered loop such that an additional 10 residues cannot be
modeled. To emphasize the missing loop, the view is different from in Figure 3, but it corresponds to the summary Figure 8.
B: A close-up view of the region around C143 shows the local structural differences between the wild-type Ero1p structure
and the C143A/C166A mutant. The Cb atom of the alanine at position 143 in the mutant is shown in blue and labeled. Side
chains of three acidic residues immediately upstream of C143 are shown as blue and red sticks. To illustrate the extent of the
conformational differences in this area, the side chain of D140 is labeled in both the mutant and wild-type structures. C: A
2Fo-Fc electron density map, contoured at 1r, illustrates the lack of interpretable density corresponding to the loop
containing the C166A mutation. D: Activities of single mutants disrupting the C143AC166 disulfide obtained from a gel-based
Trx oxidation assay. Oxidation of Trxred by the indicated Ero1p variants was blocked at various time points by addition of
PEG-maleimide 5 kD. Oxidized Trx is resistant to PEG modification and thus migrates faster than modified Trx in SDS-PAGE.
The band intensities of oxidized and reduced/modified Trx were quantified and plotted as fraction reduced. The C143A
mutant, which retains C166, is hyperactive, but the C166A mutant, retaining C143, is not. This finding points to a role for the
C166 thiol in Ero1p activation.
Heldman et al.
PROTEIN SCIENCE VOL 19:1863—1876
1871
suggests that liberating C166 is an important step
in
activating
Ero1p.
However,
the
mutagenesis
experiment did not rule out the possibility that the
C143A mutant is hyperactive for the trivial reason
that the unpaired C166 disrupts the C150AC295 di-
sulfide and produces a species that mimics C150A/
C295A. Indeed, mass spectrometry analysis of the
C143A mutant showed some C295 in reduced and
alkylated form, suggesting that C166 displaces it
and forms a disulfide with C150 in a fraction of the
molecules. It should be noted that this species was
not
observed
in
any
experiment
in
which
C143AC166 became reduced during an enzymatic
reaction, suggesting that it is an artifact of lengthy
exposure of C166 during enzyme preparation and
purification.
The
activity
of
C143A
Ero1p
on
Pdi1pred was greater than that of wild-type Ero1p
but not as rapid as C150A/C295A (data not shown).
This observation may indicate that a non-native di-
sulfide
bond
formed
when
C143
is
missing
is
reduced less effectively by Pdi1pred than by Trxred.
Together,
the
study
of
single-cysteine
mutants
C143A and C166A supports the conclusion based on
crystallographic studies that the C166 region has a
large range of motion when freed from C143, and
suggests a specific role for C166.
Discussion
The mechanism by which Ero1p activity is controlled
by encounter with reducing substrates is directly
related to the ability of the enzyme to maintain re-
dox balance in the ER. The various Ero1p disulfides
have distinct roles in catalysis and control of enzyme
activity, and the experiments presented here were
designed to dissect these roles. The prior observation
that eliminating the C150AC295 disulfide of yeast
Ero1p increases enzyme activity, coupled with the ob-
servation of cascaded reduction of Ero1p disulfides in
gel assays using Trxred as a substrate, suggested that
reduction of C150AC295 is part of the series of
events that results in activation of the enzyme.4 The
reduction of C150AC295 was considered as a possible
early event in the cascade, whereas the longest range
disulfide (i.e., C90AC349) was surmised to open later
in the reaction.4 One puzzle in this model was the
implication that the C150AC295 disulfide should be
reduced rapidly in the C90A/C349A Ero1p mutant to
yield a fully activated enzyme. The C90A/C349A mu-
tant would thus be expected to have a short lag phase
and be hyperactive, but instead it has only a slightly
shortened
lag
phase
and
essentially
wild-type
activity.
The LC-MS/MS results presented here consis-
tently showed reduction of the C143AC166 disulfide
preceding reduction of other noncatalytic disulfides.
The opening of C143AC166 first in the cascade was
unexpected, considering the lack of solvent exposure
of this disulfide in the Ero1p crystal structure6 and
the observation that the C143A/C166A double muta-
tion did not activate Ero1p.4 The disulfide mapping
further indicated that reduction of C150AC295 is
actually a late step in Ero1p activity assays on
Trxred. This result implies that the increased activity
of
the
C150A/C295A
mutant
cannot
simply
be
ascribed to removal of an initial hurdle in the reduc-
tive activation process. Whether reduction of the
C150AC295 disulfide is required as the second step
in the activation of Ero1p is difficult to test using
Trxred
as
a
substrate,
since
reduction
of
the
C150AC295 disulfide inevitably followed reduction
of C143AC166 temporally (Fig. 5). In contrast to the
relative ease of eliminating a disulfide bond by mu-
tagenesis, specifically stabilizing a disulfide to test
the effect of a lack of its reduction presents a major
experimental challenge.
Remarkably,
the
observations
presented
here
with
the
native
substrate
Pdi1pred
suggest
that
reduction of multiple Ero1p disulfides may not be
required before Ero1p can oxidize substrate. When
Pdi1pred was mixed with wild-type Ero1p, no shifted
bands corresponding to reduction of C150AC295 (or
the longer range disulfide C90AC349) were observed
at any point in the reaction [Fig. 6(A), left panel],
although wild-type Ero1p does oxidize at least one of
the domains of Pdi1pred in this time frame [Fig. 6(C),
left panel]. Stoichiometric reduction of C150AC295 is
thus not an obligatory step in generating active
Ero1p. It is possible, however, that Pdi1pred reduces
multiple disulfides in an undetectable subpopulation
of Ero1p, and this fraction is then extremely active
on Pdi1pred and performs all the substrate oxidation
observed, while the vast majority of Ero1p molecules
remain oxidized and inactive. We find this explana-
tion unlikely. If Pdi1pred were inefficient at activating
Ero1p but then served as an excellent substrate of
the activated fraction, lag phase kinetics would be
observed for oxidation of Pdi1pred. In fact, the lag
phase is more pronounced during oxidation of Trxred
than of Pdi1pred.
ince reduction of C150AC295 does not seem to be
required for Ero1p turnover, we are left with the
question of why an enzyme variant lacking this disul-
fide shows dramatically increased activity, on both
model and native substrates.4,12 The major finding
presented here is that the C150AC295 disulfide
affects the reactivity of the neighboring disulfide,
between C143 and C166, and that opening of the lat-
ter correlates with exit from the lag phase and
increased oxidase activity. The non-native substrate
Trxred reduces C143AC166 in wild-type Ero1p appa-
rently to completeness, whereas the native substrate
Pdi1pred is inefficient at reducing this disulfide or at
maintaining it reduced as Ero1p competes to re-oxi-
dize it. When C150AC295 is eliminated by mutation,
the C143AC166 disulfide is reduced more rapidly by
Trxred and more extensively by Pdi1pred. The greater
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PROTEINSCIENCE.ORG
Activation of Sulfhydryl Oxidase Ero1p
portion of the enzyme with the C143AC166 disulfide
reduced could explain the greater activity of the
C150A/C295A mutant compared to wild-type Ero1p
in the oxidation of Pdi1pred.
Differences in substrate oxidation kinetics have
also been observed for cysteine mutants in recombi-
nant human Ero1a. In particular, mutation of Ero1a
C131 increased slightly the rate of oxidation of
Pdi1p in vitro11 and increased cellular GSSG levels.5
The explanation for increased activity of the human
Ero1a C131A mutant is distinct from the mechanism
outlined herein for Ero1p C150A/C295A. Ero1a C131
was proposed to form an inhibitory disulfide with
Ero1a C94,11 which is one of the cysteines partici-
pating in the shuttle disulfide of the mammalian
enzyme. The increased activity of the Ero1a C131A
mutant was proposed to reflect loss of the inhibitory
C94AC131 disulfide and ability of the shuttle disul-
fide
between
C94
and
C99
to
form
properly.
Although yeast Ero1p does not oxidize Trxred signifi-
cantly at time zero of the reaction, an inhibitory di-
sulfide formed between a regulatory and a shuttle
cysteine cannot explain this inhibition; an intact
shuttle disulfide was observed in the crystal struc-
ture of yeast Ero1p and by disulfide mapping using
LC-MS/MS [Fig. 5(B) and Supporting information].
If a non-native disulfide were to be present during
reactions of yeast Ero1p, it would most likely be
activating rather than inhibitory. However, numer-
ous mass spectrometry samples of Ero1p trapped
during reactions with Trxred have failed to reveal
non-native disulfides, despite high peptide coverage
of the protein sequence and excellent recovery of
native disulfides. If such species exist, they may be
transient or poorly populated.
If not by engaging shuttle cysteines, then how
might Ero1p regulatory disulfides restrain enzyme
activity? For both C143AC166 and C150AC295, the
disulfide constrains the polypeptide to conformations
different from those observed when the disulfides
are removed. In particular, we observed that the
entire loop between residues 155 and 175 becomes
disordered when C166 is not tethered to C143, as
shown in the structure of the Ero1p C143A/C166A
mutant. The dramatically extended range of motion
of this loop may allow it to impact the active site. As
the C143A/C166A double mutant is not hyperactive,
however, C166 may be specifically required for these
effects. What changes in the active-site region might
increase the rate of catalysis? One hypothesis is that
the active-site disulfide (C352–C355) of Evolp is
poorly accessible for redox communication with the
shuttle disulfide (C100AC105). In the structure of
another yeast sulfhydryl oxidase, Erv2p,13 the shut-
tle disulfide was observed within dithiol-disulfide
exchange distance of the active-site disulfide. In con-
trast,
the
active-site
disulfide
of
Ero1p
is
less
approachable.6
Although
the
shuttle
disulfide
is
closer to the active site in one of the Ero1p crystal
forms than in the other, it is still not in direct con-
tact and is separated from the active site by an
intervening Tyr side chain (Tyr 191). The low sol-
vent accessibility of the active-site disulfide in Ero1p
is not altered by in silico removal of the shuttle di-
sulfide loop (data not shown), suggesting that it is
buried by other parts of the structure, occluding it
from solvent as well as from the shuttle disulfide.
Conformational or dynamic changes, propagating
from reduction of regulatory disulfides, may facili-
tate redox communication between the solvent acces-
sible shuttle disulfide and the buried active-site di-
sulfide,
forming
a
complete
redox
path
from
substrate to FAD.
In
conclusion,
we
determined
that
the
C143AC166 disulfide is the first regulatory disulfide
to be reduced during Ero1p activation and that the
facile activation of the C150A/C295A mutant can be
explained by its increased susceptibility to reduction
of C143AC166 (Fig. 8). The presence or absence of
the C150AC295 disulfide 35 A˚ away from the active
site thus affects the fate of the key C143AC166 di-
sulfide 25 A˚ from the active site, and elimination of
the
C143AC166
disulfide
allows
conformational
changes that may finally propagate to the active site
itself, with a consequent increase in thiol oxidase
activity.
Materials and Methods
Enzyme and substrate preparation
Ero1p, Ero1p C143A, Ero1p C166A, Ero1p C150A/
C295A,
and
Ero1p
C143A/C166A,
all
spanning
Figure 8. Summary of roles for regulatory disulfides distant
from the Ero1p active site. Reduction of the C150AC295
disulfide is not an early step in Ero1p activation. Instead,
the presence of this disulfide appears to protect the
neighboring C143AC166 disulfide from reduction.
Reduction of C143AC166 may be a key event in Ero1p
activation, allowing the polypeptide chain in the vicinity of
C166 in particular to sample new conformations (the arrow
indicates schematically putative motion in the direction of
the active site). Though C143AC166 is nearly 25 A˚ from the
active-site (C352AC355) disulfide in the ground-state Ero1p
structure, a liberated C166 would be present on a segment
of polypeptide that may be loosely tethered enough to
bridge this distance.
Heldman et al.
PROTEIN SCIENCE VOL 19:1863—1876
1873
residues amino acid 56–424 of the yeast protein,
were expressed in the Origami B(DE3) plysS E. coli
strain (Novagen) downstream of glutathione S-trans-
ferase (GST) and an internal His6 tag using a modi-
fied
pGEX-4T1
plasmid
(Amersham).6
Wild-type
Ero1p spanning residues 10–424 was produced from
a pGEX-4T1 vector without the additional His6 tag.
Bacteria were grown at 37C to an optical density of
0.6 at 600 nm, at which point isopropyl-1-thio-b-D-
galactopyranoside was added to a final concentration
of 0.5 mM to induce protein expression. The growth
temperature was shifted to 25C, and cells were har-
vested 12–16 h later. After cell lysis, proteins were
purified using Ni-NTA agarose (for constructs con-
taining His6 tags), cleaved with thrombin to remove
the GST, and re-purified over Ni-NTA. The longer
Ero1p construct, lacking the His6 tag, was purified
using glutathione sepharose beads, and thrombin
cleavage was performed on the column to release
Ero1p. The enzymes were then concentrated and
run on a HiLoad 16/60 Superdex 75 prep grade size
exclusion column monitored at 280 and 450 nm. The
peaks corresponding to monomeric protein were col-
lected. Enzymes used for crystallization were dia-
lyzed against 10 mM Tris, 25 mM NaCl, pH 8 and
concentrated to 400 lM (18 mg/mL).
Trx was expressed, purified, and reduced as pre-
viously described,6 except that Triton X-100 was not
used. For experiments in which gels were stained
with Invision His6-tag stain, Pdi1p was produced in
BL21(DE3) plysS E. coli cells as a fusion protein
with GST using the pGEX-4T1 plasmid, purified
over
glutathione-sepharose
beads,
cleaved
with
thrombin, and re-applied to glutathione sepharose to
remove the GST. For other experiments, Pdi1p was
produced
with
a
carboxy-terminal
His6-tag
as
described.14
For oxidation assays, Trx was reduced with 100
mM DTT for 1 h and then desalted using a PD-10
column (GE Healthcare) equilibrated in 50 mM
phosphate buffer, pH 7.5, 65 mM NaCl, 0.5 mM
EDTA. Pdi1p was reduced by incubating with 10
mM GSH from a stock titrated to pH 7.0. Proteins
were then desalted on a PD-10 column equilibrated
in 50 mM phosphate buffer, pH 7.5, 300 mM NaCl,
0.5 mM EDTA. The concentration of reduced protein
thiols was determined using Ellman’s assay.15
Crystallization and structure determination
Crystals of Ero1p C150A/C295A were grown by
hanging drop vapor diffusion at 20C in 100 mM cac-
odylic acid pH 6–6.5, 9–15 mM cadmium sulfate, 2%
methanol, 2% ethanol, and 0.8–1.6M sodium acetate.
These crystals were of space group C2221. Crystals
of Ero1p C143A/C166A were grown by hanging drop
vapor diffusion at 20C in 100 mM cacodylic acid pH
6–6.5, 12–14 mM cadmium sulfate, 2% methanol,
2% ethanol, and 1.6M sodium acetate. These crystals
were of space group P62. Before flash freezing, crys-
tals were soaked in mother liquor with 15% ethylene
glycol for a few minutes and then transferred to a
1:1 mixture of mineral oil and paratone oil (Exxon).
Diffraction data for Ero1p C150A/C295A were col-
lected at the European Synchrotron Radiation Facil-
ity beamline ID14-3. Data for Ero1p C143A/C166A
were collected using a RU-H3R generator (Rigaku)
equipped with an Raxis IVþþ image plate system
and
osmic
mirrors.
Data
were
processed
using
HKL2000.16 Phases were calculated for the C150A/
C295A mutant using the wild-type Ero1p structure
after removal of residues 146–166 and 291–302,
spanning the mutated cysteines. Phases were calcu-
lated for the C143A/C166A mutant after removal of
residues 91–121 and 136–175. Rigid body refinement
was performed before generating electron density
maps. The Ero1p C150A/C295A and Ero1p C143A/
C166A models was rebuilt in O17 and Coot,18 respec-
tively, and refined using CNS.19 The quality of the
final models were verified using MolProbity.20
Stopped-flow fluorescence
Studies were conducted at 25C on an Applied Pho-
tophysics stopped-flow apparatus fitted with an an-
aerobic adaptor. Exogenous FAD (100 lM) rather
than
oxygen
was
used
as
a
terminal
electron
acceptor, since changes in fluorescence can be moni-
tored as FAD is reduced to FADH2. Excitation was
at a wavelength of 450 nm, and an emission cut-off
filter of 495 nm was used. Enzyme and substrate
preparations were made anaerobic by N2 bubbling
and pipetting in N2 atmosphere in a glove box. To
insure anaerobic conditions, solutions were supple-
mented with 0.1% w/v glucose and trace amounts of
glucose oxidase and catalase. Samples were trans-
ferred
from the
anaerobic chamber
in gas-tight
syringes (Hamilton). Equal volumes of enzyme and
Trxred (prepared as described above) in 50 mM phos-
phate buffer, pH 7.5, 65 mM NaCl, and 1 mM EDTA
were injected into the mixing/detection chamber to
achieve final concentrations of 1 lM enzyme and 50
lM substrate.
Gel-based oxidation assays
Oxidation reactions used to analyze Ero1p electro-
phoretic mobility were performed at an enzyme
(wild-type or mutant) concentration of 2 lM. Trxred
was present at a final concentration of 100 lM (fully
reduced protein), and Pdi1pred at a concentration of
300 lM thiols (75 lM protein). Samples from the
reactions were taken at different time points and
quenched by adding 1:4 v/v sample buffer (125 mM
Tris, pH 6.8, 5% SDS, 50% glycerol, 0.1% bromphe-
nol blue) containing 5 mM AMS (Molecular Probes),
100 mM NEM, or 1 mM PEG-maleimide of 5 kD
(Nektar Therapeutics). These samples were then run
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PROTEINSCIENCE.ORG
Activation of Sulfhydryl Oxidase Ero1p
on 15% denaturing polyacrylamide gels [Figs. 4 and
5(A)], or 12% gels for the kinetics of Pdi1pred oxida-
tion [Fig. 6(C)], which were then stained with Coo-
massie.
Gels
used
to
visualize
Ero1p
or
Ero1p
C150A/C295A
in
reactions
with
Pdi1pred
were
stained with InVisionTM His-tag In-gel Stain (Invi-
trogen) and scanned using a FLA-5100 fluorescent
image analyzer (FUJI) with a 532 nm LPG filter.
Reduced samples [Fig. 4(B)] were run with an excess
of DTT.
Rates of oxidation of Trxred by various mutants
of Ero1p [Fig. 7(D)] were quantified using 0.5 lM
enzyme and 100 lM Trxred. At each time point, 10 lL
were removed and mixed with 10 lL 5 mM PEG-mal-
eimide 5 kDa in 2% SDS, 50 mM Tris, pH 6.8, 0.1%
bromphenol blue, 20% glycerol. Samples were applied
to a 15% denaturing polyacrylamide gel, and Trxred
and Trxox were visualized after separation with Coo-
massie stain. Band intensities were determined using
the ImageQuant 5.0 program.
In-gel digestion
Protein bands were excised from SDS gels that had
been stained with Coomassie and destained using
multiple washings with 40% methanol and 10% ace-
tic acid. The protein bands were either reduced,
alkylated, and in-gel digested as described,21 or else
digested directly without reduction and alkylation.
Digestions were performed in 50 mM ammonium bi-
carbonate at 37C using various combinations of the
following sequencing grade proteases (Roche Diag-
nostics): bovine trypsin, bovine chymotrypsin, and
Pseudomonas asp-N, all at a concentration of 12.5
ng/lL. Peptide mixtures were extracted from the
gels with 80% CH3CN, 1% CF3COOH, and the or-
ganic solvent was evaporated in a vacuum centri-
fuge. The resulting peptide mixtures were reconsti-
tuted in 80% formic acid and immediately diluted
1:10 with Milli-Q water before mass spectrometry
analysis.
Mass spectrometry
Samples were analyzed in an LTQ Orbitrap (Thermo
Fisher Scientific) operated in the positive ion mode
and equipped with a nanoelectrospray ion source.
Peptide mixtures were separated by online reversed-
phase nanoscale capillary LC and analyzed by tan-
dem mass spectrometry (LC-MS/MS). For the LC-MS/
MS, samples were injected onto a 15 cm reversed
phase spraying fused-silica capillary column (inner
diameter 75 lm) made in-house and packed with 3
lm ReproSil-Pur C18AQ media (Dr. Maisch GmbH,
Ammerbuch-Entringen, Germany) using an UltiMate
3000 Capillary/Nano LC System (LC Packings, Dio-
nex). The LC setup was connected to the Orbitrap.
The flow rate through the column was 250 nL/min,
and the injection volume was 5 lL. Peptides were
separated with a 50 min gradient from 5 to 65% ace-
tonitrile (buffer A: 5% acetonitrile, 0.1% formic acid,
0.005% TFA; buffer B: 90% acetonitrile, 0.2% formic
acid, 0.005% TFA). The voltage applied to the union
in order to produce an electrospray was 1.2 kV. The
mass spectrometer was operated in the data-depend-
ent mode. Survey MS scans were acquired in the
Orbitrap with the resolution set to a value of 60,000.
Up to the six most intense ions per scan were frag-
mented and analyzed in the linear trap. For the anal-
ysis of peptides, survey scans were recorded in the
FT-mode followed by data-dependent collision-induced
dissociation (CID) of the six most intense ions in the
linear ion trap (LTQ). Raw spectra were processed
using open-source software DTASuperCharge (http://
msquant.sourceforge.net). The data were searched
with MASCOT (Matrix Science, London, UK) against
a Swiss-prot database with manually incorporated
Ero1p
or
mutant
protein
(Ero1p
C150A/C295A).
Search parameters included variable modifications of
57.02146
Da
(carboxyamidomethylation)
on
Cys,
510.04028 Da (AMS; hydrolyzed) on Cys, 15.99491
Da (oxidation) in Met and 0.984016 Da (deamidation)
on Asn and Gln. The search parameters were as fol-
lows: maximum two missed cleavages, initial precur-
sor ion mass tolerance 10 ppm, fragment ion mass
tolerance 0.6 Da. The identity of the peptides were
concluded from the detected CID products by Mascot
and
Sequest software and
confirmed
by manual
inspection of the fragmentation series. The program
MassMatrix was used for disulfide identification.22
Acknowledgments
The authors thank members of the Weizmann Insti-
tute Biological Mass Spectrometry Unit and Sarah J.
Weisberg for assistance with mass spectrometry anal-
ysis. Moran Bentzur and Gideon Schreiber assisted in
the stopped-flow studies. Coordinates and structure
factors for Ero1p C150A/C295A and Ero1p C143A/
C166A have been deposited in the Protein Data Bank
with accession codes 3M31 and 3NVJ.
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PROTEINSCIENCE.ORG
Activation of Sulfhydryl Oxidase Ero1p
|
3M32
|
Structural Insight into Methyl-Coenzyme M Reductase Chemistry using Coenzyme B Analogues
|
Structural Insight into Methyl-Coenzyme M Reductase Chemistry
using Coenzyme B Analogues,†,‡
Peder E. Cedervall§,#, Mishtu Dey||,#,⊥, Arwen R. Pearson§,⊥, Stephen W. Ragsdale||, and
Carrie M. Wilmot*,||,§
§ Department of Biochemistry, Molecular Biology and Biophysics, University of Minnesota,
Minneapolis, Minnesota 55455
|| Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109
Abstract
Methyl-coenzyme M reductase (MCR) catalyzes the final and rate-limiting step in methane
biogenesis; the reduction of methyl-coenzyme M (methyl-SCoM) by coenzyme B (CoBSH) to
methane and a heterodisulfide (CoBS-SCoM). Crystallographic studies show that the active site is
deeply buried within the enzyme, and contains a highly reduced nickel-tetrapyrrole, coenzyme
F430. Methyl-SCoM must enter the active site prior to CoBSH, as species derived from analogues
of methyl-SCoM are always observed bound to the F430 nickel in the deepest part of the 30 Å long
substrate channel that leads from the protein surface to the active site. The seven-carbon
mercaptoalkanoyl chain of CoBSH binds within a 16 Å predominantly hydrophobic part of the
channel close to F430, with the CoBSH thiolate lying closest to the nickel at a distance of 8.8 Å. It
has previously been suggested that binding of CoBSH initiates catalysis by inducing a
conformational change that moves methyl-SCoM closer to the nickel promoting cleavage of the C-
S bond of methyl-SCoM. In order to better understand the structural role of CoBSH early in the
MCR mechanism, we have determined crystal structures of MCR in complex with four different
CoBSH analogues; pentanoyl-, hexanoyl-, octanoyl- and nonanoyl- derivatives of CoBSH
(CoB5SH, CoB6SH, CoB8SH and CoB9SH respectively). The data presented here reveal that the
shorter CoB5SH mercaptoalkanoyl chain overlays with that of CoBSH, but terminates two units
short of the CoBSH thiolate position. In contrast, the mercaptoalkanoyl chain of CoB6SH adopts a
different conformation, such that its thiolate is coincident with the position of the CoBSH thiolate.
†This work was supported by a Department of Energy grant DE-FG02-08ER15931 to S.W.R and supplement to C.M.W. and a
Minnesota Partnership for Biotechnology and Medical Genomics grant SPAP-05-0013-P-FY06.
‡Co-ordinates and structure factors have been deposited in the Protein Data Bank as entries 3m1v (MCRHSCoM); 3m2r
(MCRCoB5SH); 3m2u (MCRCoB6SH); 3m2v (MCRCoB8SH); 3m30 (MCRCoB9SH).
*Address correspondence to: Carrie M. Wilmot, Tel: (612) 624-2406, wilmo004@umn.edu.
⊥Current addresses: Mishtu Dey, Department of Chemistry, Massachusetts Institute of Technology, 77 Massachusetts Ave.,
Cambridge, Massachusetts 02139; Arwen Pearson, Astbury Centre for Structural Molecular Biology, Institute for Molecular and
Cellular Biology, Astbury Building. Leeds, LS2 9JT, U.K.
#These authors contributed equally to this work.
Supporting Informational material is available free of charge via the Internet at http://pubs.acs.org and contains the following:
MATERIAL AND METHODS for single crystal UV-visible microspectrophotometry, X-ray photoreduction experiment and X-ray
crystallography of the MCR-heterodisulfide product complex (MCRCoBSH + methyl-SCoM); RESULTS AND DISCUSSION for
redox changes and MCRCoBSH + methyl-SCoM crystal structure; Table S1, X-ray Data Collection, Processing and Refinement
Statistics for MCRCoBSH + methyl-SCoM; Figure S1, use of Fo-Fc electron density in modelling MCRCoB8SH; Figure S2,
illustration of electron density quality of MCRCoB5SH; Figure S3, solution and single crystal UV-visible spectra; Figure S4,
modeling of CoB6SH and CoBSH into the electron density of MCRCoB6SH; Figure S5, partially occupied HSCoM in MCRCoB6SH;
Figure S6, the active site and substrate channel of MCRCoBSH + methyl-SCoM; Figure S7, alternative conformation of Valα482 in
MCRHSCoM; Figure S8, the two conformations of the gly-rich loop in MCRHSCoM; Figure S9, propagation of conformational
changes in MCRHSCoM; Figure S10, the two conformations of CoB9SH in MCRCoB9SH; Figure S11, EPR spectra of MCRred1
sample; Scheme S1, scheme of the characterized forms of MCR.
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 September 7.
Published in final edited form as:
Biochemistry. 2010 September 7; 49(35): 7683–7693. doi:10.1021/bi100458d.
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This is consistent with the observation that CoB6SH is a slow substrate. A labile water in the
substrate channel was found to be a sensitive indicator for the presence of CoBSH and HSCoM.
The longer CoB8SH and CoB9SH analogues can be accommodated in the active site through
exclusion of this water. These analogues react with Ni(III)-methyl; a proposed MCR catalytic
intermediate of methanogenesis. The CoB8SH thiolate is 2.6 Å closer to the nickel than that of
CoBSH, but the additional carbon of CoB9SH only decreases the nickel thiolate distance a further
0.3 Å. Although the analogues did not induce any structural changes in the substrate channel, the
thiolates appeared to preferentially bind at two distinct positions in the channel; one being the
previously observed CoBSH thiolate position, and the other being at a hydrophobic annulus of
residues that lines the channel proximal to the nickel.
INTRODUCTION
Methanogenic archaea are organisms that under strict anaerobic conditions derive energy by
reducing compounds such as carbon dioxide, methylamine, acetate and methanol, to
methane (1, 2). The global production of methane by these organisms is estimated at one
billion tons annually. Microbially produced methane is not only a potential source of
renewable energy but also a potent greenhouse gas, and as such study of this process has
environmental ramifications. In these microorganisms, methyl-coenzyme M reductase
(MCR)1 is the enzyme that catalyzes the final step in methanogenesis, in which the
substrates methyl-coenzyme M (methyl-SCoM, 2-(methylthio)ethanesulfonate) and
coenzyme B (CoBSH, N-7-mercaptoheptanoylthreonine phosphate) are converted to
methane and a heterodisulfide (CoBS-SCoM) (Scheme 1) (3).
MCR is a 300 kDa protein with six subunits arranged in a α2β2γ2 oligomer (4). The known
crystal structures show that MCR has two active sites approximately 50 Å apart that are
deeply buried within the enzyme (5). The active site pocket is comprised of residues from
subunits α, α′, β and γ, with a 30 Å long substrate channel leading to the enzyme surface
(Figure 1). At the heart of the active site pocket is coenzyme F430, which is a highly reduced
nickel-containing tetrapyrrole (6–8). Currently sixteen distinct enzymatic and complexed
states of MCR have been spectroscopically characterized (Supporting Information, Scheme
S1) (9–31). In the resting active state of the enzyme, denoted MCRred1, the redox-active
nickel of F430 is present in the Ni(I) state (9, 16, 32). MCR is extremely oxygen sensitive
and upon oxygen exposure the enzyme enters an inactive Ni(II) state, denoted MCRred1-silent
(6). In this state it cannot be converted back to the active Ni(I) form by any known reducing
agent making this a challenging system to study. Additional complications involve the tight
association of coenzymes to purified MCR that are not easily displaced as demonstrated by
X-ray crystallographic and kinetic studies (5, 33–35).
Despite the fact that MCR has been studied for decades, no true catalytic intermediate has
been observed, and the actual mechanism remains elusive. Currently three general
mechanistic schemes for the enzymatic reaction have been proposed, each of which posit
different chemistry to initiate catalysis. Mechanism I involves Ni(I) acting as a nucleophile
in an SN2-type reaction that generates Ni(III)-methyl as an intermediate (Scheme 2A) (35–
38). Mechanism II starts with methyl-SCoM undergoing homolytic cleavage at the Ni(I) to
generate a methyl radical and a Ni(II)-SCoM species (Scheme 2B) (39–41). A more recently
proposed mechanism III suggests protonation of coenzyme F430 promotes reductive
cleavage of the methyl-SCoM thioether bond (42).
1Abbreviations: MCR, Methyl-coenzyme M reductase; methyl-SCoM, methyl-coenzyme M; CoBSH, coenzyme B; HSCoM,
coenzyme M; CoBS-SCoM, heterodisulfide of coenzyme B and coenzyme M; APS, Advanced Photon Source; ASU, asymmetric unit;
BPS, bromopropanesulfonate.
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Due to the stringent requirement to exclude O2, the available MCR crystal structures are all
in the inactive Ni(II) state. All but one contain CoBSH and HSCoM (demethylated methyl-
SCoM, an inhibitor and substrate analogue) in the active site (PDB codes 1hbn, 1hbo, 1hbu,
1e6y, 1e6v) (5, 33, 34). Another crystal structure has bound heterodisulfide product, CoBS-
SCoM (MCRsilent, PDB code 1hbm, Scheme 1 and Supporting Information, Scheme S1) (5,
33). All these structures reveal that both substrates access the active site through the same
channel (Figure 1). The binding site of HSCoM (and presumably methyl-SCoM) is more
deeply buried within the enzyme, and so it must enter prior to CoBSH for productive
chemistry to occur. As binding of CoBSH in the absence of co-substrate would be
inhibitory, it was suggested that a conformational change upon methyl-SCoM binding might
lower the Kd for CoBSH, and thus promote an ordered mechanism. Furthermore it has been
suggested that CoBSH binding induces a conformational change that brings the methyl-
SCoM substrate into closer proximity to the nickel, and this promotes C-S bond cleavage.
To investigate the proposed structural role of CoBSH in initiating catalysis, we have solved
the X-ray crystal structures of MCR in complex with four different CoBSH analogues.
CoBSH has a heptanoyl moiety linked to the thiol group, and the analogues are pentanoyl-,
hexanoyl-, octanoyl- or nonanoyl-containing derivatives of CoBSH (CoB5SH, CoB6SH,
CoB8SH and CoB9SH respectively; Figure 2) (3, 35, 43–47). In addition, we present a
structure in which the substrate channel predominantly lacks either CoBSH or
heterodisulfide product.
MATERIALS AND METHODS
Materials
The organism Methanothemobacter marburgensis (catalog OCM82) was obtained from the
Oregon Collection of Methanogens (Portland, OR). All buffers and media reagents were
obtained from Sigma-Aldrich (St. Louis, MO). The gases N2 (99.98%), H2/CO2 (80%/20%),
and ultra high purity H2 (99.999%) were obtained from Cryogenic Gases (Grand Rapids,
MI). Ti(III) citrate solutions were prepared from a stock solution of 246 mM Ti(III) citrate,
which was synthesized by adding sodium citrate to Ti(III) trichloride (30 wt % solution in 2
N hydrochloric acid) (Acros Organics, Morris Plains, NJ) under anaerobic conditions and
adjusting pH to 7.0 with sodium bicarbonate (48). The concentration of Ti(III) citrate was
determined by titrating against a solution of methyl viologen.
Synthesis of methyl-SCoM, CoB5SH, CoB6SH, CoBSH, CoB8SH, and CoB9SH
Methyl-SCoM was prepared from HSCoM and methyl iodide (49). The homodisulfides,
CoB5S-SCoB5, CoB6S-SCoB6, CoBS-SCoB, CoB8S-SCoB8, CoB9S-SCoB9 were prepared
as described from 5-bromovaleric acid, 6-bromohexanoic acid (Sigma-Aldrich, St. Louis,
MO), 7-bromoheptanoic acid (Karl Industries, Aurora, OH), 8-bromooctanoic acid, and 9-
bromononanoic acid (Matrix Scientific, Columbia, SC), respectively (43, 46). The free thiol
forms of CoB5SH, CoB6SH, CoBSH, CoB8SH, and CoB9SH were generated by the
reduction of the homodisulfides as previously described (45). The purity of the CoBSH
analogues was determined by 1H NMR spectroscopy. All compounds synthesized were
stored in a Vacuum Atmospheres chamber maintained at an oxygen level below 1 ppm, as
monitored continually with an oxygen analyzer (model 317, Teledyne Analytical
Instruments, City of Industry, CA) until use.
M. marburgensis Growth and MCRred1 Purification
Buffer preparations and all manipulations were performed under strict anaerobic conditions
in a Vacuum Atmospheres chamber maintained at an oxygen level below 1 ppm, as
monitored continually with an oxygen analyzer (model 317, Teledyne Analytical
Instruments, City of Industry, CA). MCRred1 was isolated from M. marburgensis cultured on
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H2/CO2 (80%/20%) at 65 °C in a 14-L fermentor (New Brunswick Scientific Co., Inc. New
Brunswick, NJ). Culture media were prepared as previously described (20, 24). MCRred1
was generated in vivo and purified as described previously (20). The purification procedure
routinely generates 65–75% MCRred1 as determined by UV-visible and EPR spectroscopy.
Spectroscopy of MCR
UV-visible spectra of the Ni(I)-containing MCRred1 were recorded in the anaerobic chamber
using a spectrophotometer (Model USB4000-UV-VIS, Ocean Optics, Dunedin, FL). EPR
spectra were recorded on a Bruker EMX spectrometer (Bruker Biospin Corp., Billerica,
MA), equipped with an Oxford ITC4 temperature controller, a Hewlett-Packard Model 5340
automatic frequency counter and Bruker gaussmeter. The EPR spectroscopic parameters
included: temperature, 70 K; microwave power, 10 mW; microwave frequency, 9.43 GHz;
receiver gain, 2 × 104; modulation amplitude, 10.0 G; modulation frequency, 100 kHz.
Double integrations of the EPR spectra were performed and referenced to a 1 mM copper
perchlorate standard. The NMR data were acquired at 298 K on a Bruker Avance DRX 500
MHz instrument equipped with a TXI cryoprobe.
Preparation of MCRred1 for Crystallization
All crystallization experiments were performed in the anaerobic chamber in which MCR
was purified unless otherwise noted. MCRred1 was prepared in 50 mM Tris, pH 7.6 and
excess HSCoM was removed by buffer exchange using an Amicon Ultra centrifuge filter
with a 50 kDa cut-off membrane (Millipore). Typically, 2–3 ml of MCRred1 was exchanged
with 10–15 ml of 50 mM Tris, pH 7.6. The enzyme was concentrated to 500–600 μl, and
this process was repeated three times. The fraction of MCRred1 in the purified MCR sample
was calculated from the UV-visible spectrum using extinction coefficients of 27.0
mM−1cm−1 at 385 nm for Ni(I)-MCRred1, and 9.15 mM−1cm−1 at 420 nm for Ni(II)-
MCRred1-silent (20). The amount of MCRred1 in samples used for crystallization was
determined to be ~80% and the concentration of total enzyme used was in the range of about
120–150 μM (~32–40 mg/ml). All crystallization experiments were performed anaerobically
by incubating 2.0 μl of enzyme solution in 50 mM Tris, pH 7.5 and 2.0 μl of reservoir
solution (100 mM Hepes-Na, pH 7.3/7.5/8.0; 150 mM magnesium acetate (Mg(CH3COO)2),
and 20/22% (w/v) PEG 400) in a sitting drop over 1 ml reservoir solution at 9 ºC. Triangular
and rectangular prismatic crystals with a bright yellowish-green color confirmed the
presence of nickel coenzyme F430. The crystals grew to a size of approximately 100–200 μm
in 4–5 days. CoBSH-depleted crystals were obtained by incubating 2 μl of a reaction
mixture containing 139 μM MCRred1 and 13 mM HSCoM with 2 μl of reservoir solution
(100 mM Hepes-Na pH 7.5, 150 mM magnesium acetate (Mg(CH3COO)2), 22% PEG 400).
Crystals of MCR complexed with the CoBSH analogues were grown by co-crystallization.
The CoB5SH co-crystals were obtained by incubating 2μl enzyme solution containing 124
μM MCRred1, 10 mM methyl-SCoM and 1 mM CoB5SH with 2μl of reservoir solution (100
mM Hepes-Na, pH 7.5, 150 mM (Mg(CH3COO)2), 22 % PEG 400). The crystals with
bound CoB6SH and CoB9SH were obtained by co-crystallization of 1 mM of analogue with
142 μM MCRred1 and equilibrated with 2μl of reservoir solution (100 mM Hepes-Na, pH
7.5, 150 mM Mg(CH3COO)2, 20 % PEG 400 for CoB6SH and 100 mM Hepes-Na, pH 7.3,
150 mM Mg(CH3COO)2), 22 % PEG 400 for CoB9SH). Crystals were cryoprotected in
reservoir solution containing 25 % (v/v) PEG 400 by soaking for 2–5 minutes before
cryocooling in liquid nitrogen in the anaerobic chamber. Crystals of CoB8SH bound to MCR
were obtained by incubating 2 μl of a mixture of 119 μM MCRred1 and 1 mM CoB8SH with
2 μl of reservoir solution (100 mM Hepes-Na, pH 7.3, 150 mM Mg(CH3COO)2), 20 % PEG
400). Before cryoprotection, the crystals were soaked for 5–10 minutes in a 100 mM
solution of methyl iodide2. The methyl iodide solution used for soaking was prepared by
adding a concentrated stock of methanolic solution of methyl iodide to the reservoir
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solution. Soaked crystals were quickly cryoprotected as described above and cryocooled in
the anaerobic chamber.
X-ray Diffraction Data Collection, Processing and Refinement
X-ray diffraction data were collected at 100 K on a ADSC Quantum-315 detector at the APS
Beamline 14-BM-C (BioCARS). The wavelength of X-rays was 0.979 Å. Data were
processed using HKL2000 (50). As in the previous X-ray crystallographic studies, the
crystals belong to the monoclinic space group P21 (a = 82 Å, b = 118 Å, c = 122 Å, β = 92°),
with one MCR molecule (two active sites) per asymmetric unit (5, 33). For refinement,
REFMAC in the Collaborative Computational Project Number 4 (CCP4) program suite was
used (51). A random sample of 5 % of the data across all resolution shells was chosen to
check refinement progress through calculation of an Rfree. The same reflections were used to
calculate Rfree for all structures, thus preventing bias due to high structural identity. The
remaining reflections were used in refinement (Rwork). Model building was done using the
Crystallographic Object-Oriented Toolkit (COOT) (52). The diffraction data and their
models are designated as MCRCoBXSH, where X is the number of carbons in the alkanoyl
portion of the analogue. Library files in CCP4 for F430 and CoBSH were incorrect, and these
were modified in Monomer Library Sketcher in the CCP4 program suite by comparison with
schematic drawings from Grabarse et al. (33). Co-ordinate and CCP4 library files for the
different CoBSH analogues were created in Monomer Library Sketcher. The general model
building and refinement strategy for all structures was as follows. It was clear from the
electron density in the substrate channel and at the active site that mixtures of species were
present in all datasets. These could be visualized with Fo-Fc and Fo-Fo difference electron
density maps (Supporting Information, Figure S1). The known positions of CoBSH and
HSCoM from the published Ni(II)-MCR crystal structures (PDB codes 1hbn, 1hbo, 1hbu
(33)) were used as guides to determine which species could be present in each dataset, and
these were then simultaneously modeled into the electron density. By alteration of their
relative occupancies (in 10% increments) followed by refinement, the ratio of occupancy
between different species was determined using the assumption that the average B-factors
for all molecular species bound should be similar to that of F430 and adjacent well-ordered
protein atoms within the active site and substrate channel. The combinations of modeled
ligands were constantly reassessed throughout refinement based on the remaining difference
electron density. This included test refinements of different ligand combinations during the
latter stages, thus using the optimized phases to check whether a different combination of
ligands could also explain the electron density. Sensible chemical structures and
interactions, along with keeping the combined occupancies of sterically mutually exclusive
species ≤ 100%, were maintained throughout refinement. The model was finally accepted
when the difference electron density map was minimal and the B-factors for the models
converged.
In practice the first structure refined was that of MCRCoB5SH. Initial phases were generated
by difference Fourier using a previously determined crystal structure (PDB code 1mro (5))
but with all non-bonded molecules, including water, removed from the model except F430.
Initial rigid body refinement followed by restrained refinement of MCRCoB5SH reduced the
Rwork to 26.5%. After model building and subsequent rounds of restrained refinement the
Rwork was 14.3% (Rfree 16.6%). Of the five structures only the CoB5SH analogue is
completely coincident with CoBSH, and so particular care had to be used in teasing apart the
ratios of the two species in modeling the MCRCoB5SH electron density. This was done by
2Methyl iodide was added with the intention of creating a Ni(III)-methyl species, which CoB8SH stabilizes. This was not achieved,
but the diffraction quality of this crystal was significantly better than crystals co-crystallized with CoB8SH alone, and so has been
included in this study.
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initially refining 100% CoBSH in the substrate channel. Positive Fo-Fc difference density
located at the carbon where the shorter CoB5SH thiol might be expected to be indicated the
presence of a more electron-rich species than carbon, which is consistent with the presence
of the CoB5SH sulfur. The refinement converged at a model containing a 50:50 mix of
CoBSH and the analogue. However, positive Fo-Fc difference density was still present at the
position of the CoBSH thiol, therefore a water molecule was added to the CoB5SH model at
50% occupancy and upon refinement this accounted for the electron density. An illustration
of the electron density quality from this structure is shown in Supporting Information,
Figure S2. An HSCoM-, CoBSH- and CoBSH analogue-free version of the refined
MCRCoB5SH structure was used as the starting model to generate initial phases for the four
other structures. After the initial round of restrained refinement the Rwork for these structures
were reduced to 14.5–15.6 %.
RESULTS AND DISCUSSION
Crystal Structures of MCR
Five crystal structures were determined, four of which are in complex with CoBSH
analogues differing in the number of carbons atoms in the alkanoyl portion of the molecule.
CoBSH is an N-7-mercaptoheptanoyl-containing molecule, whereas the four CoBSH
analogues contain N-5-mercaptopentanoyl-, N-6-mercaptohexanoyl-, N-8-mercaptooctanoyl-
or N-9-mercaptononanoyl-moieties (Figure 2). The corresponding crystal structures are
designated as MCRCoBXSH, where X is the number of carbons in the alkanoyl portion of the
analogue. The other crystal structure is of MCRred1c-silent (MCR in the Ni(II) state in
complex with HSCoM, designated here as MCRHSCoM) that is CoBSH-depleted. The
datasets have resolutions in the range from 1.35 – 1.8 Å. Although the crystallizations were
set up with the MCR solution predominantly in the Ni(I)-MCRred1 state, by the time X-ray
diffraction data were collected they had been oxidized to the Ni(II)-MCRred1-silent state
(Supporting Information). Following data collection there was no evidence for
photoreduction of the Ni(II) back to Ni(I) in any of the crystals, as assessed by single crystal
UV-visible microspectrophotometry (Supporting Information and Figure S3). Attempts to
photoreduce the crystals using different wavelengths and temperatures were unsuccessful
(Supporting Information).
Overall, the resulting structures are very similar to each other and to the previously
published structures of MCR, with differences mainly localized to the active site and
substrate channel. The two active sites in the ASU were refined independently. Unless
otherwise stated there was no difference between them. All five datasets contain a mixture
of species bound to the enzyme. There is always a background of CoBSH and HSCoM,
which co-purify with MCR and cannot be fully removed by extensive buffer exchange or by
the addition of a CoBSH analogue. HSCoM is added during purification of MCR, as it
stabilizes the resting active Ni(I) state (unpublished data), and this leads to HSCoM
occupancies between 50–100% amongst the structures (Table 1). In contrast CoBSH, which
is not added during purification, has occupancies ranging from 30–50%. As these
confounding species have all been described at high occupancy in other crystallographic
studies, the structural data of interest could be isolated (5, 33). In each case, the additional
electron density could be explained by inclusion of the appropriate CoBXSH model used in
that experiment at 50% or higher occupancy. The resulting models, along with 2Fo-Fc
electron density, are shown in Figure 3. The Rwork for the final structures range from 13.0 to
15.0 % (Rfree 15.5 to 19.5 %). The X-ray data collection, processing, refinement and model
building statistics are given in Table 1.
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Analogues shorter than CoBSH; CoB5SH and CoB6SH
CoB5SH is two methylene groups shorter than CoBSH, the MCR substrate. The
MCRCoB5SH structure is to 1.35 Å resolution. As expected the pentanoyl chain follows the
path of the CoBSH heptanoyl carbons down the substrate channel, and thus its thiol is
positioned in the same place as the second carbon preceding the CoBSH thiol (Figure 3A
and 4). There are no published MCR kinetic studies using CoB5SH, but as it binds in the
substrate channel, it is likely to be an inhibitor.
CoB6SH is one methylene shorter than CoBSH, and is a slow substrate of MCR. In this case
the 1.4 Å resolution electron density of MCRCoB6SH indicates that the analogue
unexpectedly binds in the substrate channel such that its thiol is virtually in the same
position as the thiol of CoBSH (Figure 3B and 4). The hexanoyl chain is oriented so that it
takes a shorter route down the substrate channel between carbons 2 and 5 (with the carbonyl
carbon labeled as carbon 1) than CoBSH (Figure 4, Supporting Information, Figure S4).
This short-cut is not seen in any of the other CoBXSH complex crystal structures, but
presumably arises because this CoB6SH binding conformer is energetically more favorable,
although it is not clear from the structure why this might be the case. CoB6SH binds very
tightly to MCR, with an apparent Ki value of 0.1 μM (3).
Water structure in the absence of HSCoM
The electron density for the MCRCoB6SH crystal structure only supported the modeling of 50
% bound HSCoM. In the fraction of MCR molecules where HSCoM is absent, the HSCoM
binding site is occupied by a network of four water molecules (Supporting Information,
Figure S5). Two waters are positioned close to the absent sulfonate oxygen positions of
HSCoM. Based on the presence of positive difference electron density, a third water was
modeled ligated to the Ni, and refined to a distance of ~2 Å (2.0 Å and 2.1 Å in the two
active sites of the ASU) with no distance restraint imposed between the Ni and water. This
water is in a similar position as the Ni coordinating sulfonate oxygen of the heterodisulfide
product in MCRsilent (Supporting Information, Figure S6 and PDB codes 3m32, 1hbm) (5,
33). The fourth water was in the vicinity of the expected position of a bridging water (W1)
seen in other structures (Figure 1, 3A and 3C).
Water structure in the absence of CoBSH
The 1.45 Å resolution electron density obtained for MCRHSCoM indicates that the substrate
channel contains only 30 % CoBSH. Nine ordered waters (W1–W9), along with an acetate
ion from the crystallization solution occupy the channel, with the acetate positioned where
the phosphothreonine linkage of CoBSH would be (Figure 3C). Presumably 1–2 further
waters would replace the acetate under physiological conditions. Other than W3 and W7, the
waters form hydrogen bonds with protein (Figure 5). One water (W2) occupies the same site
as the CoBSH thiol. Presumably due to the loss of favorable interactions that exist when
CoBSH is present, the hydrophobic side-chain of Valα482 adopts a second conformation
modeled at 60 % occupancy (Supporting Information, Figure S7).
Position of the “bridging” water, W1
The equivalent of W1 has previously been observed in MCRred1-silent and MCRox1-silent
crystal structures where, in the presence of CoBSH and HSCoM, it is sited equidistant (3.2
Å) between the two coenzyme thiols (PDB codes: 1hbn, 1hbo, 1hbu) and thus been termed
the “bridging water” (Figure 1) (5, 33). However, in the MCRHSCoM structure, due to the
presence of the more polarized W2 water, W1 is displaced away from HSCoM to maximize
the hydrogen bond interaction with W2 (2.9 Å to W2; 3.5 Å to HSCoM thiol, Figure 5). In
the MCRCoB5SH structure that also contained W2, the electron density indicated that this
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repositioning of W1 towards W2 also occurred. In contrast, the MCRCoB6SH structure
contained 100% thiol at the CoBSH position, but a partial occupancy of HSCoM (50%). In
this case the electron density for W1 indicated it had moved towards the nickel to form an
optimal hydrogen bond with a Ni-ligating water that was only present in the absence of
HSCoM (3.7 Å to CoB(6)SH thiol; 3.0 Å to Ni-ligating water, Supporting Information,
Figure S5). In all structures reported here, W1 (if present) appears to be a sensitive indicator
of the relative electronegativity of the Ni-ligated atom to that occupying the position of the
CoBSH thiol, and was a useful check in the crystallographic modeling and refinement
process.
Flexibility in the substrate channel: Alternative protein conformers
The binding site of HSCoM (and presumably methyl-SCoM) is more deeply buried within
the enzyme, and so it must enter prior to CoBSH for productive chemistry to occur. As
binding of CoBSH in the absence of co-substrate would be inhibitory, it was suggested that
a conformational change upon methyl-SCoM binding might lower the Kd for CoBSH, and
thus promote an ordered mechanism. Compared to the 1hbn MCRox1-silent and 1hbu
MCRred1-silent crystal structures, which both have full occupancy HSCoM, the lower
occupancy of HSCoM in the 1hbo MCRred1-silent structure was associated with significantly
greater flexibility within the channel, and the ability to model a second conformation of a
Gly-rich amino acid stretch that formed part of the CoBSH channel. This suggested that
methyl-SCoM binding might cause the channel to become more ordered, increasing the
affinity of MCR for CoBSH by conformational restriction rather than a switch mechanism
where the structure reorganizes from one well-defined conformer to another (33). In the
MCRHSCoM data containing 30 % CoBSH and 100% HSCoM, the Fo-Fc difference electron
density map at one of the two independent active sites in the ASU contained positive peaks
that suggested the presence of an alternate conformation also involving this part of the
polypeptide (Supporting Information, Figure S8). Using this as a guide, a similar second
conformation involving seven contiguous amino acid residues of the same Gly-rich amino
acid stretch (β366–372) could be modeled and refined at 20 % occupancy leaving no
residual difference density. Parts of the α′ subunit (α′111–129 and α′237–242) that are in
close proximity to this stretch of amino acids also exhibit second conformations, with the
main-chain carbonyl of α′243 in van der Waals contact with the B ring of F430 tetrapyrrole
(Supporting Information, Figure S9). Modeling these at 20% occupancy accounted for the
weak positive Fo-Fc difference electron density peaks observed in these areas. The evidence
of alternate conformers in these areas lends support to the proposal that increased flexibility
in the substrate channel propagates through the protein (33).
The MCRCoB6SH crystal structure contains 50 % CoB6SH, 50% CoBSH and 50% HSCoM.
In this case there is no evidence of an alternate loop conformation in either active site of the
ASU. However, as CoBSH and CoB6SH combined are at 100% occupancy, it is not
surprising their favorable interactions with the substrate channel would reduce
conformational disorder, despite the partial occupancy of HSCoM.
Analogues longer than CoBSH; CoB8SH and CoB9SH
Both analogues could be accommodated in the MCR substrate channel (Figures 3D and 3E).
The electron density supported final models containing 50% CoB8SH for MCRCoB8SH (1.8
Å resolution) and 60 % CoB9SH for MCRCoB9SH (1.45 Å resolution). The phosphate head-
groups are in identical positions to those of CoBSH, CoB5SH, CoB6SH (Figure 4) (5, 33).
Both analogues follow the crystallographically observed chain path of bound CoBSH, with
the extra atoms displacing the W1 water and placing the thiols closer to the nickel (Figure
6). CoB9SH does have a second conformer that deviates from the CoBSH path, but the thiol
position for this conformer and the CoBSH-tracking conformer are identical (Figure 3E and
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Supporting Information, Figure S10). Interestingly, the thiol of CoB8SH is not coincident
with the CoB9SH carbon that precedes the CoB9SH thiol (Figure 6). CoB8SH is an MCR
inhibitor with an apparent Ki of 15 μM (3). CoB9SH has never been tested for inhibition of
MCR-catalyzed methane formation, but it is reasonable to assume that it would be an
inhibitor.
CoBXSH thiol-to-nickel spatial relationship
The position of CoBSH in previous crystal structures poses a conundrum (5, 33). In all the
proposed catalytic mechanisms, CoBSH must interact with species generated at the nickel.
Perplexingly, the sulfur of the CoBSH substrate is 8.8 Å from the Ni(II) in the MCRox1-silent
and both MCRred1-silent crystal structures, and 6.4 Å from the thiol of the substrate analogue
HSCoM (Figure 1). Modeling studies demonstrated that the addition of a methyl group to
HSCoM did not bridge this gap (35, 45, 53). Therefore, a conformational change has been
postulated that would enable CoBSH to penetrate deeper into the substrate channel, and thus
approach closer to any nickelbound species. The heterodisulfide product in the MCRsilent
crystal structure has the CoBSH portion in virtually the same place as in MCRox1-silent,
giving no clue to possible structural changes that might occur to facilitate CoBSH reacting
with nickel-associated intermediates (5, 33).
Trigonometry suggests that if the alkanoyl chain of CoBSH or its analogue is in an extended
conformation, each additional unit in the chain would lead to the thiol moving ~1.2 Å
towards the Ni. Until this study there have been no crystal structures of CoBSH analogues in
complex with MCR, so mechanistic studies using different chain length analogues of
CoBSH assumed that shorter analogues would trace the observed path of CoBSH, and
longer analogues would penetrate about ~1.2 Å deeper per additional chain unit into the
channel. In the case of the shortest analogue CoB5SH, it does indeed follow the path of
CoBSH, with the thiol of CoB5SH being 2.8 Å away from the thiol position of CoBSH.
However, due to the conformation CoBSH adopts when bound in the substrate channel, the
difference in the S-Ni distance is small; the CoB5SH thiol being only 0.5 Å farther from the
Ni than CoBSH (8.8 Å for CoBSH vs. 9.3 Å for CoB5SH) (Table 2). This is due to the
alkanoyl chain of CoBSH not being in an extended conformation from carbons 4 to 6
(carbon 1 is the carbonyl carbon). CoB6SH on the other hand adopts a conformation that
places its thiol in virtually the same position as the thiol of CoBSH (Figure 4 and Table 2).
This is consistent with CoB6SH being a substrate. However, the kcat is 1000-fold lower than
for CoBSH (3, 35) although its Km value (180 μM, Dey & Ragsdale, in preparation) is
similar to that of CoBSH (Km = 75 μM (3)). The reason for this may be that the shorter
alkanoyl chain may not enable the analogue thiol to approach the nickel close enough for
efficient catalysis, and thus explain why CoB6SH is such a poor substrate.
In the case of the longer CoBXSH analogues, the sulfur of CoB8SH is 2.6 Å closer to the Ni
ion of F430 than that of CoBSH, and 2.5 Å closer to the thiol of HSCoM (Figure 6 and Table
2). The CoB9SH molecule follows the path of CoBSH, and reaches only a little further into
the substrate channel than CoB8SH, with the CoB9SH thiol positioned 2.9 Å closer to the Ni
than the thiol of CoBSH (Figure 6 and Table 2). This is only 0.3 Å closer than the distance
observed for the CoB8SH thiol, even though they are non-coincident. The distance to the
thiol of HSCoM is 2.6 Å closer than that of the substrate, CoBSH; only 0.1 Å closer than the
CoB8SH thiol. The two analogue thiols are above an annular hydrophobic aromatic
environment created by Pheα330, Tyrα333, Pheα443, Pheβ361 and Tyrβ367 that lies
between them and F430 (Figure 6). As a result, penetrating further into the channel may be
energetically unfavorable, consistent with the small difference in relative distances between
the CoB8SH/CoB9SH thiols and the HSCoM thiol/F430 nickel. The annulus is proposed to
be catalytically important in positioning methyl-SCoM and stabilizing the methane product,
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and the tyrosines have been proposed to be proton donors associated with mechanism II
(Scheme 2B) (5, 33).
Thus, there appear to be three preferential distances for thiols (including that of HSCoM)
within the MCR substrate channel; HSCoM at 2.4 Å, CoB6SH/CoBSH at 8.7 – 8.8 Å and
CoB8SH/CoB9SH at 5.9 – 6.2 Å from the nickel of F430 (Table 2).
Recent ENDOR, high-field continuous and pulse EPR work has identified changes in nickel
co-ordination when CoBSH is added to MCRred1c (active Ni(I)-MCRred1 + HSCoM) (14,
15, 18, 31). This generates up to 50% MCRred2 which is comprised of two distinct nickel co-
ordination geometries; an axial MCRred2a formally assigned as a Ni(III)-hydride, and a
rhombic MCRred2r in which the thiol of HSCoM is a Ni(I) ligand (Supporting Information,
Scheme S1). Formation of MCRred2 could also be induced by addition of CoBSH substrate
analogues CoBS-CH3 and CoBS-CF3, which have a chain length one unit longer than
substrate CoBSH (18, 53). The CoBS-CF3 enabled 19F-ENDOR studies to be performed,
and demonstrated that following CoBS-CF3 addition the remaining MCRred1 species had
Ni(I)-19F distances of 6.2 – 7.7 Å. This distance range agreed with a CoBS-CF3 model
created using the CoBSH position observed in the MCRox1-silent crystal structure (53).
However, in the MCRred2 species the Ni(I)-19F distances had shortened indicating a
movement of ~ 2 Å towards the nickel (MCRred2a Ni(I)-19 F, 4.0 – 5.5 Å; MCRred2r
Ni(I)-19F, 4.5 – 5.7 Å). In the case of the CoB8SH analogue (the closest equivalent to CoBS-
CF3 used in this study) the thiol to Ni(II) distance lies between the distance ranges observed
in the CoBS-CF3 studies, and so the fluorine(s) of the CoBS-CF3 in MCRred2 might
penetrate a little further into the hydrophobic annulus in the MCRred2 species. As the
alkanoyl chain of CoBSH is not fully extended it could easily undergo a similar
conformation change to that observed in the MCRred2 state.
Reaction of MCR Ni(III)-alkyl species’ with CoB8SH and CoB9SH
The two longer CoBXSH analogues have been shown to undergo alkylation when reacted
with MCRPS, a [Ni(III)-alkyl ↔ Ni(II)-alkylsulfonate radical] formed from reaction of
Ni(I)-MCRred1 with bromopropanesulfonate (BPS) (Supporting Information, Scheme S1)
(20, 23, 30, 45, 54). BPS is a substrate of MCRred1 in a reaction that involves a rapid
CoBSH-independent nucleophilic attack by Ni(I) on BPS to displace bromide and generate
MCRPS at a rate ~60-fold faster than generation of methane from CoBSH and methyl-
HSCoM (20, 45). Certain thiols can eliminate the propylsulfonate to yield a thioether
product and regenerate MCRred1, although at a rate 1000-fold slower than methane
formation (45). Both CoB8SH and CoB9SH can react with MCRPS to regenerate MCRred1,
but CoBSH cannot. The overall second-order rate constant for the reactivation of MCR by
CoB8SH is 160 M−1s−1, whereas for CoB9SH the reaction is more sluggish (12 M−1s−1).
CoB9SH might be expected to be closer to the proximal Ni ligand. It was therefore proposed
that this caused steric interference and explained why CoB9SH was a poorer reactivator of
MCR than CoB8SH. Our study has shown that the thiols of these two analogues are placed
such that they are approximately the same distance (~3.7 Å) from the thiol of HSCoM
ligated to the Ni atom (Table 2). The Ni(II)-HSCoM bond is 2.4 Å, whereas an Ni(III)-alkyl
bond is expected to be ~2 Å (24, 33), thus indicating that a conformational change is
required to effect the nucleophilic attack of the CoB8SH and CoB9SH thiols on an alkyl-
bound species. It would thus appear that a conformational change, such as observed in
MCRred2, is required for this chemistry also (53).
A Ni(III)-alkyl species is akin to the first intermediate in mechanism I of MCR-catalyzed
methane formation, Ni(III)-methyl (MCRMe, Supporting Information, Scheme S1, Scheme
2A) (11, 27). MCRMe has been shown to be capable of generating MCRred1 and methyl-
SCoM upon addition of HSCoM (which is the reverse of mechanism I, step 1, Scheme 2A);
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similar chemistry to the observed formation of a thioether product from the Ni(III)-alkyl.
Further addition of CoBSH following HSCoM treatment of MCRMe led to methane and
heterodisulfide formation, the natural products of methanogenesis. Although this lends
credence to mechanism I, it should be noted that like MCRPS, MCRMe in these experiments
was generated artificially. However, the MCRCoB9SH crystal structure demonstrates that the
two additional methylene units in the alkanoyl chain c.f. CoBSH, do not necessarily translate
into direct interaction of the thiol with the nickel proximal ligand. However, this could
represent the favorable position for a CoBSH thiol interacting with the methyl group of
methyl-SCoM. Just as the alkanoyl chain of CoB6SH has a more extended conformation
than CoBSH in the substrate channel, CoBSH could also adopt a more extended
conformation so that its thiol was in a similar position as the thiol of CoB8SH, priming it for
reaction with a nickel bound species.
If a significant conformational change is required early in MCR-catalyzed chemistry, which
would be a requirement of mechanism I, catalysis may well involve a rearrangement of the
aromatic amino acid annulus due to the presence of the methyl of methyl-SCoM, and this
might enable deeper penetration of CoBSH into MCR (Figure 6). All the crystal structures in
this study, and those solved previously, are of the inactive Ni(II)-MCR, which disfavors
close approach to the nickel in the absence of Ni(I)-bound methyl-SCoM, even in the case of
CoB9SH.
Conclusion
The goal of this study was to induce structural changes within the substrate channel and
active site of MCR using analogues of coenzyme CoBSH. It was hoped that this would shed
light on the nature of conformational changes that have been proposed to occur in MCR
catalysis. We have shown that that the CoBXSH analogues do not lead to any significant
conformational changes within the context of inactive Ni(II)-MCR. Therefore, it may be that
methyl-SCoM is the key coenzyme, in combination with a nickel oxidation state of 1 (and
3), that triggers a conformational change bringing the thiol of CoBSH closer to the nickel.
Thus, the crystal structure of the Ni(I)-methyl-SCoM/MCR complex may be required to
structurally define the conformational changes required for MCR-mediated chemistry.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
X-ray data were collected at beam-line 14-BM-C and photoreduction studies at 14-ID-B, BioCARS at the
Advanced Photon Source (APS), Argonne National Laboratory, Argonne, IL. We thank Vukica Srajer and Yu-
Sheng Chen for valuable assistance during data collection. Use of the Advanced Photon Source was supported by
the U.S. Department of Energy, Basic Energy Sciences, Office of Science under Contract No. DE-
AC02-06CH11357. Use of the BioCARS Sector 14 was supported by the National Institutes of Health, National
Center for Research Resources, under grant number RR007707. Computer resources were provided by the Basic
Sciences Computing Laboratory of the University of Minnesota Supercomputing Institute, and we thank Can
Ergenekan for his support. We also thank Ed Hoeffner at the Kahlert Structural Biology Laboratory (KSBL) at the
University of Minnesota. Use of the KSBL was supported by a Minnesota Partnership for Biotechnology and a
Medical Genomics Grant SPAP-05-0013-P-FY06.
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Figure 1.
The active site and substrate channel in the MCRox1-silent crystal structure (PDB code 1hbn)
(9). Coenzyme F430, CoBSH and HSCoM are drawn as stick colored by atom (carbon: dark
grey). The nickel is displayed as a green sphere, water as a red sphere. Interactions are
drawn as dashed lines, and the corresponding distance is indicted in Angstroms (Å). The
path of the substrate channel was defined in the absence of F430, CoBSH, HSCoM and
water, with the surface closest to the viewer cut away. The figure was generated using
PyMOL (http://www.pymol.org).
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Figure 2.
Drawing of CoBSH analogues; (A) N-5-mercaptopentanoylthreonine phosphate (CoB5SH);
(B) N-6-mercaptohexanoylthreonine phosphate (CoB6SH); (C) N-8-
mercaptooctanoylthreonine phosphate (CoB8SH); (D) N-9-mercaptononanoylthreonine
phosphate (CoB9SH).
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Figure 3.
The active sites and substrate channels of the MCR crystal structures.; (A) MCRCoB5SH; (B)
MCRCoB6SH; (C) MCRHSCoM; (D) MCRCoB8SH; (E) MCRCoB9SH. 2Fo-Fc electron density
map around the CoBXSH analogues, waters in the CoBSH binding part of the channel and
the acetate ion (contoured at 1σ) is shown as a blue mesh. The protein is drawn as cartoon.
CoBXSH and acetate are drawn as stick and colored by atom (carbon: CoB5SH orange;
CoB6SH pale yellow; CoB8SH light blue; CoB9SH magenta; acetate white). Coenzyme F430
and HSCoM are drawn as stick colored by atom (carbon: F430 dark grey; HSCoM medium
grey). The nickel is displayed as a green sphere, and waters as red spheres. The figure was
generated using PyMOL (http://www.pymol.org/).
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Figure 4.
Overlay of CoBSH (from PDB code 1hbn) and the different CoBSH analogues. CoBXSH are
drawn as stick with the thiol represented by a sphere, and colored CoB5SH orange; CoB6SH
pale yellow; CoBSH light green; CoB8SH light blue; CoB9SH magenta. The protein is
drawn as cartoon. Coenzyme F430 and HSCoM are drawn as stick colored by atom (carbon:
F430 dark grey; HSCoM medium grey). The nickel is displayed as a green sphere. The figure
was generated using PyMOL (http://www.pymol.org/).
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Figure 5.
Hydrogen bonding diagram for the water structure modeled in MCRHSCoM. The water
molecules are named as in Figure 3C (W1–W9); WA, WB and WC are water molecules that
are present in all structures (i.e. in concert with the substrate CoBSH and the CoBSH
analogues). Interactions between surrounding residues and the water molecules are drawn as
dashed lines, and the corresponding distance is indicated in Angstroms (Å).
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Figure 6.
Stereo image of the annulus of aromatic amino acids distal of coenzyme F430. The protein is
drawn as cartoon with the side-chains of the aromatic residues drawn as white stick.
CoBSH, (from PDB code 1hbn (9)), CoB8SH and CoB9SH are drawn as stick with the thiols
represented by spheres, and colored CoBSH light green; CoB8SH light blue; CoB9SH
magenta. Coenzyme F430 and HSCoM are drawn as stick colored by atom (carbon: F430
dark grey; HSCoM medium grey). The nickel is displayed as a green sphere. The figure was
generated using PyMOL (http://www.pymol.org/).
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Scheme 1.
Reaction catalyzed by methyl-coenzyme M reductase
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Scheme 2.
Two of the proposed catalytic mechanisms for methyl-coenzyme M reductase; (A)
mechanism I; (B) mechanism II.
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Table 1
X-ray Data Collection, Processing and Refinement Statistics
Data collection and processing statistics
Name of data set
MCRCoB5SH
MCRCoB6SH
MCRHSCoM
MCRCoB8SH
MCRCoB9SH
Measured reflections
1969388
2427498
1440665
1160543
1425506
Unique reflections
553755
446253
405349
211803
401701
Resolution (Å) a
50.0–1.30 (1.35–1.30)
50.0–1.40 (1.45–1.40)
50.0–1.45 (1.50–1.45)
50.0–1.80 (1.86–1.80)
50.0–1.45 (1.50–1.45)
Completeness (%) a
97.1 (78.1)
99.9 (100.0)
99.5 (99.7)
99.8 (100.0)
98.1 (95.4)
R-sym (%) a,b
5.5 (32.9)
7.3 (44.7)
6.2 (44.0)
8.4 (47.7)
5.6 (42.5)
I/σI a
22.3 (3.6)
20.4 (4.0)
20.2 (3.2)
21.8 (3.9)
24.3 (3.2)
Space group
P21
P21
P21
P21
P21
Refinement and model building statistics
Resolution (Å) a
20.49–1.30 (1.33–1.30)
19.89–1.40 (1.44–1.40)
20.15–1.45 (1.49–1.45)
19.93–1.80 (1.84–1.80)
20.07–1.45 (1.48–1.45)
No. of reflection in working set a
525817 (30239)
423854 (25833)
384868 (25791)
201128 (11193)
381474 (23611)
No. of reflection in test set a
27777 (1576)
22348 (1331)
20362 (1319)
10625 (557)
20163 (1210)
R-work (%) c
14.32
13.04
13.47
14.95
13.58
R-free (%) d
16.56
15.53
16.22
19.54
16.44
ESU (Å) R-work/R-free
0.044/0.046
0.049/0.051
0.056/0.059
0.121/0.119
0.057/0.060
No. protein atoms
20087
19960
20265
19750
20036
No. coenzyme atoms
218
220
180
224
272
No. ligand atoms
37
62
52
26
49
No. water molecules
2443
2352
2516
1893
2432
RMS
bond lengths (Å)
0.033
0.033
0.032
0.028
0.032
bond angles (deg.)
2.693
2.625
2.468
2.059
2.549
Ramachandran plot (%)
favored
97.8
97.5
97.6
97.2
97.7
allowed
2.1
2.4
2.3
2.7
2.1
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disallowed
0.1
0.1
0.1
0.1
0.1
Average B-factor (Å2)
protein
12.42
13.35
12.12
17.22
12.73
coenzymes
8.20
9.24
7.25
11.24
8.27
ligands
31.95
35.48
28.29
33.76
32.92
waters
22.95
24.89
23.85
26.79
24.09
over all
13.54
14.57
13.40
18.02
13.93
Occupancy of HSCoM per active site (%)e
90/90
50/50
100/100
90/90
90/85
Occupancy of CoBSH per active site (%) e
50/50
50/50
30/30
50/50
40/40
CoBSH analogue, occupancy per active site (%) e
CoB5SH, 50/50
CoB6SH, 50/50
CoB8SH, 50/50
CoB9SH, 60/60
Other molecule, occupancy per active site (%) e
Acetate, 70/70
aValues in brackets correspond to the highest resolution shell.
bR-sym = ΣhklΣNj=1|Ihkl−Ihkl(j)|/ΣhklN*Ihkl, sum over all reflections and all observations N, with Ihkl(j) intensity of the jth observation of reflection hkl and Ihkl mean intensity of the reflection hkl.
cR-work = Σ||Fo| − |Fc||/Σ|Fo|, where |Fo| = observed structure factor amplitude and |Fc| = calculated structure factor amplitude.
dR-free, R-factor based on 5% of the data excluded from refinement.
eOccupancy of model in each of the two crystallographically independent active sites in the ASU
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Table 2
Distances from analogue thiols.
CoBXS - SCoM distance (Å)
CoBXS - Ni distance (Å)
CoB5SH
7.11/7.11a
9.30/9.30
CoB6SH
6.26/6.26
8.70/8.70
CoB7SH (substrate) b
6.37/6.39
8.73/8.77
CoB8SH
3.75/3.78
6.16/6.17
CoB9SH
3.71/3.68
5.96/5.91
aDistances in the two crystallographically independent active sites in the ASU
bDistances in the 1.16 Å resolution MCRox1-silent structure (PDB code 1hbn)(33)
Biochemistry. Author manuscript; available in PMC 2011 September 7.
|
3M38
|
The roles of Glutamates and Metal ions in a rationally designed nitric oxide reductase based on myoglobin: I107E FeBMb (No metal ion binding to FeB site)
|
Roles of glutamates and metal ions in a rationally
designed nitric oxide reductase based on myoglobin
Ying-Wu Lina,2, Natasha Yeunga, Yi-Gui Gaob, Kyle D. Minerc, Shiliang Tiana, Howard Robinsond, and Yi Lua,c,1
aDepartment of Chemistry, bGeorge L. Clark X-Ray Facility and 3M Materials Laboratory, andcDepartment of Biochemistry, University of Illinois at
Urbana-Champaign, Urbana, IL 61801; and dDepartment of Biology, Brookhaven National Laboratory, Upton, NY 11973
Edited by Harry B. Gray, California Institute of Technology, Pasadena, CA, and approved April 1, 2010 (received for review January 14, 2010)
A structural and functional model of bacterial nitric oxide reductase
(NOR) has been designed by introducing two glutamates (Glu) and
three histidines (His) in sperm whale myoglobin. X-ray structural
data indicate that the three His and one Glu (V68E) residues bind
iron, mimicking the putative FeB site in NOR, while the second Glu
(I107E) interacts with a water molecule and forms a hydrogen bond-
ing network in the designed protein. Unlike the first Glu (V68E),
which lowered the heme reduction potential by ∼110 mV, the
second Glu has little effect on the heme potential, suggesting that
thenegativelychargedGluhasadifferentrolein redoxtuning.More
importantly, introducing the second Glu resulted in a ∼100% in-
crease in NOR activity, suggesting the importance of a hydrogen
bonding network in facilitating proton delivery during NOR reactiv-
ity. In addition, EPR and X-ray structural studies indicate that the
designed protein binds iron, copper, or zinc in the FeB site, each with
different effects on the structures and NOR activities, suggesting
that both redox activity and an intermediate five-coordinate
heme-NO species are important for high NOR activity. The designed
protein offers an excellent model for NOR and demonstrates the
power of using designed proteins as a simpler and more well-
defined system to address important chemical and biological issues.
biomimetic models ∣heme-copper oxidase ∣metalloprotein ∣protein design ∣
protein engineering
R
ational design of proteins that mimic both structure and func-
tion of more complex native enzymes has been a long sought-
after goal, as the process is an ultimate test of our knowledge and
an excellent means to develop advanced biocatalysts (1–3).
Although designed proteins that model the structure of native
enzymes have been known for a while (4–10), successful designs
of proteins that mimic both the structure and function of native
enzymes have been reported only recently (11–16). While being
able to design such functional proteins is laudable, the impact of
such an achievement would be greater if the designed proteins
can be used to address fundamental issues in chemistry and biol-
ogy that are difficult to tackle by other methods. One primary
example is the roles of conserved glutamates and metal ions in
bacterial nitric oxide reductase (NOR) (17–19).
NO is critical for all life (20). Bacterial denitrification is a cru-
cial part of the nitrogen cycle in nature that involves a four-step,
five-electron reduction of nitrate (NO3
−) to dinitrogen (N2)
(17, 19). Bacterial NOR is a membrane-bound protein that
catalyzes one step of this process, namely, the two-electron reduc-
tion of NO to N2O (17, 19). With no crystal or solution structure
available for bacterial NOR to date, sequence alignments and
homology modeling (21, 22) have indicated that NOR is structu-
rally homologous to the largest subunit (subunit I) of heme-
copper oxidases (HCOs) (23), enzymes that catalyze reduction
of O2 to water. The active sites of both NOR and HCO contain
a proximal histidine-coordinated heme and a distal three histi-
dine-coordinated metal center. However, the metal center in
HCOs is occupied by a copper (called CuB), whereas a nonheme
iron is present in NOR (called FeB) (23, 24). In addition, two
conserved glutamates, shown by modeling to be close to the
FeB site (21, 22), are found to be essential for NOR activity
(24, 25). Some members of HCOs such as cytochrome cbb3
oxidase display NOR activity (26–28), although the activity is
∼50-fold lower than native NOR (26). Therefore, it is important
to elucidate the structural features, specifically the roles of the
conserved glutamates close to the FeB site and metal ions (copper
vs. iron), responsible for the reduction of NO to N2O.
To address these issues, biochemical and biophysical studies of
native NOR and its variants have been carried out (24, 25, 29–37).
For example, Richardson and coworkers investigated the effects
of amino acid substitutions of the five conserved glutamates
(E122 and E125 presumed to face the periplasm and E198,
E202, and E267 located in the interior of the membrane, close
to the catalytic site) in the catalytic subunit of Paracoccus deni-
trificans, NorB. The E122A, E125A, E198A, and E267A variants
were inactive, indicating that these four glutamates are crucial for
NOR activity (24, 25, 32, 33). On the other hand, Reimann et al.
constructed a 3D model of NorB using homology modeling with
the structures of HCOs as templates and suggested a plausible
pathway consisting of these conserved glutamates for proton
delivery (22). Despite these successes, the roles of the conserved
glutamates and metal ions still remain to be fully elucidated,
partly because of the difficulty in obtaining native NOR in high
yield and the lack of a 3D structure. Even if these problems are
resolved, it is still difficult to replace iron in the native FeB site
with other metal ions, and spectroscopic studies of native NOR
are often complicated by the presence of other metal cofactors
(e.g., low-spin heme).
To overcome these limitations, a number of synthetic models of
NOR using small organic molecules as ligands, have been made
in which the nonheme FeB site can be replaced by a copper ion
(17, 38–45). In addition, since these model systems lack addi-
tional metal-binding sites, spectroscopic studies are often simpli-
fied. Therefore, studies of these synthetic models have offered
many insights. For example, Collman et al. showed that a fully
reduced heme/nonheme FeB compound can react with two
equivalents of NO leading to the formation of one equivalent
of N2O and a bis-ferric product (41). On the other hand, Karlin
and coworkers showed that a small heme/Cu complex can effi-
ciently lead to reductive coupling of NO to N2O (43). However,
it is also difficult to obtain the synthetic models in high yield due
to the multiple steps required in chemical synthesis. Because of
this limitation, no synthetic NOR model containing the two key
conserved Glu residues (E198 and E267 in NOR) has been
Author contributions: Y.-W.L., N.Y., and Y.L. designed research; Y.-W.L., N.Y., Y.-G.G.,
K.D.M., and S.T. performed research; Y.-W.L., N.Y., Y.-G.G., K.D.M., H.R., and Y.L. analyzed
data; and Y.-W.L. and Y.L. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The atomic coordinates and structure factors have been deposited in the
Protein Data Bank, www.pdb.org (PDB ID codes 3M38, 3M39, 3M3A, and 3M3B).
1To whom correspondence should be addressed. E-mail: yi-lu@illinois.edu.
2Present address: School of Chemistry and Chemical Engineering, University of South
China, Hengyang 421001, China.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1000526107/-/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.1000526107
PNAS ∣May 11, 2010 ∣vol. 107 ∣no. 19 ∣8581–8586
BIOCHEMISTRY
CHEMISTRY
reported. It is also difficult to substitute different metal ions in
the same metal-binding site without perturbing the site geometry
and distances to the heme iron, as most ligands are not as rigid as
those in native enzymes and different metal ions have different
geometric and ligand donor set preferences.
We have recently designed a structural and functional protein
model of bacterial NOR by engineering three histidines and one
glutamate into the distal pocket of sperm whale myoglobin
(swMb, L29H, F43H, H64, and V68E, named FeBMb) (14). Like
synthetic models, this “bottom-up” approach complements the
“top-down”approachofthestudyofnativeNORinthatitprovides
insights into whether certain “necessary” structural elements are
enough to impart enzyme function. Thanks in part to recent
advances in computational, molecular, and structural biology,
the designed myoglobin protein model is much easier to synthesize
and to crystallize than either native NOR or synthetic models.
Since myoglobin has often been used for the development and
calibration of numerous spectroscopic techniques (46–48), it is
an ideal choice for spectroscopic studies. More importantly, the
rigid protein network allows precise placement of either glutamate
or metal ions in myoglobin to address their roles in NOR activity.
Toward this goal, we have demonstrated that both the histidines
and one of the glutamates are essential for iron binding and NO
reduction activity (14). However, the role of the second Glu close
to the FeB site and the role of different metal ions in the FeB site
have not been addressed.
To address these important issues and to design even closer
protein models of NOR, we introduced herein the second Glu
to the second coordination sphere of the FeB site by mutating
an Ile to a Glu (named I107E FeBMb). We show that the second
Glu results in a ∼100% increase in NOR activity through hydro-
gen bonding interactions and that the two glutamates have
dramatically different effects on the heme reduction potential.
Additionally, by comparing the EPR, electrochemistry, X-ray
structures, and NOR activity of iron, copper, and zinc derivatives
of the designed protein, we have obtained deeper insights into the
roles of metal ions in NOR.
Results
Structure and Function of Fe(II)-I107E FeBMb. The X-ray crystal struc-
tures of heme-containing I107E FeBMb without metal ion in the
FeB site and with Fe2þ in the FeB site are solved at 1.42-Å and
1.65-Å resolution, respectively (Fig. 1 A and B and Table S1). In
the absence of metal ions in the FeB site, the structure shows a
water molecule in the FeB site, which forms hydrogen bonds with
NE2 atoms of all three His residues, both OE1 and OE2 atoms of
E68, and the OE2 atom of E107 (Fig. 1A). Upon binding Fe2þ,
the Fe(II)-I107E FeBMb structure shows that Fe2þ is coordinated
by three His, the OE2 atom of E68, and one water molecule.
Notably, a water molecule bridges Fe2þ in the FeB site and
the second glutamate (E107) with a distance of 2.32 Å to the
OE2 atom of E107 (Fig. 1B).
To probe the conformational changes of introducing the
second Glu (E107), we performed a structural alignment of Fe
(II)-I107E FeBMb and the previously reported FeðIIÞ-FeBMb
(14). The comparison, shown in Fig. 2, indicates that both the
polypeptide chain and the active site overlap well with each other.
In addition, the two nonheme irons are located at similar posi-
tions with a 0.36-Å separation from each other. In contrast,
E68 underwent a significant conformational rearrangement in
the presence of E107. These observations suggest that the active
site of FeBMb can be tuned by the formation of an extended
hydrogen bonding network, resulting from the introduction of
a second glutamate residue.
The binding of Fe2þ to deoxy I107E FeBMb was further mon-
itored by EPR (Fig. 3A). Since deoxy myoglobin contains Fe(II)
heme that exhibits no EPR signals in X-band EPR (14), we added
blue copper Cu(II)-azurin (49), a redox partner of native NOR
(19), to oxidize both the reduced heme and nonheme irons in Fe
(II)-I107E FeBMb to EPR-active Fe(III). Upon addition of Cu
(II)-azurin, the oxidation of deoxy I107E FeBMb resulted in
EPR signals at g ¼ 6.12 and 5.56, typical of a high-spin heme-
Fe(III). Upon addition of Fe2þ, however, a decrease of the
heme-Fe(III) EPR signals was observed, indicating that the
Fe2þ, when bound to the FeB site and oxidized by Cu(II)-azurin,
is spin-coupled to heme-Fe(III). Such a spin coupling mimics
that in NOR (35, 50–53), suggesting that I107E FeBMb models
NOR closely, at least in this respect.
To probe the role of the second Glu (E107) in NO reduction
activity, we measured the yield of N2O production by Fe(II)-
I107E FeBMb with excess NO under one turnover conditions.
We monitored N2O formation in the headspace of the solution
using GC/MS and compared this result to that of FeðIIÞ-FeBMb,
which lacks the second Glu (Fig. 4). Remarkably, Fe(II)-I107E
FeBMb
displays higher
activity than
FeðIIÞ-FeBMb.
After
∼20 hr, ∼24% N2O was produced by Fe(II)-I107E FeBMb, in
H29
A
H29
B
H64
E68
E107
3.04
2.81
3.02
2.20
2.24
2.24
2.12
2.21
H64
E68
E107
3.03
H43
2.15
3.41
3.16
2.62
2.32
H43
2.26
H93
H93
H29
C
H29
D
2.04
2.03
H64
E68
E107
2.09
2.10
2.91
2.21
H64
E68
E107
2.26
2.29
2.10
2.18
2.10
4.47
H43
2.07
3.04
H43
2.68
H93
H93
Fig. 1.
Crystal structures of I107E FeBMb (A) (PDB ID code 3M38), Fe(II)-I107E
FeBMb (B) (PDB ID code 3M39), Cu(II)-I107E FeBMb (C) (PDB ID code 3M3A),
and Zn(II)-I107E FeBMb (D) (PDB ID code 3M3B). Water molecules, Fe(II),
Cu(II), and Zn(II) are represented by red, green, orange, and gray spheres,
respectively.
Fig. 2.
Overlay of Fe(II)-I107E FeBMb (cyan) (PDB ID code 3M39) with
FeðIIÞ-FeBMb (orange) (PDB ID code 3K9Z).
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contrast to ∼10% yield for FeðIIÞ-FeBMb, strongly indicating that
the second Glu plays an important role in NO reduction, likely
facilitating proton uptake during NO reduction.
Other Metal Ions Binding to I107E FeBMb. To find out if the resting
state of the protein, i.e., oxidized or met I107E FeBMb, can bind
other metal ions, Cu2þ or Zn2þ was titrated into met I107E
FeBMb and monitored by EPR spectroscopy (Fig. 3 B and C).
In the absence of metal ions, met I107E FeBMb exhibited
high-spin heme signals at g ¼ 6.03, 5.08, and 1.98 (Fig. 3B, black
line). Upon addition of 2 eq of Cu2þ, the signals at g ¼ 6.03 and
5.08 decreased and a broad peak around g ¼ 2.95 increased,
probably due to spin coupling between heme-Fe(III) and Cu2þ
in the FeB site. In contrast, addition of Zn2þ, a metal ion with
no unpaired electrons [i.e., incapable of spin coupling to heme-
Fe(III)], produced an increase in the high-spin heme signals at
g ¼ 5.88 and 5.60 (Fig. 3C), indicating that the interaction be-
tween E68 and heme iron was weakened after metal binding.
The X-ray crystal structures of I107E FeBMb with Cu2þ or
Zn2þ in the FeB site were solved at 1.37-Å and 1.60-Å resolution,
respectively (Fig. 1 C and D and Table S1). Compared to Fe(II)-
I107E FeBMb (Fig. 1B), a similar binding site was observed
for Cu(II)-I107E FeBMb (Fig. 1C), where H29, H43, and H64
coordinate to Cu2þ with distances of 2.09, 2.10, and 2.04 Å,
respectively, slightly shorter than the corresponding distances
in the Fe2þ structure. In comparison to Fe(II)-I107E FeBMb,
the water bridging the Cu2þ and the second Glu (E107) is shifted
toward Cu2þ in the FeB site (2.03 Å) with respect to E107
(3.04 Å). Interestingly, this bridging water molecule was not
observed in Zn(II)-I107E FeBMb (Fig. 1D), but the two O atoms
of E68 coordinate to Zn2þ with similar distances (2.26 Å for OE1
and 2.29 Å for OE2). The longer distance between OE1 of E68
B
6.12
I107E FeBMb + Azurin
I107E Fe Mb + 0 5 eq Fe
2+ + Azurin
A
5.56
I107E FeBMb + 0.5 eq Fe
+ Azurin
I107E FeBMb + 1.0 eq Fe
2+ + Azurin
I107E FeBMb + 2.0 eq Fe
2+ + Azurin
1.98
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
5.08
5.38
Magnetic Field (Gauss)
5.88
I107E FeBMb
I107E FeBMb + 0.5 eq Zn
2+
I107E Fe Mb
1 0 eq Zn
2+
C
6.03
I107E FeBMb
I107E FeBMb + 0.5 eq Cu
2+
I107E Fe Mb + 1 0 eq Cu
2+
I107E FeBMb + 1.0 eq Zn
I107E FeBMb + 2.0 eq Zn
2+
I107E FeBMb + 1.0 eq Cu
I107E FeBMb + 2.0 eq Cu
2+
1.98
~2.95
1.98
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
5.60
Magnetic Field (Gauss)
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
5.08
Magnetic Field (Gauss)
Fig. 3.
EPR spectra of deoxy I107E FeBMb (0.5 mM protein in 50 mM Bis-Tris, pH 7.0) with increasing concentrations of Fe2þ in the presence of wild-type
Cu(II)-azurin (A), and oxidized I107E FeBMb with Cu2þ (B) or Zn2þ (C). Spectra were collected at 4 K, 5 mW power, and 9.05 GHz.
30
25
Fe(II)-I107E FeBMb
Fe(II)-FeBMb
15
20
10
%)
oduction (%
N2O pro
0
5
0
4
8
12
16
20
Incubation time (hr)
Fig. 4.
Time-dependent N2O production by Fe(II)-I107E FeBMb (▴) and
FeðIIÞ-FeBMb (●) with ∼50 eq. NO under single turnover conditions. The yield
was determined by a comparison of the ratio of NO∶N2O peaks from the
GC/MS chromatograms.
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and heme iron in the Zn-bound structure (2.68 Å) in comparison
to the Cu- and Fe-bound structures, is likely the result of a weaker
interaction, which is also supported by an observed increase of
the high-spin heme signals in the EPR spectra upon Zn2þ binding
(Fig. 3C). These results suggest I107E FeBMb is capable of incor-
porating different metal ions into its designed FeB site, offering
an excellent opportunity to compare the role of these metal ions
in the same protein scaffold.
Effect of Glutamates and Metal Ions on the Redox Potential of I107E
FeBMb. Since EPR and X-ray structural studies indicate metal
binding to I107E FeBMb, we used spectroelectrochemistry to
measure the effects of glutamates and metal ions on the heme
reduction potential. When there is no metal ion in the FeB site,
the I107E FeBMb displays a reduction potential of −134 3 mV
vs. the normal hydrogen electrode (NHE) (Fig. S1A), similar to
that of FeBMb (−158 4 mV) without the I107E mutation (14).
In the presence of Cu2þ, I107E FeBMb has a reduction potential
(−137 2 mV) (Fig. S1B) almost identical to that of the same
protein in the absence of metal ions in the FeB site, indicating
that copper binding to the FeB site has little effect on the reduction
potential of the heme iron. This observation is similar to that
observed for Cu2þ binding to CuBMb (CuðIIÞ-CuBMb, 80 mV
vs. CuBMb, 77 mV) (54). On the other hand, the presence of
Fe2þ and Zn2þ increased the reduction potential of I107E
FeBMb from −134 3 mV to −64 3 mV vs. NHE (Fig. S1C)
and −105 2 mV vs. NHE (Fig. S1D), respectively. The different
effects of Cu2þ, Fe2þ, and Zn2þ on the reduction potential of
I107E-FeBMb indicate that these metal ions in the FeB site may
play different roles through different coordination properties.
NOR Activity of I107E FeBMb in the Presence of Different Metal Ions.
The NO reduction activity of I107E FeBMb in the presence of
Fe2þ, Cuþ, or Zn2þ was monitored by GC/MS under single turn-
over conditions. When Fe(II)-I107E FeBMb was exposed to
excess NO, N2O could be observed to form with increased yield
over time (Fig. S2). Similarly, N2O formation was observed for
Cu(I)-I107E FeBMb, indicating that Fe or Cu binding to the
FeB site results in comparable NOR activities. It should be noted
that because of the high solubility of N2O (∼25 mM in water at
room temperature), GC/MS cannot be used to quantify the rates
of NO reduction under these conditions. In contrast, no N2O
formation was observed with redox inactive Zn2þ, which demon-
strates that redox active Fe2þ or Cuþ in the FeB site plays a
crucial role in NO reduction.
To gain deeper insight into the process of NO reduction, EPR
studies were further performed to monitor the initial process of
NO reduction. In the absence of metal ions, the EPR spectrum of
ferrous I107E FeBMb-NO shows hyperfine splitting resulting
from bound NO and the proximal histidine, indicating the forma-
tion of a six-coordinate ferrous heme-NO species (Fig. 5, top
line). After incubation of Fe(II)-I107E FeBMb with excess
NO, a distinct three-line hyperfine structure appears at 15 min
(Fig. 5A), suggesting the formation of a five-coordinate ferrous
heme-NO species as a result of cleavage of the proximal
His-Fe heme bond (55). A three-line hyperfine structure was also
observed for Cuþ and Zn2þ, except that the signal intensity is low-
er with Cu(I)-I107E FeBMb-NO (Fig. 5B) and more pronounced
in Zn(II)-I107E FeBMb-NO (Fig. 5C). The lower intensity of
the three-line hyperfine structure for Cu(I)-I107E FeBMb-NO
suggests the major species formed is a six-coordinate ferric
heme-NO complex, which is EPR silent (41). These differences
further suggest that the metal ion in the FeB site plays a key role
in formation of the intermediates, thereby tuning NOR activity.
Discussion
Using Rationally Designed Proteins to Address Important Issues in
Chemistry and Biology. Important issues such as the roles of the
conserved glutamates and nonheme FeB in NOR have been
previously addressed using biochemical and biophysical studies
or biomimetic modeling (24, 25, 27–37, 45, 56, 57). As a comple-
mentary approach, rational protein design, using small, easy-to-
produce and well-characterized proteins such as myoglobin,
offers a powerful method with which to gain insights into more
complex native enzymes such as NOR (14). Similar to synthetic
models (41, 43), the metal ion at the putative FeB site in the
protein model can be substituted freely. Better yet, Glu residues
can be placed at precise locations in the protein, including the
secondary coordination sphere, due to its rigid network. By care-
fully choosing a suitable protein template, rational protein design
could be generally applied to address other important issues in
chemistry and biology.
The Roles of Glutamates. Although two conserved glutamates
(E198 and E267) are known to be crucial for NOR activity
(24, 25), their roles are not well defined (18, 19). In a previous
study (14), we demonstrated that one Glu, E68, is important for
both iron binding and NOR activity of FeBMb. The crystal struc-
tures of both FeðIIÞ-FeBMb and Fe(II)-I107E FeBMb show that
one O atom of E68 directly coordinates to FeB (Fig. 2). In syn-
thetic models of NOR, it has also been found that the presence of
a glutamic acid mimic significantly increases the stability of iron
binding to the FeB site (40). Furthermore, a theoretical study by
Blomberg et al. (58) showed that a model with an FeB coordi-
nated by three histidines, one glutamate, and one water molecule
provides an energetically feasible reaction mechanism of NO
reduction. However, the structural model of NOR constructed
recently by Reimann et al. (22) shows that the closest conserved
Glu (E267) still has its carboxylate O atom 7 Å away from FeB,
which suggests that Glu may not bind to FeB in native NOR. One
interesting finding from our study is that the Glu (E68) under-
went a significant conformational rearrangement in the presence
of another Glu (E107) (Fig. 2). Therefore, the FeBMb provides a
viable model of NOR that is consistent with Blomberg’s model,
but cannot rule out Reimann’s model due to possible conforma-
tion changes.
While the role of the first Glu is still uncertain until a 3D struc-
ture of NOR in its active form is available, the role of the second
Glu is even less defined. We address this question by introducing
a second Glu (E107) to FeBMb. The crystal structures shown in
Fig. 1 indicate that E107 interacts with a water molecule and
forms a hydrogen bonding network in both Fe(II)-I107E
FeBMb and Cu(II)-I107E FeBMb. Interestingly, although a
similar water molecule was observed in the active site of FeðIIÞ-
FeBMb (Fig. 2), activity assay data indicate that the presence of
E107 in Fe(II)-I107E FeBMb increases NOR activity by ∼100%
C
5 min
N
t l
A
5 min
B
2+
5 min
1 min
No metal
1 min
5 min
F
2+
No metal
1
i
5 min
+
No metal
2+
Zn
2+
5 min
Fe
2+
5 min
1 min
Fe
C
+
5 min
1 min
Cu
+
Z
2
Zn
2+
15 min
F
2+
Fe
15 min
5 min
C
+
Cu
+
15 min
5 min
Zn
2+
15 min
Fe2+
5
Cu
+
15 min
3000
3100
3200
3300
3400
Magnetic Field (Gauss)
3000
3100
3200
3300
3400
Magnetic Field (Gauss)
3000
3100
3200
3300
3400
Magnetic Field (Gauss)
Fig. 5.
EPR spectra of deoxy I107E FeBMb (0.5 mM) with no metal bound in
the presence of NO after 5 min (top line), with 2 eq Fe2þ (A), Cuþ (B), or Zn2þ
(C) incubated with excess NO (∼200 eq) for 1, 5, and 15 min. Spectra were
collected in 50 mM Bis-Tris pH 7.0 at 30 K, 0.2 mW power, and 9.05 GHz.
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Lin et al.
(Fig. 4), suggesting that the second Glu may potentionally play a
role in providing one of the protons during reduction of NO to
N2O. Although free Glu outside a protein has a pKa ∼4.3, the stu-
dies of native NOR showed that the pKa of its Glu close to the
active site has a value of ∼6.6 (22, 25, 33). The hydrogen binding
network in our protein models may contribute to the fine-tuning
of the Glu pKa to be more neutral, similar to that in NOR. More-
over, it is interesting that Cu(I)-I107E FeBMb also shows NOR
activity, which provides an interesting protein model of HCOs
with NOR function (26–28), even though Glu residues are not
conserved in native HCOs.
Additionally, spectroelectrochemical studies showed that the
reduction potential of I107E FeBMb with no metal ion in the
FeB site is similar to that of FeBMb, but much lower than that
of CuBMb (77 mV) (54), which contains the same three His,
but no Glu in the metal-binding site above the heme. Since both
FeBMb and I107E FeBMb contain the V68E mutation that has
been shown to decrease the heme reduction potential of native
myoglobin from 59 mV to −137 mV (59), it is likely that the
introduction of a negatively charged Glu close to the heme group
is what is responsible for the dramatically lower heme redox
potential. A conserved glutamate, predicted to be located near
the catalytic heme b3 in NOR, was proposed to be responsible
for a ∼260-mV decrease in reduction potential (60 mV) in
comparison to the other two heme centers, heme b (345 mV)
and heme c (310 mV) (60). Our FeBMb and I107E FeBMb mod-
els mimic this feature of NOR. Notably, although introduction of
the first Glu (E68) lowered the heme potential by ∼110 mV (14),
introduction of the second Glu via the I107E mutation did not
result in a significant difference in the heme reduction potential,
suggesting that the effect of the two conserved Glu residues in
NOR on heme reduction potential is not additive, with the effects
highly dependent on the location of the Glu.
The Roles of Metal Ions. The roles of metal ions in NOR are an-
other important question as iron is found in the native FeB site
and HCO employs copper at the corresponding CuB site. With
different metal ions in the FeB site, the crystal structures clearly
show the heme and nonheme dinuclear center existing in differ-
ent local environments (Fig. 1). Although a similar hydrogen
bond network is formed in both Fe(II)-I107E FeBMb and Cu
(II)-I107E FeBMb, the conformation of E68 and E107 with re-
spect to the nonheme metal center and heme iron is different
from each other. Moreover, the coordination geometry differs
significantly with Zn2þ in the FeB site. A hydrogen bond is absent
from the Zn crystal structure, but both the O atoms of E68 act as
metal-binding ligands. These observations demonstrate that the
identity of the metal ion in the FeB site can tune the active site
through their interactions with the His and Glu ligands, resulting
in formation of different coordination geometries with different
hydrogen bonds.
In addition to structural fine-tuning, the metal ion at the FeB
site can also tune the heme iron reduction potential in I107E
FeBMb. Spectroelectrochemical studies showed that the binding
of Fe2þ or Zn2þ results in an increase in the heme reduction
potential by ∼70 mV and ∼30 mV, respectively (Fig. S1). In
the case of Cu(II)-I107E FeBMb, the crystal structure shows that
OE1 of E68 is closer to the heme iron (2.07 Å) (Fig. 1C) than its
metal-free form (2.15 Å) (Fig. 1A). The stronger interaction
from the negatively charged E68 could offset the effect of
positively charged Cu2þ binding, resulting in similar reduction
potentials
observed
for
Cu(II)-I107E
FeBMb
and
I107E
FeBMb.
In a previous study (61), EPR data showed that during NO
reduction, the binding of Cuþ to the CuB site of CuBMb can
weaken the proximal heme Fe-His bond, while complete cleavage
of the heme Fe-His bond occurred when Zn2þ was bound to
CuBMb-NO. In this study, we observed that a five-coordinate
heme-NO species was formed with Fe2þ, Cuþ, or Zn2þ bound
to the FeB site of I107E FeBMb (Fig. 5). Significantly, a five-
coordinate heme-NO species has also been observed for both
NOR (30, 31, 35) and the member of the HCO family with the
highest NO reduction activity, cytochrome cbb3 oxidases (26, 62).
However, this species was not observed for FeðIIÞ-FeBMb-NO
and CuðIÞ-FeBMb-NO, which lack the second Glu (E107). In
both these cases, the proximal heme Fe-His bond was only
weakened, as indicated by a decrease of the nine-line hyperfine
splitting signals in the EPR spectra (Fig. S3). These observations
suggest that formation of a five-coordinate heme-NO species may
play an important role in NOR reactivity.
Conclusions
We have successfully designed a structural and functional model
of NOR, by introducing a second glutamate in the vicinity of the
FeB site, named I107E FeBMb. This protein model mimics native
NOR more closely by bearing the structural feature of three his-
tidines and two glutamates in the FeB site, as predicted for native
NOR. We have demonstrated that the two glutamates can play
different roles in NO reduction activity; namely, one acts as a li-
gand to FeB (E68), and the other acts as a proton transfer group
(E107). Furthermore, by substituting different metal ions into the
nonheme metal site, we have demonstrated that FeB plays crucial
roles in fine-tuning the active site by donating electrons and by
mediating the formation of a five-coordinate heme-NO inter-
mediate during NO reduction. In the absence of a crystal struc-
ture for native NOR, this study offers an ideal protein model and
provides valuable structural as well as mechanistic information
for native NOR.
Materials and Methods
Protein Preparation. I107E FeBMb (swMb L29H/F43H/V68E/I107E) was con-
structed, expressed, and purified using the procedure described previously
(14). The purity and identity were confirmed by SDS-PAGE and electrospray
ionized MS: observed: 17; 392 1 Da; calculated: 17,391 Da.
EPR Spectroscopy. EPR spectra were recorded on a Bruker ESP 300 equipped
with an Oxford liquid helium cryostat and an ITC4 temperature controller.
The samples of met I107E FeBMb, Cu(II)-, or Zn(II)-I107E FeBMb were pre-
pared as described previously (14). The samples of NO-bound deoxy
I107E FeBMb, Cu(I)-, Fe(II)-, or Zn(II)-I107E FeBMb were prepared by inject-
ing 1 mL of purified NO gas into the EPR tube containing 300 μL of protein
(0.5 mM, 10% glycerol, in 50 mM Bis-Tris pH 7.0). The samples were then
flash frozen in liquid N2 after incubation for 1, 5, or 15 min. The molar
extinction coefficient of the Soret band of I107E FeBMb at 406 nm
(175 mM−1 · cm−1), calculated using the standard hemochromagen method
(63), was used to determine protein concentration. The metal sources of
Cu(I), Cu(II), Zn(II), and Fe(II) were ½ðCH3CNÞ4CuPF6, CuSO4, ZnSO4 · 7H2O,
and FeCl2, respectively.
Spectroelectrochemical Measurements. Protein reduction potentials were
measured using an optically transparent thin layer electrode as previously
described (64). The potential of the working electrode was applied in
the positive direction for deoxy I107E FeBMb with Fe2þ and in the negative
direction for metal free and with Cu2þ or Zn2þ. Other procedures are the
same as described previously (54).
X-Ray Crystallographic Studies. Fe(II)-I107E FeBMb was crystallized anaerobi-
cally in a glove box at room temperature using the conditions described for
FeðIIÞ-FeBMb (14). I107E FeBMb, Cu(II)-, or Zn(II)-I107E FeBMb were crystallized
aerobically. Diffraction-quality crystals were soaked in a cryoprotectant
solution of 30% PEG 400 and flash frozen in liquid nitrogen. Diffraction data
were collected at the Brookhaven National Lab Synchrotron Light Source
X12C beamline. The crystal structure was solved using the same method
as for FeðIIÞ-FeBMb (14).
NOR Activity Assay. NO reduction was monitored by GC/MS. The protein was
reduced to the deoxy form by excess dithionite that was removed with a
size-exclusion column (PD-10). Then 2 eq metal, Cu(I), Fe(II), or Zn(II), was
added to the protein solution (0.6 mM, 3 mL in 50 mM Bis-Tris buffer, pH
7.0). The samples were prepared anaerobically in a glove box. Purified NO
Lin et al.
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gas was injected into the head space of the reaction flask with the molar
ratio of NO∶protein ≈50∶1. Other procedures are the same as described
previously (14, 61).
ACKNOWLEDGMENTS. We thank Dr. Mark J. Nilges for help with EPR analysis,
and Furong Sun and Beth D. Eves for aiding in GC/MS data collection. This
work was supported by NIH Grant GM062211.
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|
3M39
|
The roles of glutamates and metal ions in a rationally designed nitric oxide reductase based on myoglobin: Fe(II)-I107E FeBMb (Fe(II) binding to FeB site)
|
Roles of glutamates and metal ions in a rationally
designed nitric oxide reductase based on myoglobin
Ying-Wu Lina,2, Natasha Yeunga, Yi-Gui Gaob, Kyle D. Minerc, Shiliang Tiana, Howard Robinsond, and Yi Lua,c,1
aDepartment of Chemistry, bGeorge L. Clark X-Ray Facility and 3M Materials Laboratory, andcDepartment of Biochemistry, University of Illinois at
Urbana-Champaign, Urbana, IL 61801; and dDepartment of Biology, Brookhaven National Laboratory, Upton, NY 11973
Edited by Harry B. Gray, California Institute of Technology, Pasadena, CA, and approved April 1, 2010 (received for review January 14, 2010)
A structural and functional model of bacterial nitric oxide reductase
(NOR) has been designed by introducing two glutamates (Glu) and
three histidines (His) in sperm whale myoglobin. X-ray structural
data indicate that the three His and one Glu (V68E) residues bind
iron, mimicking the putative FeB site in NOR, while the second Glu
(I107E) interacts with a water molecule and forms a hydrogen bond-
ing network in the designed protein. Unlike the first Glu (V68E),
which lowered the heme reduction potential by ∼110 mV, the
second Glu has little effect on the heme potential, suggesting that
thenegativelychargedGluhasadifferentrolein redoxtuning.More
importantly, introducing the second Glu resulted in a ∼100% in-
crease in NOR activity, suggesting the importance of a hydrogen
bonding network in facilitating proton delivery during NOR reactiv-
ity. In addition, EPR and X-ray structural studies indicate that the
designed protein binds iron, copper, or zinc in the FeB site, each with
different effects on the structures and NOR activities, suggesting
that both redox activity and an intermediate five-coordinate
heme-NO species are important for high NOR activity. The designed
protein offers an excellent model for NOR and demonstrates the
power of using designed proteins as a simpler and more well-
defined system to address important chemical and biological issues.
biomimetic models ∣heme-copper oxidase ∣metalloprotein ∣protein design ∣
protein engineering
R
ational design of proteins that mimic both structure and func-
tion of more complex native enzymes has been a long sought-
after goal, as the process is an ultimate test of our knowledge and
an excellent means to develop advanced biocatalysts (1–3).
Although designed proteins that model the structure of native
enzymes have been known for a while (4–10), successful designs
of proteins that mimic both the structure and function of native
enzymes have been reported only recently (11–16). While being
able to design such functional proteins is laudable, the impact of
such an achievement would be greater if the designed proteins
can be used to address fundamental issues in chemistry and biol-
ogy that are difficult to tackle by other methods. One primary
example is the roles of conserved glutamates and metal ions in
bacterial nitric oxide reductase (NOR) (17–19).
NO is critical for all life (20). Bacterial denitrification is a cru-
cial part of the nitrogen cycle in nature that involves a four-step,
five-electron reduction of nitrate (NO3
−) to dinitrogen (N2)
(17, 19). Bacterial NOR is a membrane-bound protein that
catalyzes one step of this process, namely, the two-electron reduc-
tion of NO to N2O (17, 19). With no crystal or solution structure
available for bacterial NOR to date, sequence alignments and
homology modeling (21, 22) have indicated that NOR is structu-
rally homologous to the largest subunit (subunit I) of heme-
copper oxidases (HCOs) (23), enzymes that catalyze reduction
of O2 to water. The active sites of both NOR and HCO contain
a proximal histidine-coordinated heme and a distal three histi-
dine-coordinated metal center. However, the metal center in
HCOs is occupied by a copper (called CuB), whereas a nonheme
iron is present in NOR (called FeB) (23, 24). In addition, two
conserved glutamates, shown by modeling to be close to the
FeB site (21, 22), are found to be essential for NOR activity
(24, 25). Some members of HCOs such as cytochrome cbb3
oxidase display NOR activity (26–28), although the activity is
∼50-fold lower than native NOR (26). Therefore, it is important
to elucidate the structural features, specifically the roles of the
conserved glutamates close to the FeB site and metal ions (copper
vs. iron), responsible for the reduction of NO to N2O.
To address these issues, biochemical and biophysical studies of
native NOR and its variants have been carried out (24, 25, 29–37).
For example, Richardson and coworkers investigated the effects
of amino acid substitutions of the five conserved glutamates
(E122 and E125 presumed to face the periplasm and E198,
E202, and E267 located in the interior of the membrane, close
to the catalytic site) in the catalytic subunit of Paracoccus deni-
trificans, NorB. The E122A, E125A, E198A, and E267A variants
were inactive, indicating that these four glutamates are crucial for
NOR activity (24, 25, 32, 33). On the other hand, Reimann et al.
constructed a 3D model of NorB using homology modeling with
the structures of HCOs as templates and suggested a plausible
pathway consisting of these conserved glutamates for proton
delivery (22). Despite these successes, the roles of the conserved
glutamates and metal ions still remain to be fully elucidated,
partly because of the difficulty in obtaining native NOR in high
yield and the lack of a 3D structure. Even if these problems are
resolved, it is still difficult to replace iron in the native FeB site
with other metal ions, and spectroscopic studies of native NOR
are often complicated by the presence of other metal cofactors
(e.g., low-spin heme).
To overcome these limitations, a number of synthetic models of
NOR using small organic molecules as ligands, have been made
in which the nonheme FeB site can be replaced by a copper ion
(17, 38–45). In addition, since these model systems lack addi-
tional metal-binding sites, spectroscopic studies are often simpli-
fied. Therefore, studies of these synthetic models have offered
many insights. For example, Collman et al. showed that a fully
reduced heme/nonheme FeB compound can react with two
equivalents of NO leading to the formation of one equivalent
of N2O and a bis-ferric product (41). On the other hand, Karlin
and coworkers showed that a small heme/Cu complex can effi-
ciently lead to reductive coupling of NO to N2O (43). However,
it is also difficult to obtain the synthetic models in high yield due
to the multiple steps required in chemical synthesis. Because of
this limitation, no synthetic NOR model containing the two key
conserved Glu residues (E198 and E267 in NOR) has been
Author contributions: Y.-W.L., N.Y., and Y.L. designed research; Y.-W.L., N.Y., Y.-G.G.,
K.D.M., and S.T. performed research; Y.-W.L., N.Y., Y.-G.G., K.D.M., H.R., and Y.L. analyzed
data; and Y.-W.L. and Y.L. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The atomic coordinates and structure factors have been deposited in the
Protein Data Bank, www.pdb.org (PDB ID codes 3M38, 3M39, 3M3A, and 3M3B).
1To whom correspondence should be addressed. E-mail: yi-lu@illinois.edu.
2Present address: School of Chemistry and Chemical Engineering, University of South
China, Hengyang 421001, China.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1000526107/-/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.1000526107
PNAS ∣May 11, 2010 ∣vol. 107 ∣no. 19 ∣8581–8586
BIOCHEMISTRY
CHEMISTRY
reported. It is also difficult to substitute different metal ions in
the same metal-binding site without perturbing the site geometry
and distances to the heme iron, as most ligands are not as rigid as
those in native enzymes and different metal ions have different
geometric and ligand donor set preferences.
We have recently designed a structural and functional protein
model of bacterial NOR by engineering three histidines and one
glutamate into the distal pocket of sperm whale myoglobin
(swMb, L29H, F43H, H64, and V68E, named FeBMb) (14). Like
synthetic models, this “bottom-up” approach complements the
“top-down”approachofthestudyofnativeNORinthatitprovides
insights into whether certain “necessary” structural elements are
enough to impart enzyme function. Thanks in part to recent
advances in computational, molecular, and structural biology,
the designed myoglobin protein model is much easier to synthesize
and to crystallize than either native NOR or synthetic models.
Since myoglobin has often been used for the development and
calibration of numerous spectroscopic techniques (46–48), it is
an ideal choice for spectroscopic studies. More importantly, the
rigid protein network allows precise placement of either glutamate
or metal ions in myoglobin to address their roles in NOR activity.
Toward this goal, we have demonstrated that both the histidines
and one of the glutamates are essential for iron binding and NO
reduction activity (14). However, the role of the second Glu close
to the FeB site and the role of different metal ions in the FeB site
have not been addressed.
To address these important issues and to design even closer
protein models of NOR, we introduced herein the second Glu
to the second coordination sphere of the FeB site by mutating
an Ile to a Glu (named I107E FeBMb). We show that the second
Glu results in a ∼100% increase in NOR activity through hydro-
gen bonding interactions and that the two glutamates have
dramatically different effects on the heme reduction potential.
Additionally, by comparing the EPR, electrochemistry, X-ray
structures, and NOR activity of iron, copper, and zinc derivatives
of the designed protein, we have obtained deeper insights into the
roles of metal ions in NOR.
Results
Structure and Function of Fe(II)-I107E FeBMb. The X-ray crystal struc-
tures of heme-containing I107E FeBMb without metal ion in the
FeB site and with Fe2þ in the FeB site are solved at 1.42-Å and
1.65-Å resolution, respectively (Fig. 1 A and B and Table S1). In
the absence of metal ions in the FeB site, the structure shows a
water molecule in the FeB site, which forms hydrogen bonds with
NE2 atoms of all three His residues, both OE1 and OE2 atoms of
E68, and the OE2 atom of E107 (Fig. 1A). Upon binding Fe2þ,
the Fe(II)-I107E FeBMb structure shows that Fe2þ is coordinated
by three His, the OE2 atom of E68, and one water molecule.
Notably, a water molecule bridges Fe2þ in the FeB site and
the second glutamate (E107) with a distance of 2.32 Å to the
OE2 atom of E107 (Fig. 1B).
To probe the conformational changes of introducing the
second Glu (E107), we performed a structural alignment of Fe
(II)-I107E FeBMb and the previously reported FeðIIÞ-FeBMb
(14). The comparison, shown in Fig. 2, indicates that both the
polypeptide chain and the active site overlap well with each other.
In addition, the two nonheme irons are located at similar posi-
tions with a 0.36-Å separation from each other. In contrast,
E68 underwent a significant conformational rearrangement in
the presence of E107. These observations suggest that the active
site of FeBMb can be tuned by the formation of an extended
hydrogen bonding network, resulting from the introduction of
a second glutamate residue.
The binding of Fe2þ to deoxy I107E FeBMb was further mon-
itored by EPR (Fig. 3A). Since deoxy myoglobin contains Fe(II)
heme that exhibits no EPR signals in X-band EPR (14), we added
blue copper Cu(II)-azurin (49), a redox partner of native NOR
(19), to oxidize both the reduced heme and nonheme irons in Fe
(II)-I107E FeBMb to EPR-active Fe(III). Upon addition of Cu
(II)-azurin, the oxidation of deoxy I107E FeBMb resulted in
EPR signals at g ¼ 6.12 and 5.56, typical of a high-spin heme-
Fe(III). Upon addition of Fe2þ, however, a decrease of the
heme-Fe(III) EPR signals was observed, indicating that the
Fe2þ, when bound to the FeB site and oxidized by Cu(II)-azurin,
is spin-coupled to heme-Fe(III). Such a spin coupling mimics
that in NOR (35, 50–53), suggesting that I107E FeBMb models
NOR closely, at least in this respect.
To probe the role of the second Glu (E107) in NO reduction
activity, we measured the yield of N2O production by Fe(II)-
I107E FeBMb with excess NO under one turnover conditions.
We monitored N2O formation in the headspace of the solution
using GC/MS and compared this result to that of FeðIIÞ-FeBMb,
which lacks the second Glu (Fig. 4). Remarkably, Fe(II)-I107E
FeBMb
displays higher
activity than
FeðIIÞ-FeBMb.
After
∼20 hr, ∼24% N2O was produced by Fe(II)-I107E FeBMb, in
H29
A
H29
B
H64
E68
E107
3.04
2.81
3.02
2.20
2.24
2.24
2.12
2.21
H64
E68
E107
3.03
H43
2.15
3.41
3.16
2.62
2.32
H43
2.26
H93
H93
H29
C
H29
D
2.04
2.03
H64
E68
E107
2.09
2.10
2.91
2.21
H64
E68
E107
2.26
2.29
2.10
2.18
2.10
4.47
H43
2.07
3.04
H43
2.68
H93
H93
Fig. 1.
Crystal structures of I107E FeBMb (A) (PDB ID code 3M38), Fe(II)-I107E
FeBMb (B) (PDB ID code 3M39), Cu(II)-I107E FeBMb (C) (PDB ID code 3M3A),
and Zn(II)-I107E FeBMb (D) (PDB ID code 3M3B). Water molecules, Fe(II),
Cu(II), and Zn(II) are represented by red, green, orange, and gray spheres,
respectively.
Fig. 2.
Overlay of Fe(II)-I107E FeBMb (cyan) (PDB ID code 3M39) with
FeðIIÞ-FeBMb (orange) (PDB ID code 3K9Z).
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Lin et al.
contrast to ∼10% yield for FeðIIÞ-FeBMb, strongly indicating that
the second Glu plays an important role in NO reduction, likely
facilitating proton uptake during NO reduction.
Other Metal Ions Binding to I107E FeBMb. To find out if the resting
state of the protein, i.e., oxidized or met I107E FeBMb, can bind
other metal ions, Cu2þ or Zn2þ was titrated into met I107E
FeBMb and monitored by EPR spectroscopy (Fig. 3 B and C).
In the absence of metal ions, met I107E FeBMb exhibited
high-spin heme signals at g ¼ 6.03, 5.08, and 1.98 (Fig. 3B, black
line). Upon addition of 2 eq of Cu2þ, the signals at g ¼ 6.03 and
5.08 decreased and a broad peak around g ¼ 2.95 increased,
probably due to spin coupling between heme-Fe(III) and Cu2þ
in the FeB site. In contrast, addition of Zn2þ, a metal ion with
no unpaired electrons [i.e., incapable of spin coupling to heme-
Fe(III)], produced an increase in the high-spin heme signals at
g ¼ 5.88 and 5.60 (Fig. 3C), indicating that the interaction be-
tween E68 and heme iron was weakened after metal binding.
The X-ray crystal structures of I107E FeBMb with Cu2þ or
Zn2þ in the FeB site were solved at 1.37-Å and 1.60-Å resolution,
respectively (Fig. 1 C and D and Table S1). Compared to Fe(II)-
I107E FeBMb (Fig. 1B), a similar binding site was observed
for Cu(II)-I107E FeBMb (Fig. 1C), where H29, H43, and H64
coordinate to Cu2þ with distances of 2.09, 2.10, and 2.04 Å,
respectively, slightly shorter than the corresponding distances
in the Fe2þ structure. In comparison to Fe(II)-I107E FeBMb,
the water bridging the Cu2þ and the second Glu (E107) is shifted
toward Cu2þ in the FeB site (2.03 Å) with respect to E107
(3.04 Å). Interestingly, this bridging water molecule was not
observed in Zn(II)-I107E FeBMb (Fig. 1D), but the two O atoms
of E68 coordinate to Zn2þ with similar distances (2.26 Å for OE1
and 2.29 Å for OE2). The longer distance between OE1 of E68
B
6.12
I107E FeBMb + Azurin
I107E Fe Mb + 0 5 eq Fe
2+ + Azurin
A
5.56
I107E FeBMb + 0.5 eq Fe
+ Azurin
I107E FeBMb + 1.0 eq Fe
2+ + Azurin
I107E FeBMb + 2.0 eq Fe
2+ + Azurin
1.98
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
5.08
5.38
Magnetic Field (Gauss)
5.88
I107E FeBMb
I107E FeBMb + 0.5 eq Zn
2+
I107E Fe Mb
1 0 eq Zn
2+
C
6.03
I107E FeBMb
I107E FeBMb + 0.5 eq Cu
2+
I107E Fe Mb + 1 0 eq Cu
2+
I107E FeBMb + 1.0 eq Zn
I107E FeBMb + 2.0 eq Zn
2+
I107E FeBMb + 1.0 eq Cu
I107E FeBMb + 2.0 eq Cu
2+
1.98
~2.95
1.98
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
5.60
Magnetic Field (Gauss)
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
5.08
Magnetic Field (Gauss)
Fig. 3.
EPR spectra of deoxy I107E FeBMb (0.5 mM protein in 50 mM Bis-Tris, pH 7.0) with increasing concentrations of Fe2þ in the presence of wild-type
Cu(II)-azurin (A), and oxidized I107E FeBMb with Cu2þ (B) or Zn2þ (C). Spectra were collected at 4 K, 5 mW power, and 9.05 GHz.
30
25
Fe(II)-I107E FeBMb
Fe(II)-FeBMb
15
20
10
%)
oduction (%
N2O pro
0
5
0
4
8
12
16
20
Incubation time (hr)
Fig. 4.
Time-dependent N2O production by Fe(II)-I107E FeBMb (▴) and
FeðIIÞ-FeBMb (●) with ∼50 eq. NO under single turnover conditions. The yield
was determined by a comparison of the ratio of NO∶N2O peaks from the
GC/MS chromatograms.
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and heme iron in the Zn-bound structure (2.68 Å) in comparison
to the Cu- and Fe-bound structures, is likely the result of a weaker
interaction, which is also supported by an observed increase of
the high-spin heme signals in the EPR spectra upon Zn2þ binding
(Fig. 3C). These results suggest I107E FeBMb is capable of incor-
porating different metal ions into its designed FeB site, offering
an excellent opportunity to compare the role of these metal ions
in the same protein scaffold.
Effect of Glutamates and Metal Ions on the Redox Potential of I107E
FeBMb. Since EPR and X-ray structural studies indicate metal
binding to I107E FeBMb, we used spectroelectrochemistry to
measure the effects of glutamates and metal ions on the heme
reduction potential. When there is no metal ion in the FeB site,
the I107E FeBMb displays a reduction potential of −134 3 mV
vs. the normal hydrogen electrode (NHE) (Fig. S1A), similar to
that of FeBMb (−158 4 mV) without the I107E mutation (14).
In the presence of Cu2þ, I107E FeBMb has a reduction potential
(−137 2 mV) (Fig. S1B) almost identical to that of the same
protein in the absence of metal ions in the FeB site, indicating
that copper binding to the FeB site has little effect on the reduction
potential of the heme iron. This observation is similar to that
observed for Cu2þ binding to CuBMb (CuðIIÞ-CuBMb, 80 mV
vs. CuBMb, 77 mV) (54). On the other hand, the presence of
Fe2þ and Zn2þ increased the reduction potential of I107E
FeBMb from −134 3 mV to −64 3 mV vs. NHE (Fig. S1C)
and −105 2 mV vs. NHE (Fig. S1D), respectively. The different
effects of Cu2þ, Fe2þ, and Zn2þ on the reduction potential of
I107E-FeBMb indicate that these metal ions in the FeB site may
play different roles through different coordination properties.
NOR Activity of I107E FeBMb in the Presence of Different Metal Ions.
The NO reduction activity of I107E FeBMb in the presence of
Fe2þ, Cuþ, or Zn2þ was monitored by GC/MS under single turn-
over conditions. When Fe(II)-I107E FeBMb was exposed to
excess NO, N2O could be observed to form with increased yield
over time (Fig. S2). Similarly, N2O formation was observed for
Cu(I)-I107E FeBMb, indicating that Fe or Cu binding to the
FeB site results in comparable NOR activities. It should be noted
that because of the high solubility of N2O (∼25 mM in water at
room temperature), GC/MS cannot be used to quantify the rates
of NO reduction under these conditions. In contrast, no N2O
formation was observed with redox inactive Zn2þ, which demon-
strates that redox active Fe2þ or Cuþ in the FeB site plays a
crucial role in NO reduction.
To gain deeper insight into the process of NO reduction, EPR
studies were further performed to monitor the initial process of
NO reduction. In the absence of metal ions, the EPR spectrum of
ferrous I107E FeBMb-NO shows hyperfine splitting resulting
from bound NO and the proximal histidine, indicating the forma-
tion of a six-coordinate ferrous heme-NO species (Fig. 5, top
line). After incubation of Fe(II)-I107E FeBMb with excess
NO, a distinct three-line hyperfine structure appears at 15 min
(Fig. 5A), suggesting the formation of a five-coordinate ferrous
heme-NO species as a result of cleavage of the proximal
His-Fe heme bond (55). A three-line hyperfine structure was also
observed for Cuþ and Zn2þ, except that the signal intensity is low-
er with Cu(I)-I107E FeBMb-NO (Fig. 5B) and more pronounced
in Zn(II)-I107E FeBMb-NO (Fig. 5C). The lower intensity of
the three-line hyperfine structure for Cu(I)-I107E FeBMb-NO
suggests the major species formed is a six-coordinate ferric
heme-NO complex, which is EPR silent (41). These differences
further suggest that the metal ion in the FeB site plays a key role
in formation of the intermediates, thereby tuning NOR activity.
Discussion
Using Rationally Designed Proteins to Address Important Issues in
Chemistry and Biology. Important issues such as the roles of the
conserved glutamates and nonheme FeB in NOR have been
previously addressed using biochemical and biophysical studies
or biomimetic modeling (24, 25, 27–37, 45, 56, 57). As a comple-
mentary approach, rational protein design, using small, easy-to-
produce and well-characterized proteins such as myoglobin,
offers a powerful method with which to gain insights into more
complex native enzymes such as NOR (14). Similar to synthetic
models (41, 43), the metal ion at the putative FeB site in the
protein model can be substituted freely. Better yet, Glu residues
can be placed at precise locations in the protein, including the
secondary coordination sphere, due to its rigid network. By care-
fully choosing a suitable protein template, rational protein design
could be generally applied to address other important issues in
chemistry and biology.
The Roles of Glutamates. Although two conserved glutamates
(E198 and E267) are known to be crucial for NOR activity
(24, 25), their roles are not well defined (18, 19). In a previous
study (14), we demonstrated that one Glu, E68, is important for
both iron binding and NOR activity of FeBMb. The crystal struc-
tures of both FeðIIÞ-FeBMb and Fe(II)-I107E FeBMb show that
one O atom of E68 directly coordinates to FeB (Fig. 2). In syn-
thetic models of NOR, it has also been found that the presence of
a glutamic acid mimic significantly increases the stability of iron
binding to the FeB site (40). Furthermore, a theoretical study by
Blomberg et al. (58) showed that a model with an FeB coordi-
nated by three histidines, one glutamate, and one water molecule
provides an energetically feasible reaction mechanism of NO
reduction. However, the structural model of NOR constructed
recently by Reimann et al. (22) shows that the closest conserved
Glu (E267) still has its carboxylate O atom 7 Å away from FeB,
which suggests that Glu may not bind to FeB in native NOR. One
interesting finding from our study is that the Glu (E68) under-
went a significant conformational rearrangement in the presence
of another Glu (E107) (Fig. 2). Therefore, the FeBMb provides a
viable model of NOR that is consistent with Blomberg’s model,
but cannot rule out Reimann’s model due to possible conforma-
tion changes.
While the role of the first Glu is still uncertain until a 3D struc-
ture of NOR in its active form is available, the role of the second
Glu is even less defined. We address this question by introducing
a second Glu (E107) to FeBMb. The crystal structures shown in
Fig. 1 indicate that E107 interacts with a water molecule and
forms a hydrogen bonding network in both Fe(II)-I107E
FeBMb and Cu(II)-I107E FeBMb. Interestingly, although a
similar water molecule was observed in the active site of FeðIIÞ-
FeBMb (Fig. 2), activity assay data indicate that the presence of
E107 in Fe(II)-I107E FeBMb increases NOR activity by ∼100%
C
5 min
N
t l
A
5 min
B
2+
5 min
1 min
No metal
1 min
5 min
F
2+
No metal
1
i
5 min
+
No metal
2+
Zn
2+
5 min
Fe
2+
5 min
1 min
Fe
C
+
5 min
1 min
Cu
+
Z
2
Zn
2+
15 min
F
2+
Fe
15 min
5 min
C
+
Cu
+
15 min
5 min
Zn
2+
15 min
Fe2+
5
Cu
+
15 min
3000
3100
3200
3300
3400
Magnetic Field (Gauss)
3000
3100
3200
3300
3400
Magnetic Field (Gauss)
3000
3100
3200
3300
3400
Magnetic Field (Gauss)
Fig. 5.
EPR spectra of deoxy I107E FeBMb (0.5 mM) with no metal bound in
the presence of NO after 5 min (top line), with 2 eq Fe2þ (A), Cuþ (B), or Zn2þ
(C) incubated with excess NO (∼200 eq) for 1, 5, and 15 min. Spectra were
collected in 50 mM Bis-Tris pH 7.0 at 30 K, 0.2 mW power, and 9.05 GHz.
8584
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Lin et al.
(Fig. 4), suggesting that the second Glu may potentionally play a
role in providing one of the protons during reduction of NO to
N2O. Although free Glu outside a protein has a pKa ∼4.3, the stu-
dies of native NOR showed that the pKa of its Glu close to the
active site has a value of ∼6.6 (22, 25, 33). The hydrogen binding
network in our protein models may contribute to the fine-tuning
of the Glu pKa to be more neutral, similar to that in NOR. More-
over, it is interesting that Cu(I)-I107E FeBMb also shows NOR
activity, which provides an interesting protein model of HCOs
with NOR function (26–28), even though Glu residues are not
conserved in native HCOs.
Additionally, spectroelectrochemical studies showed that the
reduction potential of I107E FeBMb with no metal ion in the
FeB site is similar to that of FeBMb, but much lower than that
of CuBMb (77 mV) (54), which contains the same three His,
but no Glu in the metal-binding site above the heme. Since both
FeBMb and I107E FeBMb contain the V68E mutation that has
been shown to decrease the heme reduction potential of native
myoglobin from 59 mV to −137 mV (59), it is likely that the
introduction of a negatively charged Glu close to the heme group
is what is responsible for the dramatically lower heme redox
potential. A conserved glutamate, predicted to be located near
the catalytic heme b3 in NOR, was proposed to be responsible
for a ∼260-mV decrease in reduction potential (60 mV) in
comparison to the other two heme centers, heme b (345 mV)
and heme c (310 mV) (60). Our FeBMb and I107E FeBMb mod-
els mimic this feature of NOR. Notably, although introduction of
the first Glu (E68) lowered the heme potential by ∼110 mV (14),
introduction of the second Glu via the I107E mutation did not
result in a significant difference in the heme reduction potential,
suggesting that the effect of the two conserved Glu residues in
NOR on heme reduction potential is not additive, with the effects
highly dependent on the location of the Glu.
The Roles of Metal Ions. The roles of metal ions in NOR are an-
other important question as iron is found in the native FeB site
and HCO employs copper at the corresponding CuB site. With
different metal ions in the FeB site, the crystal structures clearly
show the heme and nonheme dinuclear center existing in differ-
ent local environments (Fig. 1). Although a similar hydrogen
bond network is formed in both Fe(II)-I107E FeBMb and Cu
(II)-I107E FeBMb, the conformation of E68 and E107 with re-
spect to the nonheme metal center and heme iron is different
from each other. Moreover, the coordination geometry differs
significantly with Zn2þ in the FeB site. A hydrogen bond is absent
from the Zn crystal structure, but both the O atoms of E68 act as
metal-binding ligands. These observations demonstrate that the
identity of the metal ion in the FeB site can tune the active site
through their interactions with the His and Glu ligands, resulting
in formation of different coordination geometries with different
hydrogen bonds.
In addition to structural fine-tuning, the metal ion at the FeB
site can also tune the heme iron reduction potential in I107E
FeBMb. Spectroelectrochemical studies showed that the binding
of Fe2þ or Zn2þ results in an increase in the heme reduction
potential by ∼70 mV and ∼30 mV, respectively (Fig. S1). In
the case of Cu(II)-I107E FeBMb, the crystal structure shows that
OE1 of E68 is closer to the heme iron (2.07 Å) (Fig. 1C) than its
metal-free form (2.15 Å) (Fig. 1A). The stronger interaction
from the negatively charged E68 could offset the effect of
positively charged Cu2þ binding, resulting in similar reduction
potentials
observed
for
Cu(II)-I107E
FeBMb
and
I107E
FeBMb.
In a previous study (61), EPR data showed that during NO
reduction, the binding of Cuþ to the CuB site of CuBMb can
weaken the proximal heme Fe-His bond, while complete cleavage
of the heme Fe-His bond occurred when Zn2þ was bound to
CuBMb-NO. In this study, we observed that a five-coordinate
heme-NO species was formed with Fe2þ, Cuþ, or Zn2þ bound
to the FeB site of I107E FeBMb (Fig. 5). Significantly, a five-
coordinate heme-NO species has also been observed for both
NOR (30, 31, 35) and the member of the HCO family with the
highest NO reduction activity, cytochrome cbb3 oxidases (26, 62).
However, this species was not observed for FeðIIÞ-FeBMb-NO
and CuðIÞ-FeBMb-NO, which lack the second Glu (E107). In
both these cases, the proximal heme Fe-His bond was only
weakened, as indicated by a decrease of the nine-line hyperfine
splitting signals in the EPR spectra (Fig. S3). These observations
suggest that formation of a five-coordinate heme-NO species may
play an important role in NOR reactivity.
Conclusions
We have successfully designed a structural and functional model
of NOR, by introducing a second glutamate in the vicinity of the
FeB site, named I107E FeBMb. This protein model mimics native
NOR more closely by bearing the structural feature of three his-
tidines and two glutamates in the FeB site, as predicted for native
NOR. We have demonstrated that the two glutamates can play
different roles in NO reduction activity; namely, one acts as a li-
gand to FeB (E68), and the other acts as a proton transfer group
(E107). Furthermore, by substituting different metal ions into the
nonheme metal site, we have demonstrated that FeB plays crucial
roles in fine-tuning the active site by donating electrons and by
mediating the formation of a five-coordinate heme-NO inter-
mediate during NO reduction. In the absence of a crystal struc-
ture for native NOR, this study offers an ideal protein model and
provides valuable structural as well as mechanistic information
for native NOR.
Materials and Methods
Protein Preparation. I107E FeBMb (swMb L29H/F43H/V68E/I107E) was con-
structed, expressed, and purified using the procedure described previously
(14). The purity and identity were confirmed by SDS-PAGE and electrospray
ionized MS: observed: 17; 392 1 Da; calculated: 17,391 Da.
EPR Spectroscopy. EPR spectra were recorded on a Bruker ESP 300 equipped
with an Oxford liquid helium cryostat and an ITC4 temperature controller.
The samples of met I107E FeBMb, Cu(II)-, or Zn(II)-I107E FeBMb were pre-
pared as described previously (14). The samples of NO-bound deoxy
I107E FeBMb, Cu(I)-, Fe(II)-, or Zn(II)-I107E FeBMb were prepared by inject-
ing 1 mL of purified NO gas into the EPR tube containing 300 μL of protein
(0.5 mM, 10% glycerol, in 50 mM Bis-Tris pH 7.0). The samples were then
flash frozen in liquid N2 after incubation for 1, 5, or 15 min. The molar
extinction coefficient of the Soret band of I107E FeBMb at 406 nm
(175 mM−1 · cm−1), calculated using the standard hemochromagen method
(63), was used to determine protein concentration. The metal sources of
Cu(I), Cu(II), Zn(II), and Fe(II) were ½ðCH3CNÞ4CuPF6, CuSO4, ZnSO4 · 7H2O,
and FeCl2, respectively.
Spectroelectrochemical Measurements. Protein reduction potentials were
measured using an optically transparent thin layer electrode as previously
described (64). The potential of the working electrode was applied in
the positive direction for deoxy I107E FeBMb with Fe2þ and in the negative
direction for metal free and with Cu2þ or Zn2þ. Other procedures are the
same as described previously (54).
X-Ray Crystallographic Studies. Fe(II)-I107E FeBMb was crystallized anaerobi-
cally in a glove box at room temperature using the conditions described for
FeðIIÞ-FeBMb (14). I107E FeBMb, Cu(II)-, or Zn(II)-I107E FeBMb were crystallized
aerobically. Diffraction-quality crystals were soaked in a cryoprotectant
solution of 30% PEG 400 and flash frozen in liquid nitrogen. Diffraction data
were collected at the Brookhaven National Lab Synchrotron Light Source
X12C beamline. The crystal structure was solved using the same method
as for FeðIIÞ-FeBMb (14).
NOR Activity Assay. NO reduction was monitored by GC/MS. The protein was
reduced to the deoxy form by excess dithionite that was removed with a
size-exclusion column (PD-10). Then 2 eq metal, Cu(I), Fe(II), or Zn(II), was
added to the protein solution (0.6 mM, 3 mL in 50 mM Bis-Tris buffer, pH
7.0). The samples were prepared anaerobically in a glove box. Purified NO
Lin et al.
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gas was injected into the head space of the reaction flask with the molar
ratio of NO∶protein ≈50∶1. Other procedures are the same as described
previously (14, 61).
ACKNOWLEDGMENTS. We thank Dr. Mark J. Nilges for help with EPR analysis,
and Furong Sun and Beth D. Eves for aiding in GC/MS data collection. This
work was supported by NIH Grant GM062211.
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|
3M3A
|
The roles of glutamates and metal ions in a rationally designed nitric oxide reductase based on myoglobin: Cu(II)-I107E FeBMb (Cu(II) binding to FeB site)
|
Roles of glutamates and metal ions in a rationally
designed nitric oxide reductase based on myoglobin
Ying-Wu Lina,2, Natasha Yeunga, Yi-Gui Gaob, Kyle D. Minerc, Shiliang Tiana, Howard Robinsond, and Yi Lua,c,1
aDepartment of Chemistry, bGeorge L. Clark X-Ray Facility and 3M Materials Laboratory, andcDepartment of Biochemistry, University of Illinois at
Urbana-Champaign, Urbana, IL 61801; and dDepartment of Biology, Brookhaven National Laboratory, Upton, NY 11973
Edited by Harry B. Gray, California Institute of Technology, Pasadena, CA, and approved April 1, 2010 (received for review January 14, 2010)
A structural and functional model of bacterial nitric oxide reductase
(NOR) has been designed by introducing two glutamates (Glu) and
three histidines (His) in sperm whale myoglobin. X-ray structural
data indicate that the three His and one Glu (V68E) residues bind
iron, mimicking the putative FeB site in NOR, while the second Glu
(I107E) interacts with a water molecule and forms a hydrogen bond-
ing network in the designed protein. Unlike the first Glu (V68E),
which lowered the heme reduction potential by ∼110 mV, the
second Glu has little effect on the heme potential, suggesting that
thenegativelychargedGluhasadifferentrolein redoxtuning.More
importantly, introducing the second Glu resulted in a ∼100% in-
crease in NOR activity, suggesting the importance of a hydrogen
bonding network in facilitating proton delivery during NOR reactiv-
ity. In addition, EPR and X-ray structural studies indicate that the
designed protein binds iron, copper, or zinc in the FeB site, each with
different effects on the structures and NOR activities, suggesting
that both redox activity and an intermediate five-coordinate
heme-NO species are important for high NOR activity. The designed
protein offers an excellent model for NOR and demonstrates the
power of using designed proteins as a simpler and more well-
defined system to address important chemical and biological issues.
biomimetic models ∣heme-copper oxidase ∣metalloprotein ∣protein design ∣
protein engineering
R
ational design of proteins that mimic both structure and func-
tion of more complex native enzymes has been a long sought-
after goal, as the process is an ultimate test of our knowledge and
an excellent means to develop advanced biocatalysts (1–3).
Although designed proteins that model the structure of native
enzymes have been known for a while (4–10), successful designs
of proteins that mimic both the structure and function of native
enzymes have been reported only recently (11–16). While being
able to design such functional proteins is laudable, the impact of
such an achievement would be greater if the designed proteins
can be used to address fundamental issues in chemistry and biol-
ogy that are difficult to tackle by other methods. One primary
example is the roles of conserved glutamates and metal ions in
bacterial nitric oxide reductase (NOR) (17–19).
NO is critical for all life (20). Bacterial denitrification is a cru-
cial part of the nitrogen cycle in nature that involves a four-step,
five-electron reduction of nitrate (NO3
−) to dinitrogen (N2)
(17, 19). Bacterial NOR is a membrane-bound protein that
catalyzes one step of this process, namely, the two-electron reduc-
tion of NO to N2O (17, 19). With no crystal or solution structure
available for bacterial NOR to date, sequence alignments and
homology modeling (21, 22) have indicated that NOR is structu-
rally homologous to the largest subunit (subunit I) of heme-
copper oxidases (HCOs) (23), enzymes that catalyze reduction
of O2 to water. The active sites of both NOR and HCO contain
a proximal histidine-coordinated heme and a distal three histi-
dine-coordinated metal center. However, the metal center in
HCOs is occupied by a copper (called CuB), whereas a nonheme
iron is present in NOR (called FeB) (23, 24). In addition, two
conserved glutamates, shown by modeling to be close to the
FeB site (21, 22), are found to be essential for NOR activity
(24, 25). Some members of HCOs such as cytochrome cbb3
oxidase display NOR activity (26–28), although the activity is
∼50-fold lower than native NOR (26). Therefore, it is important
to elucidate the structural features, specifically the roles of the
conserved glutamates close to the FeB site and metal ions (copper
vs. iron), responsible for the reduction of NO to N2O.
To address these issues, biochemical and biophysical studies of
native NOR and its variants have been carried out (24, 25, 29–37).
For example, Richardson and coworkers investigated the effects
of amino acid substitutions of the five conserved glutamates
(E122 and E125 presumed to face the periplasm and E198,
E202, and E267 located in the interior of the membrane, close
to the catalytic site) in the catalytic subunit of Paracoccus deni-
trificans, NorB. The E122A, E125A, E198A, and E267A variants
were inactive, indicating that these four glutamates are crucial for
NOR activity (24, 25, 32, 33). On the other hand, Reimann et al.
constructed a 3D model of NorB using homology modeling with
the structures of HCOs as templates and suggested a plausible
pathway consisting of these conserved glutamates for proton
delivery (22). Despite these successes, the roles of the conserved
glutamates and metal ions still remain to be fully elucidated,
partly because of the difficulty in obtaining native NOR in high
yield and the lack of a 3D structure. Even if these problems are
resolved, it is still difficult to replace iron in the native FeB site
with other metal ions, and spectroscopic studies of native NOR
are often complicated by the presence of other metal cofactors
(e.g., low-spin heme).
To overcome these limitations, a number of synthetic models of
NOR using small organic molecules as ligands, have been made
in which the nonheme FeB site can be replaced by a copper ion
(17, 38–45). In addition, since these model systems lack addi-
tional metal-binding sites, spectroscopic studies are often simpli-
fied. Therefore, studies of these synthetic models have offered
many insights. For example, Collman et al. showed that a fully
reduced heme/nonheme FeB compound can react with two
equivalents of NO leading to the formation of one equivalent
of N2O and a bis-ferric product (41). On the other hand, Karlin
and coworkers showed that a small heme/Cu complex can effi-
ciently lead to reductive coupling of NO to N2O (43). However,
it is also difficult to obtain the synthetic models in high yield due
to the multiple steps required in chemical synthesis. Because of
this limitation, no synthetic NOR model containing the two key
conserved Glu residues (E198 and E267 in NOR) has been
Author contributions: Y.-W.L., N.Y., and Y.L. designed research; Y.-W.L., N.Y., Y.-G.G.,
K.D.M., and S.T. performed research; Y.-W.L., N.Y., Y.-G.G., K.D.M., H.R., and Y.L. analyzed
data; and Y.-W.L. and Y.L. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The atomic coordinates and structure factors have been deposited in the
Protein Data Bank, www.pdb.org (PDB ID codes 3M38, 3M39, 3M3A, and 3M3B).
1To whom correspondence should be addressed. E-mail: yi-lu@illinois.edu.
2Present address: School of Chemistry and Chemical Engineering, University of South
China, Hengyang 421001, China.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1000526107/-/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.1000526107
PNAS ∣May 11, 2010 ∣vol. 107 ∣no. 19 ∣8581–8586
BIOCHEMISTRY
CHEMISTRY
reported. It is also difficult to substitute different metal ions in
the same metal-binding site without perturbing the site geometry
and distances to the heme iron, as most ligands are not as rigid as
those in native enzymes and different metal ions have different
geometric and ligand donor set preferences.
We have recently designed a structural and functional protein
model of bacterial NOR by engineering three histidines and one
glutamate into the distal pocket of sperm whale myoglobin
(swMb, L29H, F43H, H64, and V68E, named FeBMb) (14). Like
synthetic models, this “bottom-up” approach complements the
“top-down”approachofthestudyofnativeNORinthatitprovides
insights into whether certain “necessary” structural elements are
enough to impart enzyme function. Thanks in part to recent
advances in computational, molecular, and structural biology,
the designed myoglobin protein model is much easier to synthesize
and to crystallize than either native NOR or synthetic models.
Since myoglobin has often been used for the development and
calibration of numerous spectroscopic techniques (46–48), it is
an ideal choice for spectroscopic studies. More importantly, the
rigid protein network allows precise placement of either glutamate
or metal ions in myoglobin to address their roles in NOR activity.
Toward this goal, we have demonstrated that both the histidines
and one of the glutamates are essential for iron binding and NO
reduction activity (14). However, the role of the second Glu close
to the FeB site and the role of different metal ions in the FeB site
have not been addressed.
To address these important issues and to design even closer
protein models of NOR, we introduced herein the second Glu
to the second coordination sphere of the FeB site by mutating
an Ile to a Glu (named I107E FeBMb). We show that the second
Glu results in a ∼100% increase in NOR activity through hydro-
gen bonding interactions and that the two glutamates have
dramatically different effects on the heme reduction potential.
Additionally, by comparing the EPR, electrochemistry, X-ray
structures, and NOR activity of iron, copper, and zinc derivatives
of the designed protein, we have obtained deeper insights into the
roles of metal ions in NOR.
Results
Structure and Function of Fe(II)-I107E FeBMb. The X-ray crystal struc-
tures of heme-containing I107E FeBMb without metal ion in the
FeB site and with Fe2þ in the FeB site are solved at 1.42-Å and
1.65-Å resolution, respectively (Fig. 1 A and B and Table S1). In
the absence of metal ions in the FeB site, the structure shows a
water molecule in the FeB site, which forms hydrogen bonds with
NE2 atoms of all three His residues, both OE1 and OE2 atoms of
E68, and the OE2 atom of E107 (Fig. 1A). Upon binding Fe2þ,
the Fe(II)-I107E FeBMb structure shows that Fe2þ is coordinated
by three His, the OE2 atom of E68, and one water molecule.
Notably, a water molecule bridges Fe2þ in the FeB site and
the second glutamate (E107) with a distance of 2.32 Å to the
OE2 atom of E107 (Fig. 1B).
To probe the conformational changes of introducing the
second Glu (E107), we performed a structural alignment of Fe
(II)-I107E FeBMb and the previously reported FeðIIÞ-FeBMb
(14). The comparison, shown in Fig. 2, indicates that both the
polypeptide chain and the active site overlap well with each other.
In addition, the two nonheme irons are located at similar posi-
tions with a 0.36-Å separation from each other. In contrast,
E68 underwent a significant conformational rearrangement in
the presence of E107. These observations suggest that the active
site of FeBMb can be tuned by the formation of an extended
hydrogen bonding network, resulting from the introduction of
a second glutamate residue.
The binding of Fe2þ to deoxy I107E FeBMb was further mon-
itored by EPR (Fig. 3A). Since deoxy myoglobin contains Fe(II)
heme that exhibits no EPR signals in X-band EPR (14), we added
blue copper Cu(II)-azurin (49), a redox partner of native NOR
(19), to oxidize both the reduced heme and nonheme irons in Fe
(II)-I107E FeBMb to EPR-active Fe(III). Upon addition of Cu
(II)-azurin, the oxidation of deoxy I107E FeBMb resulted in
EPR signals at g ¼ 6.12 and 5.56, typical of a high-spin heme-
Fe(III). Upon addition of Fe2þ, however, a decrease of the
heme-Fe(III) EPR signals was observed, indicating that the
Fe2þ, when bound to the FeB site and oxidized by Cu(II)-azurin,
is spin-coupled to heme-Fe(III). Such a spin coupling mimics
that in NOR (35, 50–53), suggesting that I107E FeBMb models
NOR closely, at least in this respect.
To probe the role of the second Glu (E107) in NO reduction
activity, we measured the yield of N2O production by Fe(II)-
I107E FeBMb with excess NO under one turnover conditions.
We monitored N2O formation in the headspace of the solution
using GC/MS and compared this result to that of FeðIIÞ-FeBMb,
which lacks the second Glu (Fig. 4). Remarkably, Fe(II)-I107E
FeBMb
displays higher
activity than
FeðIIÞ-FeBMb.
After
∼20 hr, ∼24% N2O was produced by Fe(II)-I107E FeBMb, in
H29
A
H29
B
H64
E68
E107
3.04
2.81
3.02
2.20
2.24
2.24
2.12
2.21
H64
E68
E107
3.03
H43
2.15
3.41
3.16
2.62
2.32
H43
2.26
H93
H93
H29
C
H29
D
2.04
2.03
H64
E68
E107
2.09
2.10
2.91
2.21
H64
E68
E107
2.26
2.29
2.10
2.18
2.10
4.47
H43
2.07
3.04
H43
2.68
H93
H93
Fig. 1.
Crystal structures of I107E FeBMb (A) (PDB ID code 3M38), Fe(II)-I107E
FeBMb (B) (PDB ID code 3M39), Cu(II)-I107E FeBMb (C) (PDB ID code 3M3A),
and Zn(II)-I107E FeBMb (D) (PDB ID code 3M3B). Water molecules, Fe(II),
Cu(II), and Zn(II) are represented by red, green, orange, and gray spheres,
respectively.
Fig. 2.
Overlay of Fe(II)-I107E FeBMb (cyan) (PDB ID code 3M39) with
FeðIIÞ-FeBMb (orange) (PDB ID code 3K9Z).
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Lin et al.
contrast to ∼10% yield for FeðIIÞ-FeBMb, strongly indicating that
the second Glu plays an important role in NO reduction, likely
facilitating proton uptake during NO reduction.
Other Metal Ions Binding to I107E FeBMb. To find out if the resting
state of the protein, i.e., oxidized or met I107E FeBMb, can bind
other metal ions, Cu2þ or Zn2þ was titrated into met I107E
FeBMb and monitored by EPR spectroscopy (Fig. 3 B and C).
In the absence of metal ions, met I107E FeBMb exhibited
high-spin heme signals at g ¼ 6.03, 5.08, and 1.98 (Fig. 3B, black
line). Upon addition of 2 eq of Cu2þ, the signals at g ¼ 6.03 and
5.08 decreased and a broad peak around g ¼ 2.95 increased,
probably due to spin coupling between heme-Fe(III) and Cu2þ
in the FeB site. In contrast, addition of Zn2þ, a metal ion with
no unpaired electrons [i.e., incapable of spin coupling to heme-
Fe(III)], produced an increase in the high-spin heme signals at
g ¼ 5.88 and 5.60 (Fig. 3C), indicating that the interaction be-
tween E68 and heme iron was weakened after metal binding.
The X-ray crystal structures of I107E FeBMb with Cu2þ or
Zn2þ in the FeB site were solved at 1.37-Å and 1.60-Å resolution,
respectively (Fig. 1 C and D and Table S1). Compared to Fe(II)-
I107E FeBMb (Fig. 1B), a similar binding site was observed
for Cu(II)-I107E FeBMb (Fig. 1C), where H29, H43, and H64
coordinate to Cu2þ with distances of 2.09, 2.10, and 2.04 Å,
respectively, slightly shorter than the corresponding distances
in the Fe2þ structure. In comparison to Fe(II)-I107E FeBMb,
the water bridging the Cu2þ and the second Glu (E107) is shifted
toward Cu2þ in the FeB site (2.03 Å) with respect to E107
(3.04 Å). Interestingly, this bridging water molecule was not
observed in Zn(II)-I107E FeBMb (Fig. 1D), but the two O atoms
of E68 coordinate to Zn2þ with similar distances (2.26 Å for OE1
and 2.29 Å for OE2). The longer distance between OE1 of E68
B
6.12
I107E FeBMb + Azurin
I107E Fe Mb + 0 5 eq Fe
2+ + Azurin
A
5.56
I107E FeBMb + 0.5 eq Fe
+ Azurin
I107E FeBMb + 1.0 eq Fe
2+ + Azurin
I107E FeBMb + 2.0 eq Fe
2+ + Azurin
1.98
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
5.08
5.38
Magnetic Field (Gauss)
5.88
I107E FeBMb
I107E FeBMb + 0.5 eq Zn
2+
I107E Fe Mb
1 0 eq Zn
2+
C
6.03
I107E FeBMb
I107E FeBMb + 0.5 eq Cu
2+
I107E Fe Mb + 1 0 eq Cu
2+
I107E FeBMb + 1.0 eq Zn
I107E FeBMb + 2.0 eq Zn
2+
I107E FeBMb + 1.0 eq Cu
I107E FeBMb + 2.0 eq Cu
2+
1.98
~2.95
1.98
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
5.60
Magnetic Field (Gauss)
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
5.08
Magnetic Field (Gauss)
Fig. 3.
EPR spectra of deoxy I107E FeBMb (0.5 mM protein in 50 mM Bis-Tris, pH 7.0) with increasing concentrations of Fe2þ in the presence of wild-type
Cu(II)-azurin (A), and oxidized I107E FeBMb with Cu2þ (B) or Zn2þ (C). Spectra were collected at 4 K, 5 mW power, and 9.05 GHz.
30
25
Fe(II)-I107E FeBMb
Fe(II)-FeBMb
15
20
10
%)
oduction (%
N2O pro
0
5
0
4
8
12
16
20
Incubation time (hr)
Fig. 4.
Time-dependent N2O production by Fe(II)-I107E FeBMb (▴) and
FeðIIÞ-FeBMb (●) with ∼50 eq. NO under single turnover conditions. The yield
was determined by a comparison of the ratio of NO∶N2O peaks from the
GC/MS chromatograms.
Lin et al.
PNAS
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and heme iron in the Zn-bound structure (2.68 Å) in comparison
to the Cu- and Fe-bound structures, is likely the result of a weaker
interaction, which is also supported by an observed increase of
the high-spin heme signals in the EPR spectra upon Zn2þ binding
(Fig. 3C). These results suggest I107E FeBMb is capable of incor-
porating different metal ions into its designed FeB site, offering
an excellent opportunity to compare the role of these metal ions
in the same protein scaffold.
Effect of Glutamates and Metal Ions on the Redox Potential of I107E
FeBMb. Since EPR and X-ray structural studies indicate metal
binding to I107E FeBMb, we used spectroelectrochemistry to
measure the effects of glutamates and metal ions on the heme
reduction potential. When there is no metal ion in the FeB site,
the I107E FeBMb displays a reduction potential of −134 3 mV
vs. the normal hydrogen electrode (NHE) (Fig. S1A), similar to
that of FeBMb (−158 4 mV) without the I107E mutation (14).
In the presence of Cu2þ, I107E FeBMb has a reduction potential
(−137 2 mV) (Fig. S1B) almost identical to that of the same
protein in the absence of metal ions in the FeB site, indicating
that copper binding to the FeB site has little effect on the reduction
potential of the heme iron. This observation is similar to that
observed for Cu2þ binding to CuBMb (CuðIIÞ-CuBMb, 80 mV
vs. CuBMb, 77 mV) (54). On the other hand, the presence of
Fe2þ and Zn2þ increased the reduction potential of I107E
FeBMb from −134 3 mV to −64 3 mV vs. NHE (Fig. S1C)
and −105 2 mV vs. NHE (Fig. S1D), respectively. The different
effects of Cu2þ, Fe2þ, and Zn2þ on the reduction potential of
I107E-FeBMb indicate that these metal ions in the FeB site may
play different roles through different coordination properties.
NOR Activity of I107E FeBMb in the Presence of Different Metal Ions.
The NO reduction activity of I107E FeBMb in the presence of
Fe2þ, Cuþ, or Zn2þ was monitored by GC/MS under single turn-
over conditions. When Fe(II)-I107E FeBMb was exposed to
excess NO, N2O could be observed to form with increased yield
over time (Fig. S2). Similarly, N2O formation was observed for
Cu(I)-I107E FeBMb, indicating that Fe or Cu binding to the
FeB site results in comparable NOR activities. It should be noted
that because of the high solubility of N2O (∼25 mM in water at
room temperature), GC/MS cannot be used to quantify the rates
of NO reduction under these conditions. In contrast, no N2O
formation was observed with redox inactive Zn2þ, which demon-
strates that redox active Fe2þ or Cuþ in the FeB site plays a
crucial role in NO reduction.
To gain deeper insight into the process of NO reduction, EPR
studies were further performed to monitor the initial process of
NO reduction. In the absence of metal ions, the EPR spectrum of
ferrous I107E FeBMb-NO shows hyperfine splitting resulting
from bound NO and the proximal histidine, indicating the forma-
tion of a six-coordinate ferrous heme-NO species (Fig. 5, top
line). After incubation of Fe(II)-I107E FeBMb with excess
NO, a distinct three-line hyperfine structure appears at 15 min
(Fig. 5A), suggesting the formation of a five-coordinate ferrous
heme-NO species as a result of cleavage of the proximal
His-Fe heme bond (55). A three-line hyperfine structure was also
observed for Cuþ and Zn2þ, except that the signal intensity is low-
er with Cu(I)-I107E FeBMb-NO (Fig. 5B) and more pronounced
in Zn(II)-I107E FeBMb-NO (Fig. 5C). The lower intensity of
the three-line hyperfine structure for Cu(I)-I107E FeBMb-NO
suggests the major species formed is a six-coordinate ferric
heme-NO complex, which is EPR silent (41). These differences
further suggest that the metal ion in the FeB site plays a key role
in formation of the intermediates, thereby tuning NOR activity.
Discussion
Using Rationally Designed Proteins to Address Important Issues in
Chemistry and Biology. Important issues such as the roles of the
conserved glutamates and nonheme FeB in NOR have been
previously addressed using biochemical and biophysical studies
or biomimetic modeling (24, 25, 27–37, 45, 56, 57). As a comple-
mentary approach, rational protein design, using small, easy-to-
produce and well-characterized proteins such as myoglobin,
offers a powerful method with which to gain insights into more
complex native enzymes such as NOR (14). Similar to synthetic
models (41, 43), the metal ion at the putative FeB site in the
protein model can be substituted freely. Better yet, Glu residues
can be placed at precise locations in the protein, including the
secondary coordination sphere, due to its rigid network. By care-
fully choosing a suitable protein template, rational protein design
could be generally applied to address other important issues in
chemistry and biology.
The Roles of Glutamates. Although two conserved glutamates
(E198 and E267) are known to be crucial for NOR activity
(24, 25), their roles are not well defined (18, 19). In a previous
study (14), we demonstrated that one Glu, E68, is important for
both iron binding and NOR activity of FeBMb. The crystal struc-
tures of both FeðIIÞ-FeBMb and Fe(II)-I107E FeBMb show that
one O atom of E68 directly coordinates to FeB (Fig. 2). In syn-
thetic models of NOR, it has also been found that the presence of
a glutamic acid mimic significantly increases the stability of iron
binding to the FeB site (40). Furthermore, a theoretical study by
Blomberg et al. (58) showed that a model with an FeB coordi-
nated by three histidines, one glutamate, and one water molecule
provides an energetically feasible reaction mechanism of NO
reduction. However, the structural model of NOR constructed
recently by Reimann et al. (22) shows that the closest conserved
Glu (E267) still has its carboxylate O atom 7 Å away from FeB,
which suggests that Glu may not bind to FeB in native NOR. One
interesting finding from our study is that the Glu (E68) under-
went a significant conformational rearrangement in the presence
of another Glu (E107) (Fig. 2). Therefore, the FeBMb provides a
viable model of NOR that is consistent with Blomberg’s model,
but cannot rule out Reimann’s model due to possible conforma-
tion changes.
While the role of the first Glu is still uncertain until a 3D struc-
ture of NOR in its active form is available, the role of the second
Glu is even less defined. We address this question by introducing
a second Glu (E107) to FeBMb. The crystal structures shown in
Fig. 1 indicate that E107 interacts with a water molecule and
forms a hydrogen bonding network in both Fe(II)-I107E
FeBMb and Cu(II)-I107E FeBMb. Interestingly, although a
similar water molecule was observed in the active site of FeðIIÞ-
FeBMb (Fig. 2), activity assay data indicate that the presence of
E107 in Fe(II)-I107E FeBMb increases NOR activity by ∼100%
C
5 min
N
t l
A
5 min
B
2+
5 min
1 min
No metal
1 min
5 min
F
2+
No metal
1
i
5 min
+
No metal
2+
Zn
2+
5 min
Fe
2+
5 min
1 min
Fe
C
+
5 min
1 min
Cu
+
Z
2
Zn
2+
15 min
F
2+
Fe
15 min
5 min
C
+
Cu
+
15 min
5 min
Zn
2+
15 min
Fe2+
5
Cu
+
15 min
3000
3100
3200
3300
3400
Magnetic Field (Gauss)
3000
3100
3200
3300
3400
Magnetic Field (Gauss)
3000
3100
3200
3300
3400
Magnetic Field (Gauss)
Fig. 5.
EPR spectra of deoxy I107E FeBMb (0.5 mM) with no metal bound in
the presence of NO after 5 min (top line), with 2 eq Fe2þ (A), Cuþ (B), or Zn2þ
(C) incubated with excess NO (∼200 eq) for 1, 5, and 15 min. Spectra were
collected in 50 mM Bis-Tris pH 7.0 at 30 K, 0.2 mW power, and 9.05 GHz.
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Lin et al.
(Fig. 4), suggesting that the second Glu may potentionally play a
role in providing one of the protons during reduction of NO to
N2O. Although free Glu outside a protein has a pKa ∼4.3, the stu-
dies of native NOR showed that the pKa of its Glu close to the
active site has a value of ∼6.6 (22, 25, 33). The hydrogen binding
network in our protein models may contribute to the fine-tuning
of the Glu pKa to be more neutral, similar to that in NOR. More-
over, it is interesting that Cu(I)-I107E FeBMb also shows NOR
activity, which provides an interesting protein model of HCOs
with NOR function (26–28), even though Glu residues are not
conserved in native HCOs.
Additionally, spectroelectrochemical studies showed that the
reduction potential of I107E FeBMb with no metal ion in the
FeB site is similar to that of FeBMb, but much lower than that
of CuBMb (77 mV) (54), which contains the same three His,
but no Glu in the metal-binding site above the heme. Since both
FeBMb and I107E FeBMb contain the V68E mutation that has
been shown to decrease the heme reduction potential of native
myoglobin from 59 mV to −137 mV (59), it is likely that the
introduction of a negatively charged Glu close to the heme group
is what is responsible for the dramatically lower heme redox
potential. A conserved glutamate, predicted to be located near
the catalytic heme b3 in NOR, was proposed to be responsible
for a ∼260-mV decrease in reduction potential (60 mV) in
comparison to the other two heme centers, heme b (345 mV)
and heme c (310 mV) (60). Our FeBMb and I107E FeBMb mod-
els mimic this feature of NOR. Notably, although introduction of
the first Glu (E68) lowered the heme potential by ∼110 mV (14),
introduction of the second Glu via the I107E mutation did not
result in a significant difference in the heme reduction potential,
suggesting that the effect of the two conserved Glu residues in
NOR on heme reduction potential is not additive, with the effects
highly dependent on the location of the Glu.
The Roles of Metal Ions. The roles of metal ions in NOR are an-
other important question as iron is found in the native FeB site
and HCO employs copper at the corresponding CuB site. With
different metal ions in the FeB site, the crystal structures clearly
show the heme and nonheme dinuclear center existing in differ-
ent local environments (Fig. 1). Although a similar hydrogen
bond network is formed in both Fe(II)-I107E FeBMb and Cu
(II)-I107E FeBMb, the conformation of E68 and E107 with re-
spect to the nonheme metal center and heme iron is different
from each other. Moreover, the coordination geometry differs
significantly with Zn2þ in the FeB site. A hydrogen bond is absent
from the Zn crystal structure, but both the O atoms of E68 act as
metal-binding ligands. These observations demonstrate that the
identity of the metal ion in the FeB site can tune the active site
through their interactions with the His and Glu ligands, resulting
in formation of different coordination geometries with different
hydrogen bonds.
In addition to structural fine-tuning, the metal ion at the FeB
site can also tune the heme iron reduction potential in I107E
FeBMb. Spectroelectrochemical studies showed that the binding
of Fe2þ or Zn2þ results in an increase in the heme reduction
potential by ∼70 mV and ∼30 mV, respectively (Fig. S1). In
the case of Cu(II)-I107E FeBMb, the crystal structure shows that
OE1 of E68 is closer to the heme iron (2.07 Å) (Fig. 1C) than its
metal-free form (2.15 Å) (Fig. 1A). The stronger interaction
from the negatively charged E68 could offset the effect of
positively charged Cu2þ binding, resulting in similar reduction
potentials
observed
for
Cu(II)-I107E
FeBMb
and
I107E
FeBMb.
In a previous study (61), EPR data showed that during NO
reduction, the binding of Cuþ to the CuB site of CuBMb can
weaken the proximal heme Fe-His bond, while complete cleavage
of the heme Fe-His bond occurred when Zn2þ was bound to
CuBMb-NO. In this study, we observed that a five-coordinate
heme-NO species was formed with Fe2þ, Cuþ, or Zn2þ bound
to the FeB site of I107E FeBMb (Fig. 5). Significantly, a five-
coordinate heme-NO species has also been observed for both
NOR (30, 31, 35) and the member of the HCO family with the
highest NO reduction activity, cytochrome cbb3 oxidases (26, 62).
However, this species was not observed for FeðIIÞ-FeBMb-NO
and CuðIÞ-FeBMb-NO, which lack the second Glu (E107). In
both these cases, the proximal heme Fe-His bond was only
weakened, as indicated by a decrease of the nine-line hyperfine
splitting signals in the EPR spectra (Fig. S3). These observations
suggest that formation of a five-coordinate heme-NO species may
play an important role in NOR reactivity.
Conclusions
We have successfully designed a structural and functional model
of NOR, by introducing a second glutamate in the vicinity of the
FeB site, named I107E FeBMb. This protein model mimics native
NOR more closely by bearing the structural feature of three his-
tidines and two glutamates in the FeB site, as predicted for native
NOR. We have demonstrated that the two glutamates can play
different roles in NO reduction activity; namely, one acts as a li-
gand to FeB (E68), and the other acts as a proton transfer group
(E107). Furthermore, by substituting different metal ions into the
nonheme metal site, we have demonstrated that FeB plays crucial
roles in fine-tuning the active site by donating electrons and by
mediating the formation of a five-coordinate heme-NO inter-
mediate during NO reduction. In the absence of a crystal struc-
ture for native NOR, this study offers an ideal protein model and
provides valuable structural as well as mechanistic information
for native NOR.
Materials and Methods
Protein Preparation. I107E FeBMb (swMb L29H/F43H/V68E/I107E) was con-
structed, expressed, and purified using the procedure described previously
(14). The purity and identity were confirmed by SDS-PAGE and electrospray
ionized MS: observed: 17; 392 1 Da; calculated: 17,391 Da.
EPR Spectroscopy. EPR spectra were recorded on a Bruker ESP 300 equipped
with an Oxford liquid helium cryostat and an ITC4 temperature controller.
The samples of met I107E FeBMb, Cu(II)-, or Zn(II)-I107E FeBMb were pre-
pared as described previously (14). The samples of NO-bound deoxy
I107E FeBMb, Cu(I)-, Fe(II)-, or Zn(II)-I107E FeBMb were prepared by inject-
ing 1 mL of purified NO gas into the EPR tube containing 300 μL of protein
(0.5 mM, 10% glycerol, in 50 mM Bis-Tris pH 7.0). The samples were then
flash frozen in liquid N2 after incubation for 1, 5, or 15 min. The molar
extinction coefficient of the Soret band of I107E FeBMb at 406 nm
(175 mM−1 · cm−1), calculated using the standard hemochromagen method
(63), was used to determine protein concentration. The metal sources of
Cu(I), Cu(II), Zn(II), and Fe(II) were ½ðCH3CNÞ4CuPF6, CuSO4, ZnSO4 · 7H2O,
and FeCl2, respectively.
Spectroelectrochemical Measurements. Protein reduction potentials were
measured using an optically transparent thin layer electrode as previously
described (64). The potential of the working electrode was applied in
the positive direction for deoxy I107E FeBMb with Fe2þ and in the negative
direction for metal free and with Cu2þ or Zn2þ. Other procedures are the
same as described previously (54).
X-Ray Crystallographic Studies. Fe(II)-I107E FeBMb was crystallized anaerobi-
cally in a glove box at room temperature using the conditions described for
FeðIIÞ-FeBMb (14). I107E FeBMb, Cu(II)-, or Zn(II)-I107E FeBMb were crystallized
aerobically. Diffraction-quality crystals were soaked in a cryoprotectant
solution of 30% PEG 400 and flash frozen in liquid nitrogen. Diffraction data
were collected at the Brookhaven National Lab Synchrotron Light Source
X12C beamline. The crystal structure was solved using the same method
as for FeðIIÞ-FeBMb (14).
NOR Activity Assay. NO reduction was monitored by GC/MS. The protein was
reduced to the deoxy form by excess dithionite that was removed with a
size-exclusion column (PD-10). Then 2 eq metal, Cu(I), Fe(II), or Zn(II), was
added to the protein solution (0.6 mM, 3 mL in 50 mM Bis-Tris buffer, pH
7.0). The samples were prepared anaerobically in a glove box. Purified NO
Lin et al.
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gas was injected into the head space of the reaction flask with the molar
ratio of NO∶protein ≈50∶1. Other procedures are the same as described
previously (14, 61).
ACKNOWLEDGMENTS. We thank Dr. Mark J. Nilges for help with EPR analysis,
and Furong Sun and Beth D. Eves for aiding in GC/MS data collection. This
work was supported by NIH Grant GM062211.
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3M3B
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The roles of glutamates and metal ions in a rationally designed nitric oxide reductase based on myoglobin: Zn(II)-I107E FeBMb (Zn(II) binding to FeB site)
|
Roles of glutamates and metal ions in a rationally
designed nitric oxide reductase based on myoglobin
Ying-Wu Lina,2, Natasha Yeunga, Yi-Gui Gaob, Kyle D. Minerc, Shiliang Tiana, Howard Robinsond, and Yi Lua,c,1
aDepartment of Chemistry, bGeorge L. Clark X-Ray Facility and 3M Materials Laboratory, andcDepartment of Biochemistry, University of Illinois at
Urbana-Champaign, Urbana, IL 61801; and dDepartment of Biology, Brookhaven National Laboratory, Upton, NY 11973
Edited by Harry B. Gray, California Institute of Technology, Pasadena, CA, and approved April 1, 2010 (received for review January 14, 2010)
A structural and functional model of bacterial nitric oxide reductase
(NOR) has been designed by introducing two glutamates (Glu) and
three histidines (His) in sperm whale myoglobin. X-ray structural
data indicate that the three His and one Glu (V68E) residues bind
iron, mimicking the putative FeB site in NOR, while the second Glu
(I107E) interacts with a water molecule and forms a hydrogen bond-
ing network in the designed protein. Unlike the first Glu (V68E),
which lowered the heme reduction potential by ∼110 mV, the
second Glu has little effect on the heme potential, suggesting that
thenegativelychargedGluhasadifferentrolein redoxtuning.More
importantly, introducing the second Glu resulted in a ∼100% in-
crease in NOR activity, suggesting the importance of a hydrogen
bonding network in facilitating proton delivery during NOR reactiv-
ity. In addition, EPR and X-ray structural studies indicate that the
designed protein binds iron, copper, or zinc in the FeB site, each with
different effects on the structures and NOR activities, suggesting
that both redox activity and an intermediate five-coordinate
heme-NO species are important for high NOR activity. The designed
protein offers an excellent model for NOR and demonstrates the
power of using designed proteins as a simpler and more well-
defined system to address important chemical and biological issues.
biomimetic models ∣heme-copper oxidase ∣metalloprotein ∣protein design ∣
protein engineering
R
ational design of proteins that mimic both structure and func-
tion of more complex native enzymes has been a long sought-
after goal, as the process is an ultimate test of our knowledge and
an excellent means to develop advanced biocatalysts (1–3).
Although designed proteins that model the structure of native
enzymes have been known for a while (4–10), successful designs
of proteins that mimic both the structure and function of native
enzymes have been reported only recently (11–16). While being
able to design such functional proteins is laudable, the impact of
such an achievement would be greater if the designed proteins
can be used to address fundamental issues in chemistry and biol-
ogy that are difficult to tackle by other methods. One primary
example is the roles of conserved glutamates and metal ions in
bacterial nitric oxide reductase (NOR) (17–19).
NO is critical for all life (20). Bacterial denitrification is a cru-
cial part of the nitrogen cycle in nature that involves a four-step,
five-electron reduction of nitrate (NO3
−) to dinitrogen (N2)
(17, 19). Bacterial NOR is a membrane-bound protein that
catalyzes one step of this process, namely, the two-electron reduc-
tion of NO to N2O (17, 19). With no crystal or solution structure
available for bacterial NOR to date, sequence alignments and
homology modeling (21, 22) have indicated that NOR is structu-
rally homologous to the largest subunit (subunit I) of heme-
copper oxidases (HCOs) (23), enzymes that catalyze reduction
of O2 to water. The active sites of both NOR and HCO contain
a proximal histidine-coordinated heme and a distal three histi-
dine-coordinated metal center. However, the metal center in
HCOs is occupied by a copper (called CuB), whereas a nonheme
iron is present in NOR (called FeB) (23, 24). In addition, two
conserved glutamates, shown by modeling to be close to the
FeB site (21, 22), are found to be essential for NOR activity
(24, 25). Some members of HCOs such as cytochrome cbb3
oxidase display NOR activity (26–28), although the activity is
∼50-fold lower than native NOR (26). Therefore, it is important
to elucidate the structural features, specifically the roles of the
conserved glutamates close to the FeB site and metal ions (copper
vs. iron), responsible for the reduction of NO to N2O.
To address these issues, biochemical and biophysical studies of
native NOR and its variants have been carried out (24, 25, 29–37).
For example, Richardson and coworkers investigated the effects
of amino acid substitutions of the five conserved glutamates
(E122 and E125 presumed to face the periplasm and E198,
E202, and E267 located in the interior of the membrane, close
to the catalytic site) in the catalytic subunit of Paracoccus deni-
trificans, NorB. The E122A, E125A, E198A, and E267A variants
were inactive, indicating that these four glutamates are crucial for
NOR activity (24, 25, 32, 33). On the other hand, Reimann et al.
constructed a 3D model of NorB using homology modeling with
the structures of HCOs as templates and suggested a plausible
pathway consisting of these conserved glutamates for proton
delivery (22). Despite these successes, the roles of the conserved
glutamates and metal ions still remain to be fully elucidated,
partly because of the difficulty in obtaining native NOR in high
yield and the lack of a 3D structure. Even if these problems are
resolved, it is still difficult to replace iron in the native FeB site
with other metal ions, and spectroscopic studies of native NOR
are often complicated by the presence of other metal cofactors
(e.g., low-spin heme).
To overcome these limitations, a number of synthetic models of
NOR using small organic molecules as ligands, have been made
in which the nonheme FeB site can be replaced by a copper ion
(17, 38–45). In addition, since these model systems lack addi-
tional metal-binding sites, spectroscopic studies are often simpli-
fied. Therefore, studies of these synthetic models have offered
many insights. For example, Collman et al. showed that a fully
reduced heme/nonheme FeB compound can react with two
equivalents of NO leading to the formation of one equivalent
of N2O and a bis-ferric product (41). On the other hand, Karlin
and coworkers showed that a small heme/Cu complex can effi-
ciently lead to reductive coupling of NO to N2O (43). However,
it is also difficult to obtain the synthetic models in high yield due
to the multiple steps required in chemical synthesis. Because of
this limitation, no synthetic NOR model containing the two key
conserved Glu residues (E198 and E267 in NOR) has been
Author contributions: Y.-W.L., N.Y., and Y.L. designed research; Y.-W.L., N.Y., Y.-G.G.,
K.D.M., and S.T. performed research; Y.-W.L., N.Y., Y.-G.G., K.D.M., H.R., and Y.L. analyzed
data; and Y.-W.L. and Y.L. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The atomic coordinates and structure factors have been deposited in the
Protein Data Bank, www.pdb.org (PDB ID codes 3M38, 3M39, 3M3A, and 3M3B).
1To whom correspondence should be addressed. E-mail: yi-lu@illinois.edu.
2Present address: School of Chemistry and Chemical Engineering, University of South
China, Hengyang 421001, China.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1000526107/-/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.1000526107
PNAS ∣May 11, 2010 ∣vol. 107 ∣no. 19 ∣8581–8586
BIOCHEMISTRY
CHEMISTRY
reported. It is also difficult to substitute different metal ions in
the same metal-binding site without perturbing the site geometry
and distances to the heme iron, as most ligands are not as rigid as
those in native enzymes and different metal ions have different
geometric and ligand donor set preferences.
We have recently designed a structural and functional protein
model of bacterial NOR by engineering three histidines and one
glutamate into the distal pocket of sperm whale myoglobin
(swMb, L29H, F43H, H64, and V68E, named FeBMb) (14). Like
synthetic models, this “bottom-up” approach complements the
“top-down”approachofthestudyofnativeNORinthatitprovides
insights into whether certain “necessary” structural elements are
enough to impart enzyme function. Thanks in part to recent
advances in computational, molecular, and structural biology,
the designed myoglobin protein model is much easier to synthesize
and to crystallize than either native NOR or synthetic models.
Since myoglobin has often been used for the development and
calibration of numerous spectroscopic techniques (46–48), it is
an ideal choice for spectroscopic studies. More importantly, the
rigid protein network allows precise placement of either glutamate
or metal ions in myoglobin to address their roles in NOR activity.
Toward this goal, we have demonstrated that both the histidines
and one of the glutamates are essential for iron binding and NO
reduction activity (14). However, the role of the second Glu close
to the FeB site and the role of different metal ions in the FeB site
have not been addressed.
To address these important issues and to design even closer
protein models of NOR, we introduced herein the second Glu
to the second coordination sphere of the FeB site by mutating
an Ile to a Glu (named I107E FeBMb). We show that the second
Glu results in a ∼100% increase in NOR activity through hydro-
gen bonding interactions and that the two glutamates have
dramatically different effects on the heme reduction potential.
Additionally, by comparing the EPR, electrochemistry, X-ray
structures, and NOR activity of iron, copper, and zinc derivatives
of the designed protein, we have obtained deeper insights into the
roles of metal ions in NOR.
Results
Structure and Function of Fe(II)-I107E FeBMb. The X-ray crystal struc-
tures of heme-containing I107E FeBMb without metal ion in the
FeB site and with Fe2þ in the FeB site are solved at 1.42-Å and
1.65-Å resolution, respectively (Fig. 1 A and B and Table S1). In
the absence of metal ions in the FeB site, the structure shows a
water molecule in the FeB site, which forms hydrogen bonds with
NE2 atoms of all three His residues, both OE1 and OE2 atoms of
E68, and the OE2 atom of E107 (Fig. 1A). Upon binding Fe2þ,
the Fe(II)-I107E FeBMb structure shows that Fe2þ is coordinated
by three His, the OE2 atom of E68, and one water molecule.
Notably, a water molecule bridges Fe2þ in the FeB site and
the second glutamate (E107) with a distance of 2.32 Å to the
OE2 atom of E107 (Fig. 1B).
To probe the conformational changes of introducing the
second Glu (E107), we performed a structural alignment of Fe
(II)-I107E FeBMb and the previously reported FeðIIÞ-FeBMb
(14). The comparison, shown in Fig. 2, indicates that both the
polypeptide chain and the active site overlap well with each other.
In addition, the two nonheme irons are located at similar posi-
tions with a 0.36-Å separation from each other. In contrast,
E68 underwent a significant conformational rearrangement in
the presence of E107. These observations suggest that the active
site of FeBMb can be tuned by the formation of an extended
hydrogen bonding network, resulting from the introduction of
a second glutamate residue.
The binding of Fe2þ to deoxy I107E FeBMb was further mon-
itored by EPR (Fig. 3A). Since deoxy myoglobin contains Fe(II)
heme that exhibits no EPR signals in X-band EPR (14), we added
blue copper Cu(II)-azurin (49), a redox partner of native NOR
(19), to oxidize both the reduced heme and nonheme irons in Fe
(II)-I107E FeBMb to EPR-active Fe(III). Upon addition of Cu
(II)-azurin, the oxidation of deoxy I107E FeBMb resulted in
EPR signals at g ¼ 6.12 and 5.56, typical of a high-spin heme-
Fe(III). Upon addition of Fe2þ, however, a decrease of the
heme-Fe(III) EPR signals was observed, indicating that the
Fe2þ, when bound to the FeB site and oxidized by Cu(II)-azurin,
is spin-coupled to heme-Fe(III). Such a spin coupling mimics
that in NOR (35, 50–53), suggesting that I107E FeBMb models
NOR closely, at least in this respect.
To probe the role of the second Glu (E107) in NO reduction
activity, we measured the yield of N2O production by Fe(II)-
I107E FeBMb with excess NO under one turnover conditions.
We monitored N2O formation in the headspace of the solution
using GC/MS and compared this result to that of FeðIIÞ-FeBMb,
which lacks the second Glu (Fig. 4). Remarkably, Fe(II)-I107E
FeBMb
displays higher
activity than
FeðIIÞ-FeBMb.
After
∼20 hr, ∼24% N2O was produced by Fe(II)-I107E FeBMb, in
H29
A
H29
B
H64
E68
E107
3.04
2.81
3.02
2.20
2.24
2.24
2.12
2.21
H64
E68
E107
3.03
H43
2.15
3.41
3.16
2.62
2.32
H43
2.26
H93
H93
H29
C
H29
D
2.04
2.03
H64
E68
E107
2.09
2.10
2.91
2.21
H64
E68
E107
2.26
2.29
2.10
2.18
2.10
4.47
H43
2.07
3.04
H43
2.68
H93
H93
Fig. 1.
Crystal structures of I107E FeBMb (A) (PDB ID code 3M38), Fe(II)-I107E
FeBMb (B) (PDB ID code 3M39), Cu(II)-I107E FeBMb (C) (PDB ID code 3M3A),
and Zn(II)-I107E FeBMb (D) (PDB ID code 3M3B). Water molecules, Fe(II),
Cu(II), and Zn(II) are represented by red, green, orange, and gray spheres,
respectively.
Fig. 2.
Overlay of Fe(II)-I107E FeBMb (cyan) (PDB ID code 3M39) with
FeðIIÞ-FeBMb (orange) (PDB ID code 3K9Z).
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Lin et al.
contrast to ∼10% yield for FeðIIÞ-FeBMb, strongly indicating that
the second Glu plays an important role in NO reduction, likely
facilitating proton uptake during NO reduction.
Other Metal Ions Binding to I107E FeBMb. To find out if the resting
state of the protein, i.e., oxidized or met I107E FeBMb, can bind
other metal ions, Cu2þ or Zn2þ was titrated into met I107E
FeBMb and monitored by EPR spectroscopy (Fig. 3 B and C).
In the absence of metal ions, met I107E FeBMb exhibited
high-spin heme signals at g ¼ 6.03, 5.08, and 1.98 (Fig. 3B, black
line). Upon addition of 2 eq of Cu2þ, the signals at g ¼ 6.03 and
5.08 decreased and a broad peak around g ¼ 2.95 increased,
probably due to spin coupling between heme-Fe(III) and Cu2þ
in the FeB site. In contrast, addition of Zn2þ, a metal ion with
no unpaired electrons [i.e., incapable of spin coupling to heme-
Fe(III)], produced an increase in the high-spin heme signals at
g ¼ 5.88 and 5.60 (Fig. 3C), indicating that the interaction be-
tween E68 and heme iron was weakened after metal binding.
The X-ray crystal structures of I107E FeBMb with Cu2þ or
Zn2þ in the FeB site were solved at 1.37-Å and 1.60-Å resolution,
respectively (Fig. 1 C and D and Table S1). Compared to Fe(II)-
I107E FeBMb (Fig. 1B), a similar binding site was observed
for Cu(II)-I107E FeBMb (Fig. 1C), where H29, H43, and H64
coordinate to Cu2þ with distances of 2.09, 2.10, and 2.04 Å,
respectively, slightly shorter than the corresponding distances
in the Fe2þ structure. In comparison to Fe(II)-I107E FeBMb,
the water bridging the Cu2þ and the second Glu (E107) is shifted
toward Cu2þ in the FeB site (2.03 Å) with respect to E107
(3.04 Å). Interestingly, this bridging water molecule was not
observed in Zn(II)-I107E FeBMb (Fig. 1D), but the two O atoms
of E68 coordinate to Zn2þ with similar distances (2.26 Å for OE1
and 2.29 Å for OE2). The longer distance between OE1 of E68
B
6.12
I107E FeBMb + Azurin
I107E Fe Mb + 0 5 eq Fe
2+ + Azurin
A
5.56
I107E FeBMb + 0.5 eq Fe
+ Azurin
I107E FeBMb + 1.0 eq Fe
2+ + Azurin
I107E FeBMb + 2.0 eq Fe
2+ + Azurin
1.98
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
5.08
5.38
Magnetic Field (Gauss)
5.88
I107E FeBMb
I107E FeBMb + 0.5 eq Zn
2+
I107E Fe Mb
1 0 eq Zn
2+
C
6.03
I107E FeBMb
I107E FeBMb + 0.5 eq Cu
2+
I107E Fe Mb + 1 0 eq Cu
2+
I107E FeBMb + 1.0 eq Zn
I107E FeBMb + 2.0 eq Zn
2+
I107E FeBMb + 1.0 eq Cu
I107E FeBMb + 2.0 eq Cu
2+
1.98
~2.95
1.98
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
5.60
Magnetic Field (Gauss)
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
5.08
Magnetic Field (Gauss)
Fig. 3.
EPR spectra of deoxy I107E FeBMb (0.5 mM protein in 50 mM Bis-Tris, pH 7.0) with increasing concentrations of Fe2þ in the presence of wild-type
Cu(II)-azurin (A), and oxidized I107E FeBMb with Cu2þ (B) or Zn2þ (C). Spectra were collected at 4 K, 5 mW power, and 9.05 GHz.
30
25
Fe(II)-I107E FeBMb
Fe(II)-FeBMb
15
20
10
%)
oduction (%
N2O pro
0
5
0
4
8
12
16
20
Incubation time (hr)
Fig. 4.
Time-dependent N2O production by Fe(II)-I107E FeBMb (▴) and
FeðIIÞ-FeBMb (●) with ∼50 eq. NO under single turnover conditions. The yield
was determined by a comparison of the ratio of NO∶N2O peaks from the
GC/MS chromatograms.
Lin et al.
PNAS
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∣
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and heme iron in the Zn-bound structure (2.68 Å) in comparison
to the Cu- and Fe-bound structures, is likely the result of a weaker
interaction, which is also supported by an observed increase of
the high-spin heme signals in the EPR spectra upon Zn2þ binding
(Fig. 3C). These results suggest I107E FeBMb is capable of incor-
porating different metal ions into its designed FeB site, offering
an excellent opportunity to compare the role of these metal ions
in the same protein scaffold.
Effect of Glutamates and Metal Ions on the Redox Potential of I107E
FeBMb. Since EPR and X-ray structural studies indicate metal
binding to I107E FeBMb, we used spectroelectrochemistry to
measure the effects of glutamates and metal ions on the heme
reduction potential. When there is no metal ion in the FeB site,
the I107E FeBMb displays a reduction potential of −134 3 mV
vs. the normal hydrogen electrode (NHE) (Fig. S1A), similar to
that of FeBMb (−158 4 mV) without the I107E mutation (14).
In the presence of Cu2þ, I107E FeBMb has a reduction potential
(−137 2 mV) (Fig. S1B) almost identical to that of the same
protein in the absence of metal ions in the FeB site, indicating
that copper binding to the FeB site has little effect on the reduction
potential of the heme iron. This observation is similar to that
observed for Cu2þ binding to CuBMb (CuðIIÞ-CuBMb, 80 mV
vs. CuBMb, 77 mV) (54). On the other hand, the presence of
Fe2þ and Zn2þ increased the reduction potential of I107E
FeBMb from −134 3 mV to −64 3 mV vs. NHE (Fig. S1C)
and −105 2 mV vs. NHE (Fig. S1D), respectively. The different
effects of Cu2þ, Fe2þ, and Zn2þ on the reduction potential of
I107E-FeBMb indicate that these metal ions in the FeB site may
play different roles through different coordination properties.
NOR Activity of I107E FeBMb in the Presence of Different Metal Ions.
The NO reduction activity of I107E FeBMb in the presence of
Fe2þ, Cuþ, or Zn2þ was monitored by GC/MS under single turn-
over conditions. When Fe(II)-I107E FeBMb was exposed to
excess NO, N2O could be observed to form with increased yield
over time (Fig. S2). Similarly, N2O formation was observed for
Cu(I)-I107E FeBMb, indicating that Fe or Cu binding to the
FeB site results in comparable NOR activities. It should be noted
that because of the high solubility of N2O (∼25 mM in water at
room temperature), GC/MS cannot be used to quantify the rates
of NO reduction under these conditions. In contrast, no N2O
formation was observed with redox inactive Zn2þ, which demon-
strates that redox active Fe2þ or Cuþ in the FeB site plays a
crucial role in NO reduction.
To gain deeper insight into the process of NO reduction, EPR
studies were further performed to monitor the initial process of
NO reduction. In the absence of metal ions, the EPR spectrum of
ferrous I107E FeBMb-NO shows hyperfine splitting resulting
from bound NO and the proximal histidine, indicating the forma-
tion of a six-coordinate ferrous heme-NO species (Fig. 5, top
line). After incubation of Fe(II)-I107E FeBMb with excess
NO, a distinct three-line hyperfine structure appears at 15 min
(Fig. 5A), suggesting the formation of a five-coordinate ferrous
heme-NO species as a result of cleavage of the proximal
His-Fe heme bond (55). A three-line hyperfine structure was also
observed for Cuþ and Zn2þ, except that the signal intensity is low-
er with Cu(I)-I107E FeBMb-NO (Fig. 5B) and more pronounced
in Zn(II)-I107E FeBMb-NO (Fig. 5C). The lower intensity of
the three-line hyperfine structure for Cu(I)-I107E FeBMb-NO
suggests the major species formed is a six-coordinate ferric
heme-NO complex, which is EPR silent (41). These differences
further suggest that the metal ion in the FeB site plays a key role
in formation of the intermediates, thereby tuning NOR activity.
Discussion
Using Rationally Designed Proteins to Address Important Issues in
Chemistry and Biology. Important issues such as the roles of the
conserved glutamates and nonheme FeB in NOR have been
previously addressed using biochemical and biophysical studies
or biomimetic modeling (24, 25, 27–37, 45, 56, 57). As a comple-
mentary approach, rational protein design, using small, easy-to-
produce and well-characterized proteins such as myoglobin,
offers a powerful method with which to gain insights into more
complex native enzymes such as NOR (14). Similar to synthetic
models (41, 43), the metal ion at the putative FeB site in the
protein model can be substituted freely. Better yet, Glu residues
can be placed at precise locations in the protein, including the
secondary coordination sphere, due to its rigid network. By care-
fully choosing a suitable protein template, rational protein design
could be generally applied to address other important issues in
chemistry and biology.
The Roles of Glutamates. Although two conserved glutamates
(E198 and E267) are known to be crucial for NOR activity
(24, 25), their roles are not well defined (18, 19). In a previous
study (14), we demonstrated that one Glu, E68, is important for
both iron binding and NOR activity of FeBMb. The crystal struc-
tures of both FeðIIÞ-FeBMb and Fe(II)-I107E FeBMb show that
one O atom of E68 directly coordinates to FeB (Fig. 2). In syn-
thetic models of NOR, it has also been found that the presence of
a glutamic acid mimic significantly increases the stability of iron
binding to the FeB site (40). Furthermore, a theoretical study by
Blomberg et al. (58) showed that a model with an FeB coordi-
nated by three histidines, one glutamate, and one water molecule
provides an energetically feasible reaction mechanism of NO
reduction. However, the structural model of NOR constructed
recently by Reimann et al. (22) shows that the closest conserved
Glu (E267) still has its carboxylate O atom 7 Å away from FeB,
which suggests that Glu may not bind to FeB in native NOR. One
interesting finding from our study is that the Glu (E68) under-
went a significant conformational rearrangement in the presence
of another Glu (E107) (Fig. 2). Therefore, the FeBMb provides a
viable model of NOR that is consistent with Blomberg’s model,
but cannot rule out Reimann’s model due to possible conforma-
tion changes.
While the role of the first Glu is still uncertain until a 3D struc-
ture of NOR in its active form is available, the role of the second
Glu is even less defined. We address this question by introducing
a second Glu (E107) to FeBMb. The crystal structures shown in
Fig. 1 indicate that E107 interacts with a water molecule and
forms a hydrogen bonding network in both Fe(II)-I107E
FeBMb and Cu(II)-I107E FeBMb. Interestingly, although a
similar water molecule was observed in the active site of FeðIIÞ-
FeBMb (Fig. 2), activity assay data indicate that the presence of
E107 in Fe(II)-I107E FeBMb increases NOR activity by ∼100%
C
5 min
N
t l
A
5 min
B
2+
5 min
1 min
No metal
1 min
5 min
F
2+
No metal
1
i
5 min
+
No metal
2+
Zn
2+
5 min
Fe
2+
5 min
1 min
Fe
C
+
5 min
1 min
Cu
+
Z
2
Zn
2+
15 min
F
2+
Fe
15 min
5 min
C
+
Cu
+
15 min
5 min
Zn
2+
15 min
Fe2+
5
Cu
+
15 min
3000
3100
3200
3300
3400
Magnetic Field (Gauss)
3000
3100
3200
3300
3400
Magnetic Field (Gauss)
3000
3100
3200
3300
3400
Magnetic Field (Gauss)
Fig. 5.
EPR spectra of deoxy I107E FeBMb (0.5 mM) with no metal bound in
the presence of NO after 5 min (top line), with 2 eq Fe2þ (A), Cuþ (B), or Zn2þ
(C) incubated with excess NO (∼200 eq) for 1, 5, and 15 min. Spectra were
collected in 50 mM Bis-Tris pH 7.0 at 30 K, 0.2 mW power, and 9.05 GHz.
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Lin et al.
(Fig. 4), suggesting that the second Glu may potentionally play a
role in providing one of the protons during reduction of NO to
N2O. Although free Glu outside a protein has a pKa ∼4.3, the stu-
dies of native NOR showed that the pKa of its Glu close to the
active site has a value of ∼6.6 (22, 25, 33). The hydrogen binding
network in our protein models may contribute to the fine-tuning
of the Glu pKa to be more neutral, similar to that in NOR. More-
over, it is interesting that Cu(I)-I107E FeBMb also shows NOR
activity, which provides an interesting protein model of HCOs
with NOR function (26–28), even though Glu residues are not
conserved in native HCOs.
Additionally, spectroelectrochemical studies showed that the
reduction potential of I107E FeBMb with no metal ion in the
FeB site is similar to that of FeBMb, but much lower than that
of CuBMb (77 mV) (54), which contains the same three His,
but no Glu in the metal-binding site above the heme. Since both
FeBMb and I107E FeBMb contain the V68E mutation that has
been shown to decrease the heme reduction potential of native
myoglobin from 59 mV to −137 mV (59), it is likely that the
introduction of a negatively charged Glu close to the heme group
is what is responsible for the dramatically lower heme redox
potential. A conserved glutamate, predicted to be located near
the catalytic heme b3 in NOR, was proposed to be responsible
for a ∼260-mV decrease in reduction potential (60 mV) in
comparison to the other two heme centers, heme b (345 mV)
and heme c (310 mV) (60). Our FeBMb and I107E FeBMb mod-
els mimic this feature of NOR. Notably, although introduction of
the first Glu (E68) lowered the heme potential by ∼110 mV (14),
introduction of the second Glu via the I107E mutation did not
result in a significant difference in the heme reduction potential,
suggesting that the effect of the two conserved Glu residues in
NOR on heme reduction potential is not additive, with the effects
highly dependent on the location of the Glu.
The Roles of Metal Ions. The roles of metal ions in NOR are an-
other important question as iron is found in the native FeB site
and HCO employs copper at the corresponding CuB site. With
different metal ions in the FeB site, the crystal structures clearly
show the heme and nonheme dinuclear center existing in differ-
ent local environments (Fig. 1). Although a similar hydrogen
bond network is formed in both Fe(II)-I107E FeBMb and Cu
(II)-I107E FeBMb, the conformation of E68 and E107 with re-
spect to the nonheme metal center and heme iron is different
from each other. Moreover, the coordination geometry differs
significantly with Zn2þ in the FeB site. A hydrogen bond is absent
from the Zn crystal structure, but both the O atoms of E68 act as
metal-binding ligands. These observations demonstrate that the
identity of the metal ion in the FeB site can tune the active site
through their interactions with the His and Glu ligands, resulting
in formation of different coordination geometries with different
hydrogen bonds.
In addition to structural fine-tuning, the metal ion at the FeB
site can also tune the heme iron reduction potential in I107E
FeBMb. Spectroelectrochemical studies showed that the binding
of Fe2þ or Zn2þ results in an increase in the heme reduction
potential by ∼70 mV and ∼30 mV, respectively (Fig. S1). In
the case of Cu(II)-I107E FeBMb, the crystal structure shows that
OE1 of E68 is closer to the heme iron (2.07 Å) (Fig. 1C) than its
metal-free form (2.15 Å) (Fig. 1A). The stronger interaction
from the negatively charged E68 could offset the effect of
positively charged Cu2þ binding, resulting in similar reduction
potentials
observed
for
Cu(II)-I107E
FeBMb
and
I107E
FeBMb.
In a previous study (61), EPR data showed that during NO
reduction, the binding of Cuþ to the CuB site of CuBMb can
weaken the proximal heme Fe-His bond, while complete cleavage
of the heme Fe-His bond occurred when Zn2þ was bound to
CuBMb-NO. In this study, we observed that a five-coordinate
heme-NO species was formed with Fe2þ, Cuþ, or Zn2þ bound
to the FeB site of I107E FeBMb (Fig. 5). Significantly, a five-
coordinate heme-NO species has also been observed for both
NOR (30, 31, 35) and the member of the HCO family with the
highest NO reduction activity, cytochrome cbb3 oxidases (26, 62).
However, this species was not observed for FeðIIÞ-FeBMb-NO
and CuðIÞ-FeBMb-NO, which lack the second Glu (E107). In
both these cases, the proximal heme Fe-His bond was only
weakened, as indicated by a decrease of the nine-line hyperfine
splitting signals in the EPR spectra (Fig. S3). These observations
suggest that formation of a five-coordinate heme-NO species may
play an important role in NOR reactivity.
Conclusions
We have successfully designed a structural and functional model
of NOR, by introducing a second glutamate in the vicinity of the
FeB site, named I107E FeBMb. This protein model mimics native
NOR more closely by bearing the structural feature of three his-
tidines and two glutamates in the FeB site, as predicted for native
NOR. We have demonstrated that the two glutamates can play
different roles in NO reduction activity; namely, one acts as a li-
gand to FeB (E68), and the other acts as a proton transfer group
(E107). Furthermore, by substituting different metal ions into the
nonheme metal site, we have demonstrated that FeB plays crucial
roles in fine-tuning the active site by donating electrons and by
mediating the formation of a five-coordinate heme-NO inter-
mediate during NO reduction. In the absence of a crystal struc-
ture for native NOR, this study offers an ideal protein model and
provides valuable structural as well as mechanistic information
for native NOR.
Materials and Methods
Protein Preparation. I107E FeBMb (swMb L29H/F43H/V68E/I107E) was con-
structed, expressed, and purified using the procedure described previously
(14). The purity and identity were confirmed by SDS-PAGE and electrospray
ionized MS: observed: 17; 392 1 Da; calculated: 17,391 Da.
EPR Spectroscopy. EPR spectra were recorded on a Bruker ESP 300 equipped
with an Oxford liquid helium cryostat and an ITC4 temperature controller.
The samples of met I107E FeBMb, Cu(II)-, or Zn(II)-I107E FeBMb were pre-
pared as described previously (14). The samples of NO-bound deoxy
I107E FeBMb, Cu(I)-, Fe(II)-, or Zn(II)-I107E FeBMb were prepared by inject-
ing 1 mL of purified NO gas into the EPR tube containing 300 μL of protein
(0.5 mM, 10% glycerol, in 50 mM Bis-Tris pH 7.0). The samples were then
flash frozen in liquid N2 after incubation for 1, 5, or 15 min. The molar
extinction coefficient of the Soret band of I107E FeBMb at 406 nm
(175 mM−1 · cm−1), calculated using the standard hemochromagen method
(63), was used to determine protein concentration. The metal sources of
Cu(I), Cu(II), Zn(II), and Fe(II) were ½ðCH3CNÞ4CuPF6, CuSO4, ZnSO4 · 7H2O,
and FeCl2, respectively.
Spectroelectrochemical Measurements. Protein reduction potentials were
measured using an optically transparent thin layer electrode as previously
described (64). The potential of the working electrode was applied in
the positive direction for deoxy I107E FeBMb with Fe2þ and in the negative
direction for metal free and with Cu2þ or Zn2þ. Other procedures are the
same as described previously (54).
X-Ray Crystallographic Studies. Fe(II)-I107E FeBMb was crystallized anaerobi-
cally in a glove box at room temperature using the conditions described for
FeðIIÞ-FeBMb (14). I107E FeBMb, Cu(II)-, or Zn(II)-I107E FeBMb were crystallized
aerobically. Diffraction-quality crystals were soaked in a cryoprotectant
solution of 30% PEG 400 and flash frozen in liquid nitrogen. Diffraction data
were collected at the Brookhaven National Lab Synchrotron Light Source
X12C beamline. The crystal structure was solved using the same method
as for FeðIIÞ-FeBMb (14).
NOR Activity Assay. NO reduction was monitored by GC/MS. The protein was
reduced to the deoxy form by excess dithionite that was removed with a
size-exclusion column (PD-10). Then 2 eq metal, Cu(I), Fe(II), or Zn(II), was
added to the protein solution (0.6 mM, 3 mL in 50 mM Bis-Tris buffer, pH
7.0). The samples were prepared anaerobically in a glove box. Purified NO
Lin et al.
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gas was injected into the head space of the reaction flask with the molar
ratio of NO∶protein ≈50∶1. Other procedures are the same as described
previously (14, 61).
ACKNOWLEDGMENTS. We thank Dr. Mark J. Nilges for help with EPR analysis,
and Furong Sun and Beth D. Eves for aiding in GC/MS data collection. This
work was supported by NIH Grant GM062211.
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3M3F
|
PEPA bound to the ligand binding domain of GluA3 (flop form)
|
The molecular mechanism of flop-selectivity and subsite
recognition for an AMPA receptor allosteric modulator:
Structures of GluA2 and GluA3 complexed with PEPA
Ahmed H. Ahmed§, Christopher P. Ptak§, and Robert E. Oswald*
Department of Molecular Medicine, Cornell University, Ithaca, NY 14853 USA
Abstract
Glutamate receptors are important potential drug targets for cognitive enhancement and the
treatment of schizophrenia in part because they are the most prevalent excitatory neurotransmitter
receptors in the vertebrate central nervous system. One approach to the application of therapeutic
agents to the AMPA subtype of glutamate receptors is the use of allosteric modulators, which
promote dimerization by binding to a dimer interface thereby reducing desensitization and
deactivation. AMPA receptors exist in two alternatively spliced variants (flip and flop) that differ
in desensitization and receptor activation profiles. Most of the structural information on
modulators of the AMPA receptor target the flip subtype. We report here the crystal structure of
the flop-selective allosteric modulator, PEPA, bound to the binding domains of the GluA2 and
GluA3 flop isoforms of AMPA receptors. Specific hydrogen bonding patterns can explain the
preference for the flop isoform. This includes a bidentate hydrogen bonding pattern between
PEPA and N754 of the flop isoforms of GluA2 and GluA3 (the corresponding position in the flip
isoform is S754). Comparison with other allosteric modulators provides a framework for the
development of new allosteric modulators with preferences for either the flip or flop isoforms. In
addition to interactions with N/S754, specific interactions of the sulfonamide with conserved
residues in the binding site are characteristics of a number of allosteric modulators. These, in
combination, with variable interactions with five subsites on the binding surface lead to different
stoichiometries, orientations within the binding pockets, and functional outcomes.
Membrane receptors are the cell's gatekeepers, allowing chemical signals access to the cell's
pathways. Through the binding of endogenous ligands, receptors identify relevant
environmental cues and facilitate cell-cell communication. The regulation of membrane
receptors has become an important goal of drug discovery efforts (1,2). By targeting the
physiological (orthosteric) ligand-binding site, agonists and antagonists control the function
of membrane receptors. Unfortunately, exogenously induced agonist-activation at the
orthosteric site can cause toxic effects from overstimulation. Allosteric modulator binding
sites use a distinct avenue for altering the natural response of a receptor. The ability of some
allosteric modulators to enhance receptor stimulation, while not actually providing the
trigger for stimulation, is a clear advantage that conserves the endogenous signaling
pathway. Being important mediators of higher-order processes such as learning and
memory, ionotropic glutamate receptors (iGluRs) have attracted a great deal of interest as
allosteric modulator targets (3–6). Of clear therapeutic importance, various
neurodegenerative disorders such as Parkinson's and Alzheimer's diseases, Huntington's
*Corresponding author; telephone: 1-607-253-3877; fax: 1-607-253-3659; email: reo1@cornell.edu.
§These authors contributed equally to this work.
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 April 7.
Published in final edited form as:
Biochemistry. 2010 April 6; 49(13): 2843–2850. doi:10.1021/bi1000678.
NIH-PA Author Manuscript
NIH-PA Author Manuscript
NIH-PA Author Manuscript
chorea, and neurologic disorders including epilepsy and ischemic brain damage have been
linked to iGluRs (7).
The crystal structure of GluA2 (8) clarifies years of speculation on the complex arrangement
of the glutamate receptor's four subunits (9). The GluA2 can be dissected into 3 functionally
distinct layers. Farthest from the membrane, the amino terminal domain (ATD) can act as a
peripheral regulatory domain but is also involved in assembly and trafficking (10,11).
Sandwiched between the ATD and the membrane domain, the ligand-binding domain (LBD)
recognizes the neurotransmitter signal and directly regulates receptor activation (12).
Structures for both isolated extracellular domains (ATD and LBD) reveal a dimeric
organization (13–15). At the membrane interface, two alternative linker conformations
transition the 2-fold symmetry, which is adopted by both extracellular domains, into the 4-
fold symmetry of a membrane-traversing cation-selective channel (8,16). For iGluRs, the
ion channel domain confers functional relevance with its ability to selectively conduct the
flow of ions across the cell's membrane. The layers of extracellular domains, each with the
potential for multiple control points, allosterically regulate the ion channel domain's function
(8). Therefore it is not surprising that the ATD, the LBD, and the LBD-channel linker have
all been shown to be effective targets of allosteric modulators (13,17,18).
Since the structures of the ATD and the full iGluR channel have only recently been solved,
allosteric drug-binding sites external to the LBD have not been fully explored in molecular
detail. However, the decade-old LBD structure has proved to be indispensable as a heavily
exploited scaffold for understanding agonist, partial agonist, and antagonist binding
interactions as well as their ability to regulate channel gating behavior (12,19,20). Although
the dimeric organization is consistent across all iGluR subtypes, the molecular details of
LBD-agonist specificity define the subtype families into N-methyl-D-aspartic acid (NMDA)
receptors (21), α-amino-3-hydroxy-5-methyl-4-isoxazole-propionic acid (AMPA) receptors
(12), and kainate receptors (22). Because all subtypes are constrained by their conserved
sensitivity to glutamate stimulation, diversity at the orthosteric site is evolutionarily limited
and most agonists display cross-subtype activity. An allosteric modulator-binding site within
the quaternary LBD structure is located along the dimer interface (18) and offers improved
discrimination by modulators. Drugs that bind to the allosteric sites on the LBD dimer
interface can enhance the activity of iGluRs (23) and increase performance on tests of
memory (24). Except for the LBD structures with modulatory ions bound to the dimer
interface (25–27), only LBD structures from the AMPA receptor subtype, GluA2, have been
reported with bound allosteric modulators (18,28–31). Within the structures, the bound
modulatory drugs stabilize the LBD dimer interface, which is required for activation of the
ion channel and is dissociated during desensitization (18).
Although the residues that line the allosteric modulator-binding pocket do not differ between
AMPA receptors subtypes (GluA1–4), the ability of allosteric modulators to stabilize the
activated state still varies (32,33). Also, AMPA receptors can be alternatively spliced into
what is referred to as flip and flop isoforms (34). Modulator selectivity (23), desensitization
(35), and channel closing rates (36) differ between flip and flop. Although several of the
amino acid differences between the two forms are located in or near the allosteric
modulator-binding site, the difference at position 754 (serine in flip, asparagine in flop)
seems to be entirely responsible for the functional differences between allosteric modulator
regulation of the flip and flop variants (23,28,32). Cyclothiazide (CTZ) and some other
thiazide derivatives have improved binding to the flip form due to a hydrogen bond between
S754 and the NH of the fused thiazide ring (28). In the case of the flop form, the
alternatively spliced sequence places an asparagine in the 754 position, which is not
optimally positioned to form a hydrogen bond. Sekiguchi et al. (33) introduced an allosteric
modulator of AMPA receptors (4-[2-(phenylsulphonylamino)ethylthio]-2,6,-
Ahmed et al.
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difluorophenoxyacetamide, PEPA) with a preference for the flop form. In fact, the relative
sensitivity of CTZ to PEPA has been used as a diagnostic for the prevalence of flip vs. flop
versions of AMPA receptor in particular cell types (37). PEPA shows potential in treatment
of post ischemic memory impairment (38) and contextual fear (39) but despite PEPA's
unique flop sensitivity, the modulator has not yet been used as a lead compound in SAR
studies.
For drug discovery to be guided by structures, understanding the possible molecular
interactions between modulators and the dimer interface is essential. We have shown
previously (31) that changes in the structures of CTZ derivatives can reorient the modulator
within the binding site. Subsequently, we proposed that the allosteric modulator site is
comprised of 5 subsites (Figure 1C). In the present study, we determine the three
dimensional structures of PEPA bound to the GluA2o and GluA3o LBDs (flop forms), and
use PEPA's binding interactions to further characterize the subsite specific binding
properties displayed by allosteric modulators. The amide group of PEPA makes a direct
hydrogen bond to N754, explaining the preferential action of PEPA on the flop form of
AMPA receptors. Another key structural element, the sulfonamide group of PEPA, is
conserved with the biarylsulfonamide class of allosteric modulators (6) and interacts with
the same residues of the dimer interface (8,30). Although previously classified as unrelated,
PEPA and the large group of biarylsulfonamide have similarities, which suggest that specific
PEPA groups (particularly the unique flop-interacting amide) can be strategically integrated
into biarylsulfonamide SAR studies.
Experimental Procedures
Materials
PEPA was purchased from Tocris (Ellisville, MO). The GluA2 S1S2J construct was
obtained from Eric Gouaux (Vollum Institute; 12).
Protein Preparation and Purification
GluA2 S1S2 consists of residues N392 - K506 and P632 - S775 of the full rat GluA2o
subunit (40), a `GA' segment at the N-terminus, and a `GT' linker connecting K506 and
P632 (12). A similar construct of GluA3 S1S2 was prepared as described previously (41).
pET-22b(+) plasmids were transformed in E. coli strain Origami B (DE3) cells and were
grown at 37°C to OD600 of 0.9 to 1.0 in LB medium supplemented with the antibiotics
(ampicillin and kanamycin). The cultures were cooled to 20°C for 20 min. and isopropyl-β-
D-thiogalactoside (IPTG) was added to a final concentration of 0.5 mM. Cultures were
allowed to grow at 20°C for 20 h. The cells were then pelleted and the S1S2 protein purified
using a Ni-NTA column, followed by a sizing column (Superose 12, XK 26/100), and
finally an HT-SP-ion exchange-Sepharose column (Amersham Pharmacia). Glutamate (1
mM) was maintained in all buffers throughout purification. After the last column, the protein
was concentrated and stored in 20 mM sodium acetate, 1 mM sodium azide, and 10 mM
glutamate at pH 5.5.
Crystallography
For crystallization trials, the protein was concentrated to 0.2 – 0.5 mM in 10 mM glutamate
using a Centricon 10 centrifugal filter (Millipore, Bedford, MA). For the PEPA-bound
structures, PEPA was added to 5 mM. The final protein concentration was 0.2 to 0.3 mM.
Crystals were grown at 4°C using the hanging drop technique, and the drops contained a 1:1
(v/v) ratio of protein solution to reservoir solution. The reservoir solution contained 14–15%
PEG 8K, 0.1 M sodium cacodylate, 0.1–0.15 M zinc acetate, and 0.25 M ammonium sulfate,
pH 6.5.
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Data were collected at the Cornell High Energy Synchrotron Source beam line A1 using a
Quantum-210 Area Detector Systems charge-coupled device detector. Data sets were
indexed and scaled with HKL-2000 (42). Structures were solved with molecular
replacement using Phenix (43). Refinement was performed with Phenix (43), and Coot 0.5
(44) was used for model building.
Results
Structure of PEPA bound to GluA2 S1S2 flop
The structure of glutamate bound to GluA2o S1S2 (3dp6; 41) was used as the initial search
probe for the molecular replacement solution of PEPA bound to GluA2o S1S2 with
glutamate in the agonist-binding site. PEPA was then modeled into two symmetrical
positions within the density found at the dimer interface, and the structure was optimized
using Phenix (43). The refinement statistics are given in Table 1. The resolution is 1.85 Å,
and three unique copies are found in the unit cell. The overall structure of the S1S2 domain
is very similar to the structure in the absence of PEPA, with contacts between glutamate and
the protein unchanged. However, PEPA clearly binds within the dimer interface, making
contacts with both monomers within the dimer. As shown in Figure 1, one PEPA molecule
binds per dimer interface. However because the dimer interface is symmetrical, two
equivalent orientations (related by a 180° rotation) are possible. Electron density for both is
seen in the crystal structure, although the intensity of one orientation is greater than the
other.
The binding of PEPA to the dimer interface increases the distance between the two
monomers that form the dimer by approximately 1.5 Å. This allows the relatively large
PEPA molecule to fit within the interface, but also increases the separation between the
linkers to the ion channel (the distance increases from 39.4 Å to 41 Å; Figure 1A). Relative
to the core of Lobe 1, both the J/K helices and one β strand (P105-G110) connecting the two
lobes are displaced slightly away from the dimer interface (Figure 1B). In addition, Lobe 2
is slightly twisted relative to glutamate-bound S1S2 in the absence of PEPA (3dp6; 41).
PEPA binds at the bottom of a water-filled, inverted U-shaped cleft with five subsites (A, B/
B′, and C/C′; 31). Upon binding, crystallographic waters are displaced from the central A
subsite and more buried C/C′ sites, with the waters in the B/B′ subsite remaining (Figure
1C). This displacement of presumably ordered water would be likely to contribute a
favorable entropy component to binding.
The sidechains of P494 are at the center of the interface and the edge of the two proline
rings from each monomer form the base of the binding site in which the difluorophenyl ring
resides (Figure 2A). This is close to the position of the methoxybenzoyl ring of aniracetam
in its structure bound to GluA2-S1S2(FW) (29). The other side of the ring is exposed to
S497 and S729. The sidechain hydroxyl of S497 is oriented toward the dimer interface in the
absence of PEPA, but rotates out toward the solvent to accommodate the difluorophenyl ring
of PEPA (Figure 1B). The amide of PEPA is involved in a network of hydrogen bonds with
sidechain hydroxyl of Y424, the backbone carbonyl of F495, the sidechain carboxyl of
D760, the sidechain amide of N754, and two water molecules (Figure 2A). The most
striking of these hydrogen bond pairs is with N754. This represents the only difference
between the flip (S754) and flop (N754) isoforms in the PEPA binding site and is almost
certainly a major source of the preference for the flop isoform. The phenyl-sulfonylamide
side of PEPA inserts into a hydrophobic pocket formed by sidechain methyls of I481 and
L751 as well as methylene groups contributed by K493, N754, and E755 (Figure 2B). It is
possible that the contribution by methylene group of N754 provides a more hydrophobic
pocket than S754 in the flip form, further contributing to the preference for the flop form.
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Because the dimer interface is symmetrical, PEPA can bind in two orientations and both are
observed in the crystal. For this reason, changes in the protein due to a specific interaction
with PEPA can be partially masked because each monomer is a weighted average of two
orientations of bound PEPA. However, one orientation has a stronger density than the other,
providing some insight into the extent of changes in the dimer interface that are produced by
PEPA binding. As shown in Figure 2C, the two monomers comprising the dimer differ more
within the PEPA binding site than the corresponding monomers in the absence of PEPA.
One turn of helix J (L751 to N754) contains important determinants for both orientations of
PEPA. In one orientation the amide group of PEPA interacts with the sidechain of N754,
and in the other, the aromatic ring of PEPA inserts between a hydrophobic pocket formed by
the sidechain of L751 and the methylene group of N754. In the orientation for which the
density of the amide of PEPA is stronger, N754 is better positioned to form an H-bond
(Figure 2D); whereas, in the other side of the interface, N754 is oriented to form an H-bond
with the carbonyl of S729. This change in orientation facilitates the insertion of the aromatic
ring of PEPA into the hydrophobic pocket, which is accompanied by a small shift in the
sidechain of L751 to accommodate the aromatic ring (Figure 2D). Since these structures are
weighted averages, it is possible that the actual positions of these sidechains involve an even
greater movement than is seen from the asymmetry of the crystal.
Structure of PEPA bound to GluA3 S1S2 flop
In studies of the physiological effects of PEPA, a significant difference between subtypes
has been observed, with GluA3 being most susceptible to modulation (33). The structure of
GluA3i S1S2 bound (flip form) to glutamate has been reported previously (41). Since PEPA
preferentially binds to the flop form, the GluA3o structure was determined bound to
glutamate with and without PEPA (Figure 3A). Like GluA2o, in the absence of PEPA,
GluA3o has three copies in the asymmetric unit. Comparing lobe closure between GluA3i
and GluA3o, the flop form is slightly more closed (1.6° ± 0.7°).
In the presence of PEPA, GluA3o was present in one copy in the asymmetric unit, and PEPA
was observed with the same density in two symmetrical orientations. Like GluA2o bound to
PEPA, the dimer interface (assessed using the symmetrical molecule in the crystal) was
displaced relative to the unbound from (Figure 3A) by approximately 2.5 Å at the position
of the linker replacing the ion channel domain. Within the binding site, three sidechains
exhibited different rotamers compared with the GluA2o structure bound to PEPA (Figure
3B). For PEPA-bound GluA3o, both S497 and S729 assumed rotameric states that differed
both from GluA2o bound to PEPA and from GluA2o and GluA3o in the absence of PEPA. In
the case of S729, the rotameric state in combination with a slight movement of the amide of
PEPA (relative to the GluA2o structure) would make an H-bond with the sidechain of S729
(shown in Figure 2A for GluA2o) unlikely. In the case of N754, the sidechain is displaced
relative to the GluA2o-PEPA structure so that only one H-bond is made to the amide of
PEPA. This may be a result of averaging of the two orientations of PEPA only one of which
forms a bidentate H-bond with N754.
Discussion
The goal of allosteric modulation, like orthosteric modulation, is often to stabilize a
conformational state of a dynamic protein (45). The activated state of iGluRs is naturally
unstable allowing the channel to desensitize (46). Disruption of the symmetrical dimer
interface between LBDs is thought to initiate desensitization-mediated channel closure (47).
By maintaining the LBD dimer, positive allosteric modulators can prevent desensitization
and prolong activation (18). Currently, 15 crystal structures of the GluA2 LBD with bound
allosteric modulators are deposited in the Protein Data Bank (48). All of these modulators
bind to a large crevice with 2-fold symmetry along the symmetric dimer interface (18). The
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large variation in structure among allosteric modulators results in significant variations in
binding orientations and interactions. At least four distinct binding modes have been
identified: (1) A-subsite class (aniracetam, CX614 (29)), (2) classical thiazide (cyclothiazide
(18), TCMZ, ALTZ (31)), (3) the shifted thiazide class (IDRA-21, HCMZ, HFMZ; (31)),
and (4) the full spanning class (PEPA (this paper), dimeric biarylpropylsulfonamide (30),
LY404187 (8)). Overlaying modulators from these structural classes has led to the proposal
that the allosteric modulator site is comprised of a series of subsites (Figure 1C;31).
Positioned at the center of the binding-site, the symmetric A subsite is narrow and allows
entrance to only one molecule. Two subsites (B and C) lie at each end of the A subsite with
the hydrophobic C subsite located more deeply in the pocket effectively defining five
subsites (A, B, B′, C, and C′).
In the open state, the subsites are filled with water, which may act to weakly stabilize the
dimer. Allosteric modulators generate stronger interactions across the subsites thereby
increasing the linkages between the monomers. The simplest modulator class, including
aniracetam and CX614, fills the A subsite with one molecule but does not enter the
peripheral B and C subsites (29). The two classes of thiazide-based modulators account for
10 of the 15 solved allosteric modulator-GluA2 crystal structure complexes (18,28,31). The
classical thiazide (CTZ-like) binding class and the shifted thiazide (IDRA-21-like) binding
class are positioned respectively in the B and C subsite or mainly the C subsite. Most of the
thiazide modulators do not extend across the A subsite and therefore can bind two molecules
per dimer. However, a few of the newly described shifted thiazides (HFMZ, HCTZ; 31)
enter the A subsite but only enough to impair binding of a second modulator. The dimeric
biarylpropylsulfonamide compound ((R,R)-N,N-(2,2'-[Biphenyl-4-4'-Diyl]Bis[Propane-2,1-
Diyl]) Dimethanesulfonamide) described by Kaae et al. (30) was the first allosteric
modulator shown by crystallography to extend along the entire length of the inner dimer
cavity from C to C′ subsites. PEPA also interacts with J helices from both monomers, which
cap the ends of the modulator-binding pocket. The density occupied by both symmetrical
copies of PEPA overlays the dimeric biarylsulfonamide compound as both modulators
represent the full spanning class (Figure 4B).
The GluA flip and flop splice variants differ by only a few residues along the J helix in the
LBD; however, residue 754 (Asn in flop and Ser in flip) is positioned between the B and C
subsites. For thiazides, a clear preference in binding to the flip-form is mediated by a
hydrogen bond between the hydrobenzothiadiazide ring and S754 (28). In contrast, PEPA is
flop-selective and the PEPA-bound structure provides the first structure containing a direct
interaction between a modulator and the flop form's N754. The amide of PEPA extends
straight out from the A subsite and across the B and C subsite interface to make an amide-
amide hydrogen bond with N754 (Figure 2A). Unlike most other AMPA modulators, PEPA
fills neither the B nor the C subsites but interacts directly with the J helix. A similar
interaction is seen with LY404187 (49) bound to GluA2i (8). Strong hydrogen bonding can
occur between two amides (50) and has been shown to be responsible for driving
oligomerization of transmembrane leucine zippers (51). The distances between the
interacting amides in the PEPA-bound structure support a bidentate hydrogen-bonding
pattern, which is much stronger and more specific than a typical hydrogen bond. While
PEPA is selective for the AMPA receptor's flop form, a weaker but still existent potentiation
of the flip form has been observed (33,52). Replacing N754 (flop) with S754 (flip) would
not prevent PEPA from binding; however, serine would provide only one hydrogen-bonding
partner for PEPA's amide with an extended interaction distance. In contrast, LY404187
displays a preference for the flip isoform (53), and its cyano group extends out to interact
directly with S754. The cyano-S754 interaction is a clear flip analog of the flop-selective
PEPA amide-N754 interaction (Figure 4A).
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Opposite to the amide on the PEPA molecule, a sulfonamide is tethered to the
difluorophenyl ring (Figure 4A). Within the dimer interface, the sulfonamide is positioned
so the nitrogen can hydrogen bond directly with the carbonyl of P494 (Figure 4C). A
sulfonamide oxygen points toward the amide nitrogen of G731. The angle of the peptide
plane is perpendicular to the sulfonamide oxygen, making a hydrogen bonding interaction
unlikely (Figure 4D). Instead, a dipole-dipole or charge-dipole interaction may occur. The
amide nitrogen of a polypeptide supports at least a partial positive charge (54), which would
interact with the strongly electronegative sulfonamide oxygen (55). Interestingly, both the
dimeric biarylsulfonamide (30) and LY404187 (8), other members of the full spanning
modulator class, also have a sulfonamide that interacts with the same backbone atoms of
P494 and G731 as PEPA (Figure 4C).
A large number of biarylsulfonamides have been identified that modulate AMPA receptors
and are being evaluated for therapeutic use in the treatment of depression and Parkinson's
disease (56). The conserved sulfonamide reveals a previously unidentified relationship
between PEPA and the biarylsulfonamide modulators. When the perpendicular peptide bond
plane including G731 is fixed, the sulfonamide on three overlayed modulators varies by 1.2
Å along the length of the interface with the PEPA sulfonamide being positioned closer to the
A subsite (Figure 4D). A shift of the sulfonamide also results in a shift in the corresponding
P494 across the interface presumably to maintain the hydrogen bond with the modulator's
amine. The sulfonamide forms an important bridge between the two dimer halves. For
PEPA, a phenyl-sulfonamide replaces the methyl-sulfonamide in the dimeric
biarylsulfonamide and fits snuggly against L751. Based on the orientation-induced
asymmetry within the GluA2-complex structure, the phenyl pushes the J helix away from
PEPA thereby affecting the C subsite (Figure 2C and D). Residues lining the C subsite are
on the same beta strand as G731, which must shift if the C subsite is to remain together and
presumably explain the 1.2 Å shift relative to the dimeric biarylsulfonamide. In fact, the
same phenyl-sulfonamide group substitution in a biarylpropylsulfonamide decreases the
modulatory effect of the derivative in SAR studies (57). For biarylpropylsulfonamides, the
optimal sulfonamide substitution was found to be either an ethyl or an iso-propyl group,
which should both fit without significantly disrupting the J helix or C subsite (57).
The PEPA-bound crystal structure from AMPA receptor subtypes, GluA2 and GluA3, do
not display major differences in binding interactions even though PEPA exhibits a stronger
effect on GluA3 (33). For GluA2, an asymmetry in the receptor-binding pocket was
observed while no significant difference in PEPA density was seen for the each orientation
within the GluA3 crystal structure. In addition, a number of side chains exhibit different
rotameric states between the two structures, although it is unlikely that these small changes
significantly impact the differential effects on the two subtypes. Although no structural
differences have been identified between GluA2 and GluA3 that would obviously impact
PEPA affinity, the possibility exists that subtle differences arising from the sequence
differences peripheral to the binding site may be important as has been described in the case
of the agonist binding site of GluA4 (58).
We have explored how PEPA (this paper) and other allosteric modulators (31) interact with
the GluA interface in the context of drug design. Together the identification of a conserved
group between PEPA (this paper) and biarylpropylsulfonamides (8,30) and the regional
nature of various subsite-functional group interactions provide a backdrop to extend
biarylpropylsulfonamide SAR studies (57) to include PEPA and biarylpropylsulfonamide
chimeras. Although optimizing the stability of the dimer interface provides a starting point
for SAR studies, additional constraints should be considered including the ability of the
modulator to enter the cavity, the dynamic structure of the dimer interface during closed,
open, and desensitized state transitions, and the ability of the modulator to cross the blood-
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brain barrier before being metabolized. This definition of the allosteric modulator binding-
site should provide guidance in glutamate receptor allosteric modulator pharmacology.
Acknowledgments
We thank Prof. Eric Gouaux (Vollum Institute) for the GluA2 S1S2J construct, and Prof. Linda Nowak (Cornell)
for the full-length GluA3 construct.
This work was supported by a grants from the National Institutes of Health (R01-GM068935, R01 NS049223, and
R21 NS067562). This work is based upon research conducted at the Cornell High Energy Synchrotron Source
(CHESS), which is supported by the National Science Foundation under award DMR 0225180, using the
Macromolecular Diffraction at the CHESS (MacCHESS) facility, which is supported by award RR-01646 from the
National Institutes of Health, through its National Center for Research Resources.
Abbreviations
ALTZ
althiazide
AMPA
α-amino-3-hydroxy-5-methyl-4-isoxazole-propionic acid
CLTZ
chlorothiazide
CX614
pyrrolidino-1,3-oxazino benzo-1,4-dioxan-10-one
CTZ
cyclothiazide
FW
(S)-5-fluorowillardiine
flip and flop
alternatively spliced versions of AMPA receptors that vary in rates of
desensitization and sensitivity to allosteric modulators
iGluR
ionotropic glutamate receptor
GluA1-4
four subtypes of AMPA receptor
HCTZ
hydrochlorothiazide
HFMZ
hydroflumethiazide
IDRA-21
7-chloro-3-methyl-3,4-dihydro-2H-benzo[e][1,2,4]thiadiazine 1,1-dioxide
IPTG
isopropyl-β-D-thiogalactoside
LY404187
N-[2-(4′-cyanobiphenyl-4-yl)propyl]propane-2-sulfamide
PEPA
4-[2-(phenylsulphonylamino)ethylthio]-2,6,-difluorophenoxy acetamide
NMDA
N-methyl-D-aspartic acid
S1S2
extracellular ligand-binding domain of GluA2 and GluA3
SAR
structure-activity relationships
TCMZ
trichlormethiazide
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55. Cazenave-Gassiot A, Boughtflower R, Caldwell J, Coxhead R, Hitzel L, Lane S, Oakley P,
Holyoak C, Pullen F, Langley GJ. Prediction of retention for sulfonamides in supercritical fluid
chromatography. J Chromatogr A. 2008; 1189:254–265. [PubMed: 17977551]
56. O'Neill MJ, Witkin JM. AMPA receptor potentiators: application for depression and Parkinson's
disease. Current drug targets. 2007; 8:603–620. [PubMed: 17504104]
57. Ornstein PL, Zimmerman DM, Arnold MB, Bleisch TJ, Cantrell B, Simon R, Zarrinmayeh H,
Baker SR, Gates M, Tizzano JP, Bleakman D, Mandelzys A, Jarvie KR, Ho K, Deverill M,
Kamboj RK. Biarylpropylsulfonamides as novel, potent potentiators of 2-amino-3- (5-methyl-3-
hydroxyisoxazol-4-yl)- propanoic acid (AMPA) receptors. J Med Chem. 2000; 43:4354–4358.
[PubMed: 11087558]
58. Gill A, Birdsey-Benson A, Jones BL, Henderson LP, Madden DR. Correlating AMPA receptor
activation and cleft closure across subunits: crystal structures of the GluR4 ligand-binding domain
in complex with full and partial agonists. Biochemistry. 2008; 47:13831–13841. [PubMed:
19102704]
Ahmed et al.
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Figure 1.
(A) Comparison of glutamate-bound GluA2o S1S2 in the presence (blue) and the absence of
PEPA (green) in two orientations. Both orientations of PEPA are shown. Note that the
binding of PEPA results in a separation of the two components of the dimer (distance
between the Cα atoms of the threonine in the linker) by approximately 1.5 Å. (B) One
monomer of GluA2o S1S2 in the presence (blue) and the absence of PEPA (green) with one
orientation of PEPA shown. Both the J/K helices and the strand near S497 are displaced
upon binding PEPA. Also, the sidechains of S497 and S729 change rotameric states. (C)
Comparison of the water molecules at the dimer interface in the presence (tan spheres) and
the absence of PEPA (red spheres). PEPA is shown in both orientations. Despite the greater
separation of the dimer interface, a number of the ordered water molecules found in the
absence of PEPA are displaced by PEPA. The black circles delineate subsites of the
allosteric modulator binding site as described previously (31).
Ahmed et al.
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Figure 2.
The PEPA binding site, emphasizing the important interactions, shown in two orientations.
(A) A view of the amide side of PEPA bound to GluA2 S1S2. The hydrogen bonding
network with the amide of PEPA is shown as dotted lines. The H-bond with the sidechain of
S729 is difficult to display in the orientation used in the figure. (B) A view of the phenyl
group of PEPA inserted into a hydrophobic pocket in GluA2 S1S2. (C) RMS plot showing
more variability in the J/K helices for the PEPA-bound structure than the unbound structure.
(D) J/K helix showing where differences in the two orientations were analyzed. The amide
of PEPA-N754 interaction (blue) maintains the position of the J helix in the absence of
PEPA (green) The J helix is displaced on the phenyl side of PEPA (red).
Ahmed et al.
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Figure 3.
(A) Comparison of glutamate-bound GluA3o S1S2 in the presence (blue) and the absence of
PEPA (green) in two orientations. Both orientations of PEPA are shown. Note that the
binding of PEPA results in a separation of the two components of the dimer (distance
between the Cα atoms of the threonine in the linker) by approximately 2.5 Å. (B) One
monomer of GluA3o S1S2 in the presence (blue) and the absence of PEPA (green) with one
orientation of PEPA shown. Shown for comparison is the PEPA-bound form of GluA2o
(red). Both the J/K helices and the strand near S497 are displaced upon binding PEPA for
both GluA2o and GluA3o. Also, the sidechains of S497 and S729 are in different rotameric
states for GluA3o bound to PEPA compared with GluA3o in the absence of PEPA and
GluA2o bound to PEPA. Also, N754 is displaced in PEPA-bound GluA3o, such that only
one H-bond is possible with the amide of PEPA.
Ahmed et al.
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Figure 4.
(A) Members of the full spanning class of allosteric modulators. The shape-highlighted
regions of the modulators illuminate key contact points to the specific binding pocket
residues and subsites (as labeled for PEPA). (B) Overlay of the full spanning modulator
structures. The structures were aligned at both sets of P494 and G731 residues. PEPA (gray)
occupies a similar arrangement of subsites as the dimeric biarylsulfonamide (PDB entry
3bbr, cyan, 30) and LY404187 (PDB entry 3kgc, magenta, 8). (C) The sulfonamide bridges
the two monomers in both PEPA and the dimeric biarylsulfonamide with the same
interactions to P494 and G731. (D) The hydrogen bond between the carbonyl of P494 and
the sulfonamide is maintained when the modulator is in a shifted position relative to the
peptide plane of K730 and G731 (green disk) located on the opposite monomer.
Ahmed et al.
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Table 1
Structural Statistics
GluA2o (PEPA)
GluA3o (PEPA)
GluA3o
Space Group
P22121
P222
P222
Unit Cell (Å)
a=47.13 b=113.92 c=164.81
a=46.95 b=52.26 c=115.98
a=46.03 b=110.33 c=161.192
X-ray source
CHESS (A1)
CHESS (A1)
CHESS (A1)
Wavelength (Å)
0.977
0.977
0.977
Resolution (Å)
50–2.0 (2.03–2.00)
50–2.5 (2.54–2.00)
50–1.85 (1.88–1.85)
Measured reflections (#)
817961
62549
344340
Unique reflections (#)
70175
9141
69379
Data redundancy
6.9 (7.1)
6.0 (4.2)
4.7 (3.0)
Completeness (%)
99.9 (100.0)
99.5 (89.8)
96.5 (73.5)
Rsym (%)
11.4 (34.2)
13.0 (45.5)
7.5 (24.6)
I/σi
33.3 (7.1)
19.2 (2.5)
34.3 (3.3)
PDB ID
*
*
*
Current Model Refinement Statistics
Phasing
MR
MR
MR
Molecules/AU
3 (no NCS applied)
1
3 (no NCS applied)
Rwork/Rfree (%)
18.7/24.2
18.9/28.5
20.1/23.4
Free R test set size (#/%)
2000 (2.85)
914 (10.0)
2000 (2.88)
Number of protein atoms
5979
2030
6091
Number of heteroatoms
111
62
30
Rmsd bond length (Å)
0.011
0.015
0.009
Rmsd bond angles (°)
1.3
1.8
1.3
*to be submitted to RCSB Protein Data Bank
Biochemistry. Author manuscript; available in PMC 2011 April 7.
|
3M3J
|
A new crystal form of Lys48-linked diubiquitin
|
structural communications
994
doi:10.1107/S1744309110027600
Acta Cryst. (2010). F66, 994–998
Acta Crystallographica Section F
Structural Biology
and Crystallization
Communications
ISSN 1744-3091
A new crystal form of Lys48-linked diubiquitin
Jean-Franc¸ois Trempe,a*
Nicholas R. Brown,b Martin E. M.
Nobleb and Jane A. Endicottb
aDepartment of Biochemistry, McGill
University, 3649 Promenade Sir William Osler,
Montreal, Que´bec H3G 0B1, Canada, and
bLaboratory of Molecular Biophysics,
Department of Biochemistry, South Parks Road,
Oxford OX1 3QU, England
Correspondence e-mail:
jean.trempe@mail.mcgill.ca
Received 10 March 2010
Accepted 12 July 2010
PDB Reference: Lys48-linked diubiquitin, 3m3j.
Lys48-linked polyubiquitin chains are recognized by the proteasome as a tag for
the degradation of the attached substrates. Here, a new crystal form of Lys48-
linked diubiquitin (Ub2) was obtained and the crystal structure was refined to
1.6 A˚ resolution. The structure reveals an ordered isopeptide bond in a trans
configuration. All three molecules in the asymmetric unit were in the same
closed conformation, in which the hydrophobic patches of both the distal and
the proximal moieties interact with each other. Despite the different crystal-
lization conditions and different crystal packing, the new crystal structure of Ub2
is similar to the previously published structure of diubiquitin, but differences are
observed in the conformation of the flexible isopeptide linkage.
1. Introduction
The ubiquitin–proteasome pathway is a fundamental cellular process
in eukaryotes that controls protein degradation. Substrates are
tagged with ubiquitin through a cascade of enzymatic reactions that is
initiated by the activation of ubiquitin by the E1 enzyme, followed
by ubiquitin conjugation to E2 and finally transfer of the activated
ubiquitin from E2 to a specific substrate via an E3 ligase (Hershko &
Ciechanover, 1998). Ubiquitin molecules are assembled through the
formation of an isopeptide bond between the carboxyl-terminal
group of ubiquitin and the side-chain "-amino group of a lysine in
another ubiquitin molecule (termed the distal and proximal moieties,
respectively) or on the substrate. The 26S proteasome is able to
recognize and degrade substrates tagged with a Lys48-linked poly-
ubiquitin chain (Finley, 2009).
Several proteasomal ubiquitin receptors have been described,
including the 19S regulatory particle base subunits S5a/Rpn10
(Deveraux et al., 1994) and Rpn13 (Husnjak et al., 2008), as well as
the UBL-UBA-containing proteins HHR23/Rad23, Dsk2/Dph1 and
Ddi1/Mud1 (Bertolaet et al., 2001; Wilkinson et al., 2001). The inter-
actions of ubiquitin receptors with Lys48-linked polyubiquitin have
been characterized at the structural level (Schreiner et al., 2008;
Trempe et al., 2005; Varadan et al., 2005; Zhang, Chen et al., 2009;
Zhang, Wang et al., 2009), but as yet a crystal structure of a Lys48-
linked polyubiquitin chain bound to its receptor has not been
reported. In an attempt to obtain the structure of Lys48-linked di-
ubiquitin (Ub2) bound to the Mud1 UBA domain (Trempe et al.,
2005), cocrystallization trials were performed. Diffracting crystals
were obtained, but subsequent structure determination revealed that
the crystals were solely composed of Ub2. The Ub2 subunits in the
new crystal structure adopt the closed conformation, as observed in
the previous crystal structure (Cook et al., 1992) and in solution
(Varadan et al., 2002). The packing in the new crystal form differs
from that in the previous crystal structure and the structure reveals
differences in the conformation of the isopeptide linkage and the
loop connecting 1 and 2.
2. Materials and methods
2.1. Purification and crystallization
Ub2 was synthesized in vitro as described previously (Piotrowski et
al., 1997; Trempe et al., 2005). Briefly, the reaction mixture contained
50 mM Tris–HCl pH 8.0, 2 mM ATP, 5 mM MgSO4, 0.5 mM bovine
ubiquitin, 0.5 mM recombinant human His6-E1 and 50 mM recombi-
nant budding yeast His10-Cdc34. The synthesis reaction was per-
formed at 310 K overnight. Bovine ubiquitin was purchased as a
lyophilized powder (Sigma–Aldrich), His6-E1 ubiquitin-conjugating
enzyme was expressed from a recombinant baculovirus in Sf9 insect
cells and recombinant His10-Cdc34 was expressed in BL21 (DE3)
Escherichia coli cells from a pET16 expression plasmid. Both His-
tagged proteins were purified using Ni–NTA agarose resin (Qiagen).
The amino-acid sequence of bovine ubiquitin is identical to that
of human ubiquitin and yeast Cdc34 has previously been shown to
synthesize Lys48-linked polyubiquitin chains in vitro with human E1
(Wu et al., 2002).
The Ub2 purification method was a modification of a previously
published protocol (Chen & Pickart, 1990). After completion, the
synthesis reaction mixture was dialysed against 50 mM ammonium
acetate pH 4.5. The mixture was filtered and loaded at 1.0 ml min1
onto a Mono-S cation-exchange chromatography column (HR 5/5,
GE Healthcare). The polyubiquitin chains were then eluted with a
linear gradient of 0–0.4 M KCl over 60 ml. Elution fractions were
collected and further purified by size-exclusion chromatography on a
Superdex 75 16/60 column (GE Healthcare) equilibrated in crystal-
lization buffer (20 mM Tris–HCl pH 8.0, 50 mM NaCl, 0.01% NaN3).
The purity of the different polyubiquitin chains (Ub1, Ub2, Ub3 and
Ub4) was assessed by SDS–PAGE. The Ub2 concentration was
determined using UVabsorbance at 276 nm. The Mud1 UBA domain
(residues
293–332)
was
expressed
and
purified
as
described
previously (Trempe et al., 2005) and dialyzed against crystallization
buffer.
Cocrystallization trials of Mud1 UBA with Ub2 were performed at
a final concentration of 0.5 mM Ub2 and 0.5–0.75 mM Mud1 UBA
using Structure Screens 1 and 2 (Molecular Dimensions). Crystals
were grown at 295 K by vapour diffusion using the sitting-drop
method (1.0 ml drops). Thin rectangular plate-shaped crystals
(300 100 30 mm) were grown in 30% PEG 4000, 0.2 M Li2SO4,
0.1 M Tris–HCl pH 8.5 from a 1.5:1 molar ratio of UBA:Ub2.
Conditions with less or no Mud1 UBAyielded smaller crystals of poor
diffraction quality.
2.2. Data collection and processing
A crystal was cryoprotected using mother liquor supplemented
with 15% ethylene glycol and frozen in liquid nitrogen. Data were
collected at 100 K on beamline ID-29 at ESRF, Grenoble. Data-
collection statistics are shown in Table 1. Reflections were indexed
and integrated using the program MOSFLM (Leslie, 2006) and the
intensities were scaled and merged using SCALA (Evans, 2006).
2.3. Structure solution and refinement
The phase problem was solved by molecular replacement using
the program Phaser (McCoy et al., 2007). The crystal structure of
monoubiquitin (PDB code 1ubq; Vijay-Kumar et al., 1987) was used
as a search model, excluding the flexible residues 73–76. Six copies of
ubiquitin were found, giving a solvent content of 41%. After rigid-
body refinement in REFMAC5 (Murshudov et al., 1997), no addi-
tional density was observed that could accommodate the UBA
domain. Water molecules were added automatically using ARP/
wARP (Perrakis et al., 1997). The model was then adjusted in the
electron-density map using the program Coot (Emsley & Cowtan,
2004). The bulk solvent was modelled using the Babinet method with
a mask. After a few cycles of restrained refinement in REFMAC5
and model building, a final model was obtained with good overall
geometry and a satisfactory fit to the experimental amplitudes
(Table 1). The distal moieties of the three Ub2 molecules in the
asymmetric unit were named A, C and E and their respective cova-
lently bound proximal moieties were named B, D and F. The co-
ordinates and structure factors were deposited in the Protein Data
Bank under accession code 3m3j.
3. Results and discussion
The asymmetric unit of the new crystal form contained three Ub2
molecules, which all adopt the same conformation in which the
hydrophobic patches of the proximal and distal ubiquitin moieties,
centred around Ile44, interact with each other (Fig. 1a). Most
ubiquitin-binding domains interact with the hydrophobic patch of
ubiquitin (Hicke et al., 2005) and thus the conformation in which the
patch is buried will be referred to as the closed conformation. More
specifically, the side chains of Leu8, Ile44, His68 and Val70 of one
moiety fit snugly onto a surface formed by the same amino acids on
the other moiety (Fig. 1b). Moreover, the same seven hydrogen bonds
were found in each of the three distal–proximal pairs, notably
between the carbonyl O atoms of Gly47 and Leu71 and the backbone
amides of Leu71 and Gln49, respectively. The overall arrangement of
the distal and proximal moieties is thus remarkably similar among the
three Ub2 molecules in the asymmetric unit (Fig. 1c), with C root-
structural communications
Acta Cryst. (2010). F66, 994–998
Trempe et al.
Lys48-linked diubiquitin
995
Table 1
X-ray data-collection and refinement statistics for Ub2.
Values in parentheses are for the last shell.
X-ray source
ESRF ID29
Wavelength (A˚ )
0.97625
Space group
C2
Unit-cell parameters (A˚ , )
a = 58.7, b = 78.7, c = 93.1,
= = 90, = 97.9
Mosaicity ()
0.30
Images
180
Oscillation angle ()
1.0
Resolution (A˚ )
39.90–1.60 (1.69–1.60)
Unique reflections
54118 (7792)
Completeness (%)
97.9 (96.8)
Multiplicity
3.8 (3.8)
hIi/h(I)i
16.1 (3.2)
Rmerge†
0.057 (0.432)
Solvent content (%)
41
No. of reflections in Rfree set (5%)
2738
Rwork
0.183
Rfree
0.229
FOM
0.851
R.m.s. deviations from ideal values‡
Bond lengths (A˚ )
0.012
Bond angles ()
1.5
Torsion angles ()
6.1
Protein atoms
3962
Water atoms
360
Ligand atoms (1 ethylene glycol, 3 sulfate ions)
19
Disordered residues (not modelled)
Chain B, 76; chains D, F, 74, 75, 76§
Average B factors (A˚ 2)
Protein main chain
19
Protein side chain
21
Water
32
Ethylene glycol
28
Sulfate ions
58
Ramachandran outliers}
1 [Gln62 in chain D]
Estimated coordinate error†† (A˚ )
0.18
PDB code
3m3j
† P
hkl
P
i jIiðhklÞ hIðhklÞij=P
hkl
P
i IiðhklÞ, where Ii(hkl) is the intensity of the ith
measurement of reflection hkl and hI(hkl)i is the mean value for all i measure-
ments.
‡ Ideal values as reported in Engh & Huber (2001).
§ These residues
correspond to the C-termini of proximal ubiquitin moieties.
} Residues for which the
backbone torsion angles are outside the core region of the Ramachandran plot (Kleywegt
& Jones, 1996).
†† Coordinate error estimated from a Luzzati plot (R/Rfree versus
resolution) as reported by SFCHECK (Vaguine et al., 1999).
mean-square deviation (r.m.s.d.) values that are between 0.39 and
0.53 A˚ .
A previously reported crystal structure of Ub2 (Cook et al., 1992)
has a single molecule in the asymmetric unit, which also adopts the
closed conformation (Fig. 1c). C r.m.s.d. values of 0.68–0.89 A˚ were
calculated between the previous structure (PDB code 1aar; Cook et
al., 1992) and each of the Ub2 subunits in the new crystal structure.
The previous crystal form was obtained by crystallizing Ub2 in the
presence of 2-methyl-2,4-pentanediol (MPD) and sodium citrate at
pH 5.0, instead of PEG 4000, Li2SO4 and Tris at pH 8.5 as used in the
current study. Despite these different conditions, the same set of
hydrophobic interactions and hydrogen bonds were found as in the
previous Ub2 crystal structure. The closed conformation was also
observed in one of the crystal forms of Ub4 (Phillips et al., 2001) but
not in the other (Cook et al., 1994). Similar to the case reported here,
the more recent Ub4 crystal structure was obtained from a crystal
grown in the presence of a peptide derived from a ubiquitin-binding
protein (S5a), which was not incorporated into the crystal but yielded
Ub4 crystals in a different space group (Phillips et al., 2001). NMR
residual dipolar couplings and relaxation-anisotropy studies have
shown that the closed conformation of Ub2 predominates in solution
at pH values above 6.8 and is in rapid equilibrium with an open form
(Varadan et al., 2002). The solution structure of the closed confor-
mation, which was determined by a docking approach using chemical
shift perturbation data and residual dipolar coupling restraints (PDB
code 2bgf; van Dijk et al., 2005), superposes with an average C
r.m.s.d. of 1.5 A˚ with the three Ub2 conjugates observed in the
present crystal structure. This shows that the overall arrangement of
the Ub2 conjugate in the crystal is similar to that observed in solution.
Although Ub2 adopts the closed conformation in both crystal
forms (this study and Cook et al., 1992), differences are observed in
the configuration of the isopeptide linkage. Well defined electron
density was observed for the isopeptide linkage in the new crystal
structure (Fig. 2a), with B factors near main-chain levels for the atoms
involved (between 15 and 25 A˚ 2, compared with 10–20 A˚ 2 for main-
chain atoms). This contrasts with the previously published Ub2 crystal
structure, which showed slight disorder for these residues (B factors
of >30 A˚ 2, compared with 10–20 A˚ 2 for main-chain atoms), although
electron density was also visible for the isopeptide bond (Cook et al.,
1992). The crystal packing probably induces this order in the new
crystal form, since isopeptide linkages from molecules within or
between different asymmetric units make a number of reciprocal
interactions (Fig. 2b). The "-amide group of Lys48 in the distal
subunit (involved in the isopeptide bond) makes a hydrogen bond to
the backbone carbonyl O atom of Ala46 in a neighbouring subunit
and the side chain of Leu73 in the proximal subunit intercalates
between Leu71 and Leu73 in the neighbouring subunit (Fig. 2c).
These interactions were not observed in the previous structure owing
to different crystal packing. A network of intramolecular hydrogen
bonds and water molecules that were not observed in the previous
crystal structure further stabilizes the isopeptide-linkage conforma-
tion. A water molecule makes hydrogen bonds to the carbonyl O
atoms of Gly76 and Gln49 in the distal and proximal moieties,
respectively, and another water molecule bridges the side chain of
Glu51 with the carbonyl O atom of Gly76 (Fig. 2c). Finally, the
carbonyl O atom of Leu73 makes a hydrogen bond to the amide
group of Gly76 in the distal moiety. These interactions were observed
in all three isopeptide linkages in the asymmetric unit, which thus
adopt nearly identical conformations with residues 73–76 (distal) and
Lys48 (proximal) forming a long U-shaped loop (Fig. 1c). The con-
formation of the isopeptide linkage in the previous structure is
similar, but shows significant differences in the backbone torsion
structural communications
996
Trempe et al.
Lys48-linked diubiquitin
Acta Cryst. (2010). F66, 994–998
Figure 1
Crystal structure of Lys48-linked Ub2. (a) Cartoon representation of a Ub2 molecule in the crystal structure. The proximal and distal moieties are coloured magenta and cyan,
respectively. The atoms forming the isopeptide bond as well as the interface residues Ile44 and Val70 are shown as sticks. Residues labelled with primes belong to the distal
moiety. (b) Close-up view of the residues forming the interface between the distal and proximal subunits. The molecular surface of the proximal subunit is displayed in
transparent white. (c) Cross-eye stereoview ribbon display of the overlaid Ub2 crystal structures. The three chains in the new crystal structure are shaded yellow, blue and red
for A–B, C–D and E–F, respectively. The previously reported crystal structure of Ub2 is shaded in magenta (PDB code 1aar; Cook et al., 1992). Residues that have different
conformations in different subunits are labelled. The disordered C-termini of the proximal moieties are labelled ‘C’.
angles for residues 73–76 (Fig. 3a). The isopeptide bond is in a trans
configuration in both crystal structures, but the carbonyl O atom of
Gly76 points in opposite directions, which imposes a reconfiguration
of Gly75 and Gly76. This emphasizes the flexibility of the isopeptide
linkage, which is essential for the function of Ub2 because ubiquitin-
binding domains need to access the hydrophobic patches of ubiquitin
that are occluded in the closed conformation (Fig. 1a). Solution NMR
dynamics studies have indeed shown that the closed conformation
of Ub2 experiences fast interdomain motion on a 10 ns timescale
(Ryabov & Fushman, 2006).
Additional differences are found in the backbones of different Ub2
subunits, notably at the free C-termini of the proximal moieties (B, D
and F), which show variable levels of disorder for residues Arg74–
Gly76 (Fig. 1c and Table 1). The loop residues Thr9 and Gly10, which
are located between the 1 and 2 strands, also adopt a different
conformation in chain B compared with the other chains (Figs. 1c and
3b) and the electron density around these residues is weaker in chain
B in comparison with the other chains. In the previous crystal
structure this loop adopts the conformation observed in chains A, C,
D, E and F in the new crystal structure. Interestingly, the chemical
environment around Thr9 and Gly10 is nearly identical for all chains,
including chain B, with Thr9 being in proximity to Ala46/Gly47 and
Ser57/Asp58 in two different neighbouring subunits (not shown).
This suggests that the two conformations observed have similar
potential energy, with the most frequent being slightly more
stable. This loop shows significant backbone dynamics in solution
(Lakomek et al., 2006), which is consistent with the variability
observed here.
structural communications
Acta Cryst. (2010). F66, 994–998
Trempe et al.
Lys48-linked diubiquitin
997
Figure 2
Conformation of the isopeptide bond in the crystal structure of Ub2. (a) Cross-eye stereoview of the A-weighted 2Fo Fc electron-density map at the isopeptide linkage
contoured in blue at 0.35 e A˚ 3. The atomic model is drawn as sticks. Water molecules are drawn as red spheres. (b) The three Lys48-linked Ub2 molecules in one asymmetric
unit are coloured yellow for chains A–B, blue for chains C–D and red for chains E–F. Distal (A, C and E) and proximal (B, D and F) ubiquitin moieties are distinguished by
pale and dark shades, respectively. Chains C0 and D0 are from an adjacent asymmetric unit and are labelled in pale and dark cyan, respectively. The isopeptide linkages are
shown as spheres coloured by atom type (white, carbon; blue, nitrogen; red, oxygen). (c) Cross-eye stereoview of the isopeptide bond and its interactions. Residues labelled
with primes belong to a distal moiety. Hydrogen bonds are shown as dashed lines. C atoms of chains A–B and E–F are shown in yellow and salmon red, respectively.
4. Conclusions
A new crystal form of Lys48-linked Ub2 was obtained and its struc-
ture was determined by X-ray crystallography to 1.6 A˚ resolution.
The asymmetric unit is composed of three Ub2 molecules that all
adopt the closed conformation, as observed in solution (Varadan et
al., 2002) and in the previous crystal structure (Cook et al., 1992),
despite the different crystallization conditions and crystal packing.
The new crystal form reveals a new conformation for the isopeptide
linkage, which interacts with other isopeptide linkages in the other
subunits. A new conformation was also observed for the loop
between the 1 and 2 strands. These local differences emphasize the
flexibility of the isopeptide linkage and the 1–2 loop.
We would like to thank Professor Kazuhiro Iwai for providing the
recombinant baculovirus to express mouse E1, Dr Randy Poon for
constructs for the expression of human E2 and the staff at ESRF
beamline ID29 for providing excellent facilities for data collection.
This work was supported by an MRC grant to JAE and Wellcome
Trust and British Overseas Research Studentships to JFT.
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structural communications
998
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Lys48-linked diubiquitin
Acta Cryst. (2010). F66, 994–998
Figure 3
Comparison of loop conformations in different Ub2 crystal structures. (a) Comparison of the isopeptide-bond conformation in the two Ub2 crystal structures. Chains A–B of
the new crystal structure are coloured yellow and the previous structure (PDB code 1aar; Cook et al., 1992) is coloured magenta. Residues labelled with primes belong to a
distal moiety. The conformation of the isopeptide bond in chains C–D and E–F is similar to that in chains A–B. (b) Comparison of the 1–2 loop conformation in chain B
(yellow) and the previous crystal structure (magenta). The conformation of this loop in chains C–D and E–F of the new structure is similar to that shown in magenta.
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3M3K
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Ligand binding domain (S1S2) of GluA3 (flop)
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The molecular mechanism of flop-selectivity and subsite
recognition for an AMPA receptor allosteric modulator:
Structures of GluA2 and GluA3 complexed with PEPA
Ahmed H. Ahmed§, Christopher P. Ptak§, and Robert E. Oswald*
Department of Molecular Medicine, Cornell University, Ithaca, NY 14853 USA
Abstract
Glutamate receptors are important potential drug targets for cognitive enhancement and the
treatment of schizophrenia in part because they are the most prevalent excitatory neurotransmitter
receptors in the vertebrate central nervous system. One approach to the application of therapeutic
agents to the AMPA subtype of glutamate receptors is the use of allosteric modulators, which
promote dimerization by binding to a dimer interface thereby reducing desensitization and
deactivation. AMPA receptors exist in two alternatively spliced variants (flip and flop) that differ
in desensitization and receptor activation profiles. Most of the structural information on
modulators of the AMPA receptor target the flip subtype. We report here the crystal structure of
the flop-selective allosteric modulator, PEPA, bound to the binding domains of the GluA2 and
GluA3 flop isoforms of AMPA receptors. Specific hydrogen bonding patterns can explain the
preference for the flop isoform. This includes a bidentate hydrogen bonding pattern between
PEPA and N754 of the flop isoforms of GluA2 and GluA3 (the corresponding position in the flip
isoform is S754). Comparison with other allosteric modulators provides a framework for the
development of new allosteric modulators with preferences for either the flip or flop isoforms. In
addition to interactions with N/S754, specific interactions of the sulfonamide with conserved
residues in the binding site are characteristics of a number of allosteric modulators. These, in
combination, with variable interactions with five subsites on the binding surface lead to different
stoichiometries, orientations within the binding pockets, and functional outcomes.
Membrane receptors are the cell's gatekeepers, allowing chemical signals access to the cell's
pathways. Through the binding of endogenous ligands, receptors identify relevant
environmental cues and facilitate cell-cell communication. The regulation of membrane
receptors has become an important goal of drug discovery efforts (1,2). By targeting the
physiological (orthosteric) ligand-binding site, agonists and antagonists control the function
of membrane receptors. Unfortunately, exogenously induced agonist-activation at the
orthosteric site can cause toxic effects from overstimulation. Allosteric modulator binding
sites use a distinct avenue for altering the natural response of a receptor. The ability of some
allosteric modulators to enhance receptor stimulation, while not actually providing the
trigger for stimulation, is a clear advantage that conserves the endogenous signaling
pathway. Being important mediators of higher-order processes such as learning and
memory, ionotropic glutamate receptors (iGluRs) have attracted a great deal of interest as
allosteric modulator targets (3–6). Of clear therapeutic importance, various
neurodegenerative disorders such as Parkinson's and Alzheimer's diseases, Huntington's
*Corresponding author; telephone: 1-607-253-3877; fax: 1-607-253-3659; email: reo1@cornell.edu.
§These authors contributed equally to this work.
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Published in final edited form as:
Biochemistry. 2010 April 6; 49(13): 2843–2850. doi:10.1021/bi1000678.
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chorea, and neurologic disorders including epilepsy and ischemic brain damage have been
linked to iGluRs (7).
The crystal structure of GluA2 (8) clarifies years of speculation on the complex arrangement
of the glutamate receptor's four subunits (9). The GluA2 can be dissected into 3 functionally
distinct layers. Farthest from the membrane, the amino terminal domain (ATD) can act as a
peripheral regulatory domain but is also involved in assembly and trafficking (10,11).
Sandwiched between the ATD and the membrane domain, the ligand-binding domain (LBD)
recognizes the neurotransmitter signal and directly regulates receptor activation (12).
Structures for both isolated extracellular domains (ATD and LBD) reveal a dimeric
organization (13–15). At the membrane interface, two alternative linker conformations
transition the 2-fold symmetry, which is adopted by both extracellular domains, into the 4-
fold symmetry of a membrane-traversing cation-selective channel (8,16). For iGluRs, the
ion channel domain confers functional relevance with its ability to selectively conduct the
flow of ions across the cell's membrane. The layers of extracellular domains, each with the
potential for multiple control points, allosterically regulate the ion channel domain's function
(8). Therefore it is not surprising that the ATD, the LBD, and the LBD-channel linker have
all been shown to be effective targets of allosteric modulators (13,17,18).
Since the structures of the ATD and the full iGluR channel have only recently been solved,
allosteric drug-binding sites external to the LBD have not been fully explored in molecular
detail. However, the decade-old LBD structure has proved to be indispensable as a heavily
exploited scaffold for understanding agonist, partial agonist, and antagonist binding
interactions as well as their ability to regulate channel gating behavior (12,19,20). Although
the dimeric organization is consistent across all iGluR subtypes, the molecular details of
LBD-agonist specificity define the subtype families into N-methyl-D-aspartic acid (NMDA)
receptors (21), α-amino-3-hydroxy-5-methyl-4-isoxazole-propionic acid (AMPA) receptors
(12), and kainate receptors (22). Because all subtypes are constrained by their conserved
sensitivity to glutamate stimulation, diversity at the orthosteric site is evolutionarily limited
and most agonists display cross-subtype activity. An allosteric modulator-binding site within
the quaternary LBD structure is located along the dimer interface (18) and offers improved
discrimination by modulators. Drugs that bind to the allosteric sites on the LBD dimer
interface can enhance the activity of iGluRs (23) and increase performance on tests of
memory (24). Except for the LBD structures with modulatory ions bound to the dimer
interface (25–27), only LBD structures from the AMPA receptor subtype, GluA2, have been
reported with bound allosteric modulators (18,28–31). Within the structures, the bound
modulatory drugs stabilize the LBD dimer interface, which is required for activation of the
ion channel and is dissociated during desensitization (18).
Although the residues that line the allosteric modulator-binding pocket do not differ between
AMPA receptors subtypes (GluA1–4), the ability of allosteric modulators to stabilize the
activated state still varies (32,33). Also, AMPA receptors can be alternatively spliced into
what is referred to as flip and flop isoforms (34). Modulator selectivity (23), desensitization
(35), and channel closing rates (36) differ between flip and flop. Although several of the
amino acid differences between the two forms are located in or near the allosteric
modulator-binding site, the difference at position 754 (serine in flip, asparagine in flop)
seems to be entirely responsible for the functional differences between allosteric modulator
regulation of the flip and flop variants (23,28,32). Cyclothiazide (CTZ) and some other
thiazide derivatives have improved binding to the flip form due to a hydrogen bond between
S754 and the NH of the fused thiazide ring (28). In the case of the flop form, the
alternatively spliced sequence places an asparagine in the 754 position, which is not
optimally positioned to form a hydrogen bond. Sekiguchi et al. (33) introduced an allosteric
modulator of AMPA receptors (4-[2-(phenylsulphonylamino)ethylthio]-2,6,-
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difluorophenoxyacetamide, PEPA) with a preference for the flop form. In fact, the relative
sensitivity of CTZ to PEPA has been used as a diagnostic for the prevalence of flip vs. flop
versions of AMPA receptor in particular cell types (37). PEPA shows potential in treatment
of post ischemic memory impairment (38) and contextual fear (39) but despite PEPA's
unique flop sensitivity, the modulator has not yet been used as a lead compound in SAR
studies.
For drug discovery to be guided by structures, understanding the possible molecular
interactions between modulators and the dimer interface is essential. We have shown
previously (31) that changes in the structures of CTZ derivatives can reorient the modulator
within the binding site. Subsequently, we proposed that the allosteric modulator site is
comprised of 5 subsites (Figure 1C). In the present study, we determine the three
dimensional structures of PEPA bound to the GluA2o and GluA3o LBDs (flop forms), and
use PEPA's binding interactions to further characterize the subsite specific binding
properties displayed by allosteric modulators. The amide group of PEPA makes a direct
hydrogen bond to N754, explaining the preferential action of PEPA on the flop form of
AMPA receptors. Another key structural element, the sulfonamide group of PEPA, is
conserved with the biarylsulfonamide class of allosteric modulators (6) and interacts with
the same residues of the dimer interface (8,30). Although previously classified as unrelated,
PEPA and the large group of biarylsulfonamide have similarities, which suggest that specific
PEPA groups (particularly the unique flop-interacting amide) can be strategically integrated
into biarylsulfonamide SAR studies.
Experimental Procedures
Materials
PEPA was purchased from Tocris (Ellisville, MO). The GluA2 S1S2J construct was
obtained from Eric Gouaux (Vollum Institute; 12).
Protein Preparation and Purification
GluA2 S1S2 consists of residues N392 - K506 and P632 - S775 of the full rat GluA2o
subunit (40), a `GA' segment at the N-terminus, and a `GT' linker connecting K506 and
P632 (12). A similar construct of GluA3 S1S2 was prepared as described previously (41).
pET-22b(+) plasmids were transformed in E. coli strain Origami B (DE3) cells and were
grown at 37°C to OD600 of 0.9 to 1.0 in LB medium supplemented with the antibiotics
(ampicillin and kanamycin). The cultures were cooled to 20°C for 20 min. and isopropyl-β-
D-thiogalactoside (IPTG) was added to a final concentration of 0.5 mM. Cultures were
allowed to grow at 20°C for 20 h. The cells were then pelleted and the S1S2 protein purified
using a Ni-NTA column, followed by a sizing column (Superose 12, XK 26/100), and
finally an HT-SP-ion exchange-Sepharose column (Amersham Pharmacia). Glutamate (1
mM) was maintained in all buffers throughout purification. After the last column, the protein
was concentrated and stored in 20 mM sodium acetate, 1 mM sodium azide, and 10 mM
glutamate at pH 5.5.
Crystallography
For crystallization trials, the protein was concentrated to 0.2 – 0.5 mM in 10 mM glutamate
using a Centricon 10 centrifugal filter (Millipore, Bedford, MA). For the PEPA-bound
structures, PEPA was added to 5 mM. The final protein concentration was 0.2 to 0.3 mM.
Crystals were grown at 4°C using the hanging drop technique, and the drops contained a 1:1
(v/v) ratio of protein solution to reservoir solution. The reservoir solution contained 14–15%
PEG 8K, 0.1 M sodium cacodylate, 0.1–0.15 M zinc acetate, and 0.25 M ammonium sulfate,
pH 6.5.
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Data were collected at the Cornell High Energy Synchrotron Source beam line A1 using a
Quantum-210 Area Detector Systems charge-coupled device detector. Data sets were
indexed and scaled with HKL-2000 (42). Structures were solved with molecular
replacement using Phenix (43). Refinement was performed with Phenix (43), and Coot 0.5
(44) was used for model building.
Results
Structure of PEPA bound to GluA2 S1S2 flop
The structure of glutamate bound to GluA2o S1S2 (3dp6; 41) was used as the initial search
probe for the molecular replacement solution of PEPA bound to GluA2o S1S2 with
glutamate in the agonist-binding site. PEPA was then modeled into two symmetrical
positions within the density found at the dimer interface, and the structure was optimized
using Phenix (43). The refinement statistics are given in Table 1. The resolution is 1.85 Å,
and three unique copies are found in the unit cell. The overall structure of the S1S2 domain
is very similar to the structure in the absence of PEPA, with contacts between glutamate and
the protein unchanged. However, PEPA clearly binds within the dimer interface, making
contacts with both monomers within the dimer. As shown in Figure 1, one PEPA molecule
binds per dimer interface. However because the dimer interface is symmetrical, two
equivalent orientations (related by a 180° rotation) are possible. Electron density for both is
seen in the crystal structure, although the intensity of one orientation is greater than the
other.
The binding of PEPA to the dimer interface increases the distance between the two
monomers that form the dimer by approximately 1.5 Å. This allows the relatively large
PEPA molecule to fit within the interface, but also increases the separation between the
linkers to the ion channel (the distance increases from 39.4 Å to 41 Å; Figure 1A). Relative
to the core of Lobe 1, both the J/K helices and one β strand (P105-G110) connecting the two
lobes are displaced slightly away from the dimer interface (Figure 1B). In addition, Lobe 2
is slightly twisted relative to glutamate-bound S1S2 in the absence of PEPA (3dp6; 41).
PEPA binds at the bottom of a water-filled, inverted U-shaped cleft with five subsites (A, B/
B′, and C/C′; 31). Upon binding, crystallographic waters are displaced from the central A
subsite and more buried C/C′ sites, with the waters in the B/B′ subsite remaining (Figure
1C). This displacement of presumably ordered water would be likely to contribute a
favorable entropy component to binding.
The sidechains of P494 are at the center of the interface and the edge of the two proline
rings from each monomer form the base of the binding site in which the difluorophenyl ring
resides (Figure 2A). This is close to the position of the methoxybenzoyl ring of aniracetam
in its structure bound to GluA2-S1S2(FW) (29). The other side of the ring is exposed to
S497 and S729. The sidechain hydroxyl of S497 is oriented toward the dimer interface in the
absence of PEPA, but rotates out toward the solvent to accommodate the difluorophenyl ring
of PEPA (Figure 1B). The amide of PEPA is involved in a network of hydrogen bonds with
sidechain hydroxyl of Y424, the backbone carbonyl of F495, the sidechain carboxyl of
D760, the sidechain amide of N754, and two water molecules (Figure 2A). The most
striking of these hydrogen bond pairs is with N754. This represents the only difference
between the flip (S754) and flop (N754) isoforms in the PEPA binding site and is almost
certainly a major source of the preference for the flop isoform. The phenyl-sulfonylamide
side of PEPA inserts into a hydrophobic pocket formed by sidechain methyls of I481 and
L751 as well as methylene groups contributed by K493, N754, and E755 (Figure 2B). It is
possible that the contribution by methylene group of N754 provides a more hydrophobic
pocket than S754 in the flip form, further contributing to the preference for the flop form.
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Because the dimer interface is symmetrical, PEPA can bind in two orientations and both are
observed in the crystal. For this reason, changes in the protein due to a specific interaction
with PEPA can be partially masked because each monomer is a weighted average of two
orientations of bound PEPA. However, one orientation has a stronger density than the other,
providing some insight into the extent of changes in the dimer interface that are produced by
PEPA binding. As shown in Figure 2C, the two monomers comprising the dimer differ more
within the PEPA binding site than the corresponding monomers in the absence of PEPA.
One turn of helix J (L751 to N754) contains important determinants for both orientations of
PEPA. In one orientation the amide group of PEPA interacts with the sidechain of N754,
and in the other, the aromatic ring of PEPA inserts between a hydrophobic pocket formed by
the sidechain of L751 and the methylene group of N754. In the orientation for which the
density of the amide of PEPA is stronger, N754 is better positioned to form an H-bond
(Figure 2D); whereas, in the other side of the interface, N754 is oriented to form an H-bond
with the carbonyl of S729. This change in orientation facilitates the insertion of the aromatic
ring of PEPA into the hydrophobic pocket, which is accompanied by a small shift in the
sidechain of L751 to accommodate the aromatic ring (Figure 2D). Since these structures are
weighted averages, it is possible that the actual positions of these sidechains involve an even
greater movement than is seen from the asymmetry of the crystal.
Structure of PEPA bound to GluA3 S1S2 flop
In studies of the physiological effects of PEPA, a significant difference between subtypes
has been observed, with GluA3 being most susceptible to modulation (33). The structure of
GluA3i S1S2 bound (flip form) to glutamate has been reported previously (41). Since PEPA
preferentially binds to the flop form, the GluA3o structure was determined bound to
glutamate with and without PEPA (Figure 3A). Like GluA2o, in the absence of PEPA,
GluA3o has three copies in the asymmetric unit. Comparing lobe closure between GluA3i
and GluA3o, the flop form is slightly more closed (1.6° ± 0.7°).
In the presence of PEPA, GluA3o was present in one copy in the asymmetric unit, and PEPA
was observed with the same density in two symmetrical orientations. Like GluA2o bound to
PEPA, the dimer interface (assessed using the symmetrical molecule in the crystal) was
displaced relative to the unbound from (Figure 3A) by approximately 2.5 Å at the position
of the linker replacing the ion channel domain. Within the binding site, three sidechains
exhibited different rotamers compared with the GluA2o structure bound to PEPA (Figure
3B). For PEPA-bound GluA3o, both S497 and S729 assumed rotameric states that differed
both from GluA2o bound to PEPA and from GluA2o and GluA3o in the absence of PEPA. In
the case of S729, the rotameric state in combination with a slight movement of the amide of
PEPA (relative to the GluA2o structure) would make an H-bond with the sidechain of S729
(shown in Figure 2A for GluA2o) unlikely. In the case of N754, the sidechain is displaced
relative to the GluA2o-PEPA structure so that only one H-bond is made to the amide of
PEPA. This may be a result of averaging of the two orientations of PEPA only one of which
forms a bidentate H-bond with N754.
Discussion
The goal of allosteric modulation, like orthosteric modulation, is often to stabilize a
conformational state of a dynamic protein (45). The activated state of iGluRs is naturally
unstable allowing the channel to desensitize (46). Disruption of the symmetrical dimer
interface between LBDs is thought to initiate desensitization-mediated channel closure (47).
By maintaining the LBD dimer, positive allosteric modulators can prevent desensitization
and prolong activation (18). Currently, 15 crystal structures of the GluA2 LBD with bound
allosteric modulators are deposited in the Protein Data Bank (48). All of these modulators
bind to a large crevice with 2-fold symmetry along the symmetric dimer interface (18). The
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large variation in structure among allosteric modulators results in significant variations in
binding orientations and interactions. At least four distinct binding modes have been
identified: (1) A-subsite class (aniracetam, CX614 (29)), (2) classical thiazide (cyclothiazide
(18), TCMZ, ALTZ (31)), (3) the shifted thiazide class (IDRA-21, HCMZ, HFMZ; (31)),
and (4) the full spanning class (PEPA (this paper), dimeric biarylpropylsulfonamide (30),
LY404187 (8)). Overlaying modulators from these structural classes has led to the proposal
that the allosteric modulator site is comprised of a series of subsites (Figure 1C;31).
Positioned at the center of the binding-site, the symmetric A subsite is narrow and allows
entrance to only one molecule. Two subsites (B and C) lie at each end of the A subsite with
the hydrophobic C subsite located more deeply in the pocket effectively defining five
subsites (A, B, B′, C, and C′).
In the open state, the subsites are filled with water, which may act to weakly stabilize the
dimer. Allosteric modulators generate stronger interactions across the subsites thereby
increasing the linkages between the monomers. The simplest modulator class, including
aniracetam and CX614, fills the A subsite with one molecule but does not enter the
peripheral B and C subsites (29). The two classes of thiazide-based modulators account for
10 of the 15 solved allosteric modulator-GluA2 crystal structure complexes (18,28,31). The
classical thiazide (CTZ-like) binding class and the shifted thiazide (IDRA-21-like) binding
class are positioned respectively in the B and C subsite or mainly the C subsite. Most of the
thiazide modulators do not extend across the A subsite and therefore can bind two molecules
per dimer. However, a few of the newly described shifted thiazides (HFMZ, HCTZ; 31)
enter the A subsite but only enough to impair binding of a second modulator. The dimeric
biarylpropylsulfonamide compound ((R,R)-N,N-(2,2'-[Biphenyl-4-4'-Diyl]Bis[Propane-2,1-
Diyl]) Dimethanesulfonamide) described by Kaae et al. (30) was the first allosteric
modulator shown by crystallography to extend along the entire length of the inner dimer
cavity from C to C′ subsites. PEPA also interacts with J helices from both monomers, which
cap the ends of the modulator-binding pocket. The density occupied by both symmetrical
copies of PEPA overlays the dimeric biarylsulfonamide compound as both modulators
represent the full spanning class (Figure 4B).
The GluA flip and flop splice variants differ by only a few residues along the J helix in the
LBD; however, residue 754 (Asn in flop and Ser in flip) is positioned between the B and C
subsites. For thiazides, a clear preference in binding to the flip-form is mediated by a
hydrogen bond between the hydrobenzothiadiazide ring and S754 (28). In contrast, PEPA is
flop-selective and the PEPA-bound structure provides the first structure containing a direct
interaction between a modulator and the flop form's N754. The amide of PEPA extends
straight out from the A subsite and across the B and C subsite interface to make an amide-
amide hydrogen bond with N754 (Figure 2A). Unlike most other AMPA modulators, PEPA
fills neither the B nor the C subsites but interacts directly with the J helix. A similar
interaction is seen with LY404187 (49) bound to GluA2i (8). Strong hydrogen bonding can
occur between two amides (50) and has been shown to be responsible for driving
oligomerization of transmembrane leucine zippers (51). The distances between the
interacting amides in the PEPA-bound structure support a bidentate hydrogen-bonding
pattern, which is much stronger and more specific than a typical hydrogen bond. While
PEPA is selective for the AMPA receptor's flop form, a weaker but still existent potentiation
of the flip form has been observed (33,52). Replacing N754 (flop) with S754 (flip) would
not prevent PEPA from binding; however, serine would provide only one hydrogen-bonding
partner for PEPA's amide with an extended interaction distance. In contrast, LY404187
displays a preference for the flip isoform (53), and its cyano group extends out to interact
directly with S754. The cyano-S754 interaction is a clear flip analog of the flop-selective
PEPA amide-N754 interaction (Figure 4A).
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Opposite to the amide on the PEPA molecule, a sulfonamide is tethered to the
difluorophenyl ring (Figure 4A). Within the dimer interface, the sulfonamide is positioned
so the nitrogen can hydrogen bond directly with the carbonyl of P494 (Figure 4C). A
sulfonamide oxygen points toward the amide nitrogen of G731. The angle of the peptide
plane is perpendicular to the sulfonamide oxygen, making a hydrogen bonding interaction
unlikely (Figure 4D). Instead, a dipole-dipole or charge-dipole interaction may occur. The
amide nitrogen of a polypeptide supports at least a partial positive charge (54), which would
interact with the strongly electronegative sulfonamide oxygen (55). Interestingly, both the
dimeric biarylsulfonamide (30) and LY404187 (8), other members of the full spanning
modulator class, also have a sulfonamide that interacts with the same backbone atoms of
P494 and G731 as PEPA (Figure 4C).
A large number of biarylsulfonamides have been identified that modulate AMPA receptors
and are being evaluated for therapeutic use in the treatment of depression and Parkinson's
disease (56). The conserved sulfonamide reveals a previously unidentified relationship
between PEPA and the biarylsulfonamide modulators. When the perpendicular peptide bond
plane including G731 is fixed, the sulfonamide on three overlayed modulators varies by 1.2
Å along the length of the interface with the PEPA sulfonamide being positioned closer to the
A subsite (Figure 4D). A shift of the sulfonamide also results in a shift in the corresponding
P494 across the interface presumably to maintain the hydrogen bond with the modulator's
amine. The sulfonamide forms an important bridge between the two dimer halves. For
PEPA, a phenyl-sulfonamide replaces the methyl-sulfonamide in the dimeric
biarylsulfonamide and fits snuggly against L751. Based on the orientation-induced
asymmetry within the GluA2-complex structure, the phenyl pushes the J helix away from
PEPA thereby affecting the C subsite (Figure 2C and D). Residues lining the C subsite are
on the same beta strand as G731, which must shift if the C subsite is to remain together and
presumably explain the 1.2 Å shift relative to the dimeric biarylsulfonamide. In fact, the
same phenyl-sulfonamide group substitution in a biarylpropylsulfonamide decreases the
modulatory effect of the derivative in SAR studies (57). For biarylpropylsulfonamides, the
optimal sulfonamide substitution was found to be either an ethyl or an iso-propyl group,
which should both fit without significantly disrupting the J helix or C subsite (57).
The PEPA-bound crystal structure from AMPA receptor subtypes, GluA2 and GluA3, do
not display major differences in binding interactions even though PEPA exhibits a stronger
effect on GluA3 (33). For GluA2, an asymmetry in the receptor-binding pocket was
observed while no significant difference in PEPA density was seen for the each orientation
within the GluA3 crystal structure. In addition, a number of side chains exhibit different
rotameric states between the two structures, although it is unlikely that these small changes
significantly impact the differential effects on the two subtypes. Although no structural
differences have been identified between GluA2 and GluA3 that would obviously impact
PEPA affinity, the possibility exists that subtle differences arising from the sequence
differences peripheral to the binding site may be important as has been described in the case
of the agonist binding site of GluA4 (58).
We have explored how PEPA (this paper) and other allosteric modulators (31) interact with
the GluA interface in the context of drug design. Together the identification of a conserved
group between PEPA (this paper) and biarylpropylsulfonamides (8,30) and the regional
nature of various subsite-functional group interactions provide a backdrop to extend
biarylpropylsulfonamide SAR studies (57) to include PEPA and biarylpropylsulfonamide
chimeras. Although optimizing the stability of the dimer interface provides a starting point
for SAR studies, additional constraints should be considered including the ability of the
modulator to enter the cavity, the dynamic structure of the dimer interface during closed,
open, and desensitized state transitions, and the ability of the modulator to cross the blood-
Ahmed et al.
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brain barrier before being metabolized. This definition of the allosteric modulator binding-
site should provide guidance in glutamate receptor allosteric modulator pharmacology.
Acknowledgments
We thank Prof. Eric Gouaux (Vollum Institute) for the GluA2 S1S2J construct, and Prof. Linda Nowak (Cornell)
for the full-length GluA3 construct.
This work was supported by a grants from the National Institutes of Health (R01-GM068935, R01 NS049223, and
R21 NS067562). This work is based upon research conducted at the Cornell High Energy Synchrotron Source
(CHESS), which is supported by the National Science Foundation under award DMR 0225180, using the
Macromolecular Diffraction at the CHESS (MacCHESS) facility, which is supported by award RR-01646 from the
National Institutes of Health, through its National Center for Research Resources.
Abbreviations
ALTZ
althiazide
AMPA
α-amino-3-hydroxy-5-methyl-4-isoxazole-propionic acid
CLTZ
chlorothiazide
CX614
pyrrolidino-1,3-oxazino benzo-1,4-dioxan-10-one
CTZ
cyclothiazide
FW
(S)-5-fluorowillardiine
flip and flop
alternatively spliced versions of AMPA receptors that vary in rates of
desensitization and sensitivity to allosteric modulators
iGluR
ionotropic glutamate receptor
GluA1-4
four subtypes of AMPA receptor
HCTZ
hydrochlorothiazide
HFMZ
hydroflumethiazide
IDRA-21
7-chloro-3-methyl-3,4-dihydro-2H-benzo[e][1,2,4]thiadiazine 1,1-dioxide
IPTG
isopropyl-β-D-thiogalactoside
LY404187
N-[2-(4′-cyanobiphenyl-4-yl)propyl]propane-2-sulfamide
PEPA
4-[2-(phenylsulphonylamino)ethylthio]-2,6,-difluorophenoxy acetamide
NMDA
N-methyl-D-aspartic acid
S1S2
extracellular ligand-binding domain of GluA2 and GluA3
SAR
structure-activity relationships
TCMZ
trichlormethiazide
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Figure 1.
(A) Comparison of glutamate-bound GluA2o S1S2 in the presence (blue) and the absence of
PEPA (green) in two orientations. Both orientations of PEPA are shown. Note that the
binding of PEPA results in a separation of the two components of the dimer (distance
between the Cα atoms of the threonine in the linker) by approximately 1.5 Å. (B) One
monomer of GluA2o S1S2 in the presence (blue) and the absence of PEPA (green) with one
orientation of PEPA shown. Both the J/K helices and the strand near S497 are displaced
upon binding PEPA. Also, the sidechains of S497 and S729 change rotameric states. (C)
Comparison of the water molecules at the dimer interface in the presence (tan spheres) and
the absence of PEPA (red spheres). PEPA is shown in both orientations. Despite the greater
separation of the dimer interface, a number of the ordered water molecules found in the
absence of PEPA are displaced by PEPA. The black circles delineate subsites of the
allosteric modulator binding site as described previously (31).
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Figure 2.
The PEPA binding site, emphasizing the important interactions, shown in two orientations.
(A) A view of the amide side of PEPA bound to GluA2 S1S2. The hydrogen bonding
network with the amide of PEPA is shown as dotted lines. The H-bond with the sidechain of
S729 is difficult to display in the orientation used in the figure. (B) A view of the phenyl
group of PEPA inserted into a hydrophobic pocket in GluA2 S1S2. (C) RMS plot showing
more variability in the J/K helices for the PEPA-bound structure than the unbound structure.
(D) J/K helix showing where differences in the two orientations were analyzed. The amide
of PEPA-N754 interaction (blue) maintains the position of the J helix in the absence of
PEPA (green) The J helix is displaced on the phenyl side of PEPA (red).
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Figure 3.
(A) Comparison of glutamate-bound GluA3o S1S2 in the presence (blue) and the absence of
PEPA (green) in two orientations. Both orientations of PEPA are shown. Note that the
binding of PEPA results in a separation of the two components of the dimer (distance
between the Cα atoms of the threonine in the linker) by approximately 2.5 Å. (B) One
monomer of GluA3o S1S2 in the presence (blue) and the absence of PEPA (green) with one
orientation of PEPA shown. Shown for comparison is the PEPA-bound form of GluA2o
(red). Both the J/K helices and the strand near S497 are displaced upon binding PEPA for
both GluA2o and GluA3o. Also, the sidechains of S497 and S729 are in different rotameric
states for GluA3o bound to PEPA compared with GluA3o in the absence of PEPA and
GluA2o bound to PEPA. Also, N754 is displaced in PEPA-bound GluA3o, such that only
one H-bond is possible with the amide of PEPA.
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Figure 4.
(A) Members of the full spanning class of allosteric modulators. The shape-highlighted
regions of the modulators illuminate key contact points to the specific binding pocket
residues and subsites (as labeled for PEPA). (B) Overlay of the full spanning modulator
structures. The structures were aligned at both sets of P494 and G731 residues. PEPA (gray)
occupies a similar arrangement of subsites as the dimeric biarylsulfonamide (PDB entry
3bbr, cyan, 30) and LY404187 (PDB entry 3kgc, magenta, 8). (C) The sulfonamide bridges
the two monomers in both PEPA and the dimeric biarylsulfonamide with the same
interactions to P494 and G731. (D) The hydrogen bond between the carbonyl of P494 and
the sulfonamide is maintained when the modulator is in a shifted position relative to the
peptide plane of K730 and G731 (green disk) located on the opposite monomer.
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Table 1
Structural Statistics
GluA2o (PEPA)
GluA3o (PEPA)
GluA3o
Space Group
P22121
P222
P222
Unit Cell (Å)
a=47.13 b=113.92 c=164.81
a=46.95 b=52.26 c=115.98
a=46.03 b=110.33 c=161.192
X-ray source
CHESS (A1)
CHESS (A1)
CHESS (A1)
Wavelength (Å)
0.977
0.977
0.977
Resolution (Å)
50–2.0 (2.03–2.00)
50–2.5 (2.54–2.00)
50–1.85 (1.88–1.85)
Measured reflections (#)
817961
62549
344340
Unique reflections (#)
70175
9141
69379
Data redundancy
6.9 (7.1)
6.0 (4.2)
4.7 (3.0)
Completeness (%)
99.9 (100.0)
99.5 (89.8)
96.5 (73.5)
Rsym (%)
11.4 (34.2)
13.0 (45.5)
7.5 (24.6)
I/σi
33.3 (7.1)
19.2 (2.5)
34.3 (3.3)
PDB ID
*
*
*
Current Model Refinement Statistics
Phasing
MR
MR
MR
Molecules/AU
3 (no NCS applied)
1
3 (no NCS applied)
Rwork/Rfree (%)
18.7/24.2
18.9/28.5
20.1/23.4
Free R test set size (#/%)
2000 (2.85)
914 (10.0)
2000 (2.88)
Number of protein atoms
5979
2030
6091
Number of heteroatoms
111
62
30
Rmsd bond length (Å)
0.011
0.015
0.009
Rmsd bond angles (°)
1.3
1.8
1.3
*to be submitted to RCSB Protein Data Bank
Biochemistry. Author manuscript; available in PMC 2011 April 7.
|
3M3L
|
PEPA bound to the ligand binding domain of GluA2 (flop form)
|
The molecular mechanism of flop-selectivity and subsite
recognition for an AMPA receptor allosteric modulator:
Structures of GluA2 and GluA3 complexed with PEPA
Ahmed H. Ahmed§, Christopher P. Ptak§, and Robert E. Oswald*
Department of Molecular Medicine, Cornell University, Ithaca, NY 14853 USA
Abstract
Glutamate receptors are important potential drug targets for cognitive enhancement and the
treatment of schizophrenia in part because they are the most prevalent excitatory neurotransmitter
receptors in the vertebrate central nervous system. One approach to the application of therapeutic
agents to the AMPA subtype of glutamate receptors is the use of allosteric modulators, which
promote dimerization by binding to a dimer interface thereby reducing desensitization and
deactivation. AMPA receptors exist in two alternatively spliced variants (flip and flop) that differ
in desensitization and receptor activation profiles. Most of the structural information on
modulators of the AMPA receptor target the flip subtype. We report here the crystal structure of
the flop-selective allosteric modulator, PEPA, bound to the binding domains of the GluA2 and
GluA3 flop isoforms of AMPA receptors. Specific hydrogen bonding patterns can explain the
preference for the flop isoform. This includes a bidentate hydrogen bonding pattern between
PEPA and N754 of the flop isoforms of GluA2 and GluA3 (the corresponding position in the flip
isoform is S754). Comparison with other allosteric modulators provides a framework for the
development of new allosteric modulators with preferences for either the flip or flop isoforms. In
addition to interactions with N/S754, specific interactions of the sulfonamide with conserved
residues in the binding site are characteristics of a number of allosteric modulators. These, in
combination, with variable interactions with five subsites on the binding surface lead to different
stoichiometries, orientations within the binding pockets, and functional outcomes.
Membrane receptors are the cell's gatekeepers, allowing chemical signals access to the cell's
pathways. Through the binding of endogenous ligands, receptors identify relevant
environmental cues and facilitate cell-cell communication. The regulation of membrane
receptors has become an important goal of drug discovery efforts (1,2). By targeting the
physiological (orthosteric) ligand-binding site, agonists and antagonists control the function
of membrane receptors. Unfortunately, exogenously induced agonist-activation at the
orthosteric site can cause toxic effects from overstimulation. Allosteric modulator binding
sites use a distinct avenue for altering the natural response of a receptor. The ability of some
allosteric modulators to enhance receptor stimulation, while not actually providing the
trigger for stimulation, is a clear advantage that conserves the endogenous signaling
pathway. Being important mediators of higher-order processes such as learning and
memory, ionotropic glutamate receptors (iGluRs) have attracted a great deal of interest as
allosteric modulator targets (3–6). Of clear therapeutic importance, various
neurodegenerative disorders such as Parkinson's and Alzheimer's diseases, Huntington's
*Corresponding author; telephone: 1-607-253-3877; fax: 1-607-253-3659; email: reo1@cornell.edu.
§These authors contributed equally to this work.
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Published in final edited form as:
Biochemistry. 2010 April 6; 49(13): 2843–2850. doi:10.1021/bi1000678.
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chorea, and neurologic disorders including epilepsy and ischemic brain damage have been
linked to iGluRs (7).
The crystal structure of GluA2 (8) clarifies years of speculation on the complex arrangement
of the glutamate receptor's four subunits (9). The GluA2 can be dissected into 3 functionally
distinct layers. Farthest from the membrane, the amino terminal domain (ATD) can act as a
peripheral regulatory domain but is also involved in assembly and trafficking (10,11).
Sandwiched between the ATD and the membrane domain, the ligand-binding domain (LBD)
recognizes the neurotransmitter signal and directly regulates receptor activation (12).
Structures for both isolated extracellular domains (ATD and LBD) reveal a dimeric
organization (13–15). At the membrane interface, two alternative linker conformations
transition the 2-fold symmetry, which is adopted by both extracellular domains, into the 4-
fold symmetry of a membrane-traversing cation-selective channel (8,16). For iGluRs, the
ion channel domain confers functional relevance with its ability to selectively conduct the
flow of ions across the cell's membrane. The layers of extracellular domains, each with the
potential for multiple control points, allosterically regulate the ion channel domain's function
(8). Therefore it is not surprising that the ATD, the LBD, and the LBD-channel linker have
all been shown to be effective targets of allosteric modulators (13,17,18).
Since the structures of the ATD and the full iGluR channel have only recently been solved,
allosteric drug-binding sites external to the LBD have not been fully explored in molecular
detail. However, the decade-old LBD structure has proved to be indispensable as a heavily
exploited scaffold for understanding agonist, partial agonist, and antagonist binding
interactions as well as their ability to regulate channel gating behavior (12,19,20). Although
the dimeric organization is consistent across all iGluR subtypes, the molecular details of
LBD-agonist specificity define the subtype families into N-methyl-D-aspartic acid (NMDA)
receptors (21), α-amino-3-hydroxy-5-methyl-4-isoxazole-propionic acid (AMPA) receptors
(12), and kainate receptors (22). Because all subtypes are constrained by their conserved
sensitivity to glutamate stimulation, diversity at the orthosteric site is evolutionarily limited
and most agonists display cross-subtype activity. An allosteric modulator-binding site within
the quaternary LBD structure is located along the dimer interface (18) and offers improved
discrimination by modulators. Drugs that bind to the allosteric sites on the LBD dimer
interface can enhance the activity of iGluRs (23) and increase performance on tests of
memory (24). Except for the LBD structures with modulatory ions bound to the dimer
interface (25–27), only LBD structures from the AMPA receptor subtype, GluA2, have been
reported with bound allosteric modulators (18,28–31). Within the structures, the bound
modulatory drugs stabilize the LBD dimer interface, which is required for activation of the
ion channel and is dissociated during desensitization (18).
Although the residues that line the allosteric modulator-binding pocket do not differ between
AMPA receptors subtypes (GluA1–4), the ability of allosteric modulators to stabilize the
activated state still varies (32,33). Also, AMPA receptors can be alternatively spliced into
what is referred to as flip and flop isoforms (34). Modulator selectivity (23), desensitization
(35), and channel closing rates (36) differ between flip and flop. Although several of the
amino acid differences between the two forms are located in or near the allosteric
modulator-binding site, the difference at position 754 (serine in flip, asparagine in flop)
seems to be entirely responsible for the functional differences between allosteric modulator
regulation of the flip and flop variants (23,28,32). Cyclothiazide (CTZ) and some other
thiazide derivatives have improved binding to the flip form due to a hydrogen bond between
S754 and the NH of the fused thiazide ring (28). In the case of the flop form, the
alternatively spliced sequence places an asparagine in the 754 position, which is not
optimally positioned to form a hydrogen bond. Sekiguchi et al. (33) introduced an allosteric
modulator of AMPA receptors (4-[2-(phenylsulphonylamino)ethylthio]-2,6,-
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difluorophenoxyacetamide, PEPA) with a preference for the flop form. In fact, the relative
sensitivity of CTZ to PEPA has been used as a diagnostic for the prevalence of flip vs. flop
versions of AMPA receptor in particular cell types (37). PEPA shows potential in treatment
of post ischemic memory impairment (38) and contextual fear (39) but despite PEPA's
unique flop sensitivity, the modulator has not yet been used as a lead compound in SAR
studies.
For drug discovery to be guided by structures, understanding the possible molecular
interactions between modulators and the dimer interface is essential. We have shown
previously (31) that changes in the structures of CTZ derivatives can reorient the modulator
within the binding site. Subsequently, we proposed that the allosteric modulator site is
comprised of 5 subsites (Figure 1C). In the present study, we determine the three
dimensional structures of PEPA bound to the GluA2o and GluA3o LBDs (flop forms), and
use PEPA's binding interactions to further characterize the subsite specific binding
properties displayed by allosteric modulators. The amide group of PEPA makes a direct
hydrogen bond to N754, explaining the preferential action of PEPA on the flop form of
AMPA receptors. Another key structural element, the sulfonamide group of PEPA, is
conserved with the biarylsulfonamide class of allosteric modulators (6) and interacts with
the same residues of the dimer interface (8,30). Although previously classified as unrelated,
PEPA and the large group of biarylsulfonamide have similarities, which suggest that specific
PEPA groups (particularly the unique flop-interacting amide) can be strategically integrated
into biarylsulfonamide SAR studies.
Experimental Procedures
Materials
PEPA was purchased from Tocris (Ellisville, MO). The GluA2 S1S2J construct was
obtained from Eric Gouaux (Vollum Institute; 12).
Protein Preparation and Purification
GluA2 S1S2 consists of residues N392 - K506 and P632 - S775 of the full rat GluA2o
subunit (40), a `GA' segment at the N-terminus, and a `GT' linker connecting K506 and
P632 (12). A similar construct of GluA3 S1S2 was prepared as described previously (41).
pET-22b(+) plasmids were transformed in E. coli strain Origami B (DE3) cells and were
grown at 37°C to OD600 of 0.9 to 1.0 in LB medium supplemented with the antibiotics
(ampicillin and kanamycin). The cultures were cooled to 20°C for 20 min. and isopropyl-β-
D-thiogalactoside (IPTG) was added to a final concentration of 0.5 mM. Cultures were
allowed to grow at 20°C for 20 h. The cells were then pelleted and the S1S2 protein purified
using a Ni-NTA column, followed by a sizing column (Superose 12, XK 26/100), and
finally an HT-SP-ion exchange-Sepharose column (Amersham Pharmacia). Glutamate (1
mM) was maintained in all buffers throughout purification. After the last column, the protein
was concentrated and stored in 20 mM sodium acetate, 1 mM sodium azide, and 10 mM
glutamate at pH 5.5.
Crystallography
For crystallization trials, the protein was concentrated to 0.2 – 0.5 mM in 10 mM glutamate
using a Centricon 10 centrifugal filter (Millipore, Bedford, MA). For the PEPA-bound
structures, PEPA was added to 5 mM. The final protein concentration was 0.2 to 0.3 mM.
Crystals were grown at 4°C using the hanging drop technique, and the drops contained a 1:1
(v/v) ratio of protein solution to reservoir solution. The reservoir solution contained 14–15%
PEG 8K, 0.1 M sodium cacodylate, 0.1–0.15 M zinc acetate, and 0.25 M ammonium sulfate,
pH 6.5.
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Data were collected at the Cornell High Energy Synchrotron Source beam line A1 using a
Quantum-210 Area Detector Systems charge-coupled device detector. Data sets were
indexed and scaled with HKL-2000 (42). Structures were solved with molecular
replacement using Phenix (43). Refinement was performed with Phenix (43), and Coot 0.5
(44) was used for model building.
Results
Structure of PEPA bound to GluA2 S1S2 flop
The structure of glutamate bound to GluA2o S1S2 (3dp6; 41) was used as the initial search
probe for the molecular replacement solution of PEPA bound to GluA2o S1S2 with
glutamate in the agonist-binding site. PEPA was then modeled into two symmetrical
positions within the density found at the dimer interface, and the structure was optimized
using Phenix (43). The refinement statistics are given in Table 1. The resolution is 1.85 Å,
and three unique copies are found in the unit cell. The overall structure of the S1S2 domain
is very similar to the structure in the absence of PEPA, with contacts between glutamate and
the protein unchanged. However, PEPA clearly binds within the dimer interface, making
contacts with both monomers within the dimer. As shown in Figure 1, one PEPA molecule
binds per dimer interface. However because the dimer interface is symmetrical, two
equivalent orientations (related by a 180° rotation) are possible. Electron density for both is
seen in the crystal structure, although the intensity of one orientation is greater than the
other.
The binding of PEPA to the dimer interface increases the distance between the two
monomers that form the dimer by approximately 1.5 Å. This allows the relatively large
PEPA molecule to fit within the interface, but also increases the separation between the
linkers to the ion channel (the distance increases from 39.4 Å to 41 Å; Figure 1A). Relative
to the core of Lobe 1, both the J/K helices and one β strand (P105-G110) connecting the two
lobes are displaced slightly away from the dimer interface (Figure 1B). In addition, Lobe 2
is slightly twisted relative to glutamate-bound S1S2 in the absence of PEPA (3dp6; 41).
PEPA binds at the bottom of a water-filled, inverted U-shaped cleft with five subsites (A, B/
B′, and C/C′; 31). Upon binding, crystallographic waters are displaced from the central A
subsite and more buried C/C′ sites, with the waters in the B/B′ subsite remaining (Figure
1C). This displacement of presumably ordered water would be likely to contribute a
favorable entropy component to binding.
The sidechains of P494 are at the center of the interface and the edge of the two proline
rings from each monomer form the base of the binding site in which the difluorophenyl ring
resides (Figure 2A). This is close to the position of the methoxybenzoyl ring of aniracetam
in its structure bound to GluA2-S1S2(FW) (29). The other side of the ring is exposed to
S497 and S729. The sidechain hydroxyl of S497 is oriented toward the dimer interface in the
absence of PEPA, but rotates out toward the solvent to accommodate the difluorophenyl ring
of PEPA (Figure 1B). The amide of PEPA is involved in a network of hydrogen bonds with
sidechain hydroxyl of Y424, the backbone carbonyl of F495, the sidechain carboxyl of
D760, the sidechain amide of N754, and two water molecules (Figure 2A). The most
striking of these hydrogen bond pairs is with N754. This represents the only difference
between the flip (S754) and flop (N754) isoforms in the PEPA binding site and is almost
certainly a major source of the preference for the flop isoform. The phenyl-sulfonylamide
side of PEPA inserts into a hydrophobic pocket formed by sidechain methyls of I481 and
L751 as well as methylene groups contributed by K493, N754, and E755 (Figure 2B). It is
possible that the contribution by methylene group of N754 provides a more hydrophobic
pocket than S754 in the flip form, further contributing to the preference for the flop form.
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Because the dimer interface is symmetrical, PEPA can bind in two orientations and both are
observed in the crystal. For this reason, changes in the protein due to a specific interaction
with PEPA can be partially masked because each monomer is a weighted average of two
orientations of bound PEPA. However, one orientation has a stronger density than the other,
providing some insight into the extent of changes in the dimer interface that are produced by
PEPA binding. As shown in Figure 2C, the two monomers comprising the dimer differ more
within the PEPA binding site than the corresponding monomers in the absence of PEPA.
One turn of helix J (L751 to N754) contains important determinants for both orientations of
PEPA. In one orientation the amide group of PEPA interacts with the sidechain of N754,
and in the other, the aromatic ring of PEPA inserts between a hydrophobic pocket formed by
the sidechain of L751 and the methylene group of N754. In the orientation for which the
density of the amide of PEPA is stronger, N754 is better positioned to form an H-bond
(Figure 2D); whereas, in the other side of the interface, N754 is oriented to form an H-bond
with the carbonyl of S729. This change in orientation facilitates the insertion of the aromatic
ring of PEPA into the hydrophobic pocket, which is accompanied by a small shift in the
sidechain of L751 to accommodate the aromatic ring (Figure 2D). Since these structures are
weighted averages, it is possible that the actual positions of these sidechains involve an even
greater movement than is seen from the asymmetry of the crystal.
Structure of PEPA bound to GluA3 S1S2 flop
In studies of the physiological effects of PEPA, a significant difference between subtypes
has been observed, with GluA3 being most susceptible to modulation (33). The structure of
GluA3i S1S2 bound (flip form) to glutamate has been reported previously (41). Since PEPA
preferentially binds to the flop form, the GluA3o structure was determined bound to
glutamate with and without PEPA (Figure 3A). Like GluA2o, in the absence of PEPA,
GluA3o has three copies in the asymmetric unit. Comparing lobe closure between GluA3i
and GluA3o, the flop form is slightly more closed (1.6° ± 0.7°).
In the presence of PEPA, GluA3o was present in one copy in the asymmetric unit, and PEPA
was observed with the same density in two symmetrical orientations. Like GluA2o bound to
PEPA, the dimer interface (assessed using the symmetrical molecule in the crystal) was
displaced relative to the unbound from (Figure 3A) by approximately 2.5 Å at the position
of the linker replacing the ion channel domain. Within the binding site, three sidechains
exhibited different rotamers compared with the GluA2o structure bound to PEPA (Figure
3B). For PEPA-bound GluA3o, both S497 and S729 assumed rotameric states that differed
both from GluA2o bound to PEPA and from GluA2o and GluA3o in the absence of PEPA. In
the case of S729, the rotameric state in combination with a slight movement of the amide of
PEPA (relative to the GluA2o structure) would make an H-bond with the sidechain of S729
(shown in Figure 2A for GluA2o) unlikely. In the case of N754, the sidechain is displaced
relative to the GluA2o-PEPA structure so that only one H-bond is made to the amide of
PEPA. This may be a result of averaging of the two orientations of PEPA only one of which
forms a bidentate H-bond with N754.
Discussion
The goal of allosteric modulation, like orthosteric modulation, is often to stabilize a
conformational state of a dynamic protein (45). The activated state of iGluRs is naturally
unstable allowing the channel to desensitize (46). Disruption of the symmetrical dimer
interface between LBDs is thought to initiate desensitization-mediated channel closure (47).
By maintaining the LBD dimer, positive allosteric modulators can prevent desensitization
and prolong activation (18). Currently, 15 crystal structures of the GluA2 LBD with bound
allosteric modulators are deposited in the Protein Data Bank (48). All of these modulators
bind to a large crevice with 2-fold symmetry along the symmetric dimer interface (18). The
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large variation in structure among allosteric modulators results in significant variations in
binding orientations and interactions. At least four distinct binding modes have been
identified: (1) A-subsite class (aniracetam, CX614 (29)), (2) classical thiazide (cyclothiazide
(18), TCMZ, ALTZ (31)), (3) the shifted thiazide class (IDRA-21, HCMZ, HFMZ; (31)),
and (4) the full spanning class (PEPA (this paper), dimeric biarylpropylsulfonamide (30),
LY404187 (8)). Overlaying modulators from these structural classes has led to the proposal
that the allosteric modulator site is comprised of a series of subsites (Figure 1C;31).
Positioned at the center of the binding-site, the symmetric A subsite is narrow and allows
entrance to only one molecule. Two subsites (B and C) lie at each end of the A subsite with
the hydrophobic C subsite located more deeply in the pocket effectively defining five
subsites (A, B, B′, C, and C′).
In the open state, the subsites are filled with water, which may act to weakly stabilize the
dimer. Allosteric modulators generate stronger interactions across the subsites thereby
increasing the linkages between the monomers. The simplest modulator class, including
aniracetam and CX614, fills the A subsite with one molecule but does not enter the
peripheral B and C subsites (29). The two classes of thiazide-based modulators account for
10 of the 15 solved allosteric modulator-GluA2 crystal structure complexes (18,28,31). The
classical thiazide (CTZ-like) binding class and the shifted thiazide (IDRA-21-like) binding
class are positioned respectively in the B and C subsite or mainly the C subsite. Most of the
thiazide modulators do not extend across the A subsite and therefore can bind two molecules
per dimer. However, a few of the newly described shifted thiazides (HFMZ, HCTZ; 31)
enter the A subsite but only enough to impair binding of a second modulator. The dimeric
biarylpropylsulfonamide compound ((R,R)-N,N-(2,2'-[Biphenyl-4-4'-Diyl]Bis[Propane-2,1-
Diyl]) Dimethanesulfonamide) described by Kaae et al. (30) was the first allosteric
modulator shown by crystallography to extend along the entire length of the inner dimer
cavity from C to C′ subsites. PEPA also interacts with J helices from both monomers, which
cap the ends of the modulator-binding pocket. The density occupied by both symmetrical
copies of PEPA overlays the dimeric biarylsulfonamide compound as both modulators
represent the full spanning class (Figure 4B).
The GluA flip and flop splice variants differ by only a few residues along the J helix in the
LBD; however, residue 754 (Asn in flop and Ser in flip) is positioned between the B and C
subsites. For thiazides, a clear preference in binding to the flip-form is mediated by a
hydrogen bond between the hydrobenzothiadiazide ring and S754 (28). In contrast, PEPA is
flop-selective and the PEPA-bound structure provides the first structure containing a direct
interaction between a modulator and the flop form's N754. The amide of PEPA extends
straight out from the A subsite and across the B and C subsite interface to make an amide-
amide hydrogen bond with N754 (Figure 2A). Unlike most other AMPA modulators, PEPA
fills neither the B nor the C subsites but interacts directly with the J helix. A similar
interaction is seen with LY404187 (49) bound to GluA2i (8). Strong hydrogen bonding can
occur between two amides (50) and has been shown to be responsible for driving
oligomerization of transmembrane leucine zippers (51). The distances between the
interacting amides in the PEPA-bound structure support a bidentate hydrogen-bonding
pattern, which is much stronger and more specific than a typical hydrogen bond. While
PEPA is selective for the AMPA receptor's flop form, a weaker but still existent potentiation
of the flip form has been observed (33,52). Replacing N754 (flop) with S754 (flip) would
not prevent PEPA from binding; however, serine would provide only one hydrogen-bonding
partner for PEPA's amide with an extended interaction distance. In contrast, LY404187
displays a preference for the flip isoform (53), and its cyano group extends out to interact
directly with S754. The cyano-S754 interaction is a clear flip analog of the flop-selective
PEPA amide-N754 interaction (Figure 4A).
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Opposite to the amide on the PEPA molecule, a sulfonamide is tethered to the
difluorophenyl ring (Figure 4A). Within the dimer interface, the sulfonamide is positioned
so the nitrogen can hydrogen bond directly with the carbonyl of P494 (Figure 4C). A
sulfonamide oxygen points toward the amide nitrogen of G731. The angle of the peptide
plane is perpendicular to the sulfonamide oxygen, making a hydrogen bonding interaction
unlikely (Figure 4D). Instead, a dipole-dipole or charge-dipole interaction may occur. The
amide nitrogen of a polypeptide supports at least a partial positive charge (54), which would
interact with the strongly electronegative sulfonamide oxygen (55). Interestingly, both the
dimeric biarylsulfonamide (30) and LY404187 (8), other members of the full spanning
modulator class, also have a sulfonamide that interacts with the same backbone atoms of
P494 and G731 as PEPA (Figure 4C).
A large number of biarylsulfonamides have been identified that modulate AMPA receptors
and are being evaluated for therapeutic use in the treatment of depression and Parkinson's
disease (56). The conserved sulfonamide reveals a previously unidentified relationship
between PEPA and the biarylsulfonamide modulators. When the perpendicular peptide bond
plane including G731 is fixed, the sulfonamide on three overlayed modulators varies by 1.2
Å along the length of the interface with the PEPA sulfonamide being positioned closer to the
A subsite (Figure 4D). A shift of the sulfonamide also results in a shift in the corresponding
P494 across the interface presumably to maintain the hydrogen bond with the modulator's
amine. The sulfonamide forms an important bridge between the two dimer halves. For
PEPA, a phenyl-sulfonamide replaces the methyl-sulfonamide in the dimeric
biarylsulfonamide and fits snuggly against L751. Based on the orientation-induced
asymmetry within the GluA2-complex structure, the phenyl pushes the J helix away from
PEPA thereby affecting the C subsite (Figure 2C and D). Residues lining the C subsite are
on the same beta strand as G731, which must shift if the C subsite is to remain together and
presumably explain the 1.2 Å shift relative to the dimeric biarylsulfonamide. In fact, the
same phenyl-sulfonamide group substitution in a biarylpropylsulfonamide decreases the
modulatory effect of the derivative in SAR studies (57). For biarylpropylsulfonamides, the
optimal sulfonamide substitution was found to be either an ethyl or an iso-propyl group,
which should both fit without significantly disrupting the J helix or C subsite (57).
The PEPA-bound crystal structure from AMPA receptor subtypes, GluA2 and GluA3, do
not display major differences in binding interactions even though PEPA exhibits a stronger
effect on GluA3 (33). For GluA2, an asymmetry in the receptor-binding pocket was
observed while no significant difference in PEPA density was seen for the each orientation
within the GluA3 crystal structure. In addition, a number of side chains exhibit different
rotameric states between the two structures, although it is unlikely that these small changes
significantly impact the differential effects on the two subtypes. Although no structural
differences have been identified between GluA2 and GluA3 that would obviously impact
PEPA affinity, the possibility exists that subtle differences arising from the sequence
differences peripheral to the binding site may be important as has been described in the case
of the agonist binding site of GluA4 (58).
We have explored how PEPA (this paper) and other allosteric modulators (31) interact with
the GluA interface in the context of drug design. Together the identification of a conserved
group between PEPA (this paper) and biarylpropylsulfonamides (8,30) and the regional
nature of various subsite-functional group interactions provide a backdrop to extend
biarylpropylsulfonamide SAR studies (57) to include PEPA and biarylpropylsulfonamide
chimeras. Although optimizing the stability of the dimer interface provides a starting point
for SAR studies, additional constraints should be considered including the ability of the
modulator to enter the cavity, the dynamic structure of the dimer interface during closed,
open, and desensitized state transitions, and the ability of the modulator to cross the blood-
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brain barrier before being metabolized. This definition of the allosteric modulator binding-
site should provide guidance in glutamate receptor allosteric modulator pharmacology.
Acknowledgments
We thank Prof. Eric Gouaux (Vollum Institute) for the GluA2 S1S2J construct, and Prof. Linda Nowak (Cornell)
for the full-length GluA3 construct.
This work was supported by a grants from the National Institutes of Health (R01-GM068935, R01 NS049223, and
R21 NS067562). This work is based upon research conducted at the Cornell High Energy Synchrotron Source
(CHESS), which is supported by the National Science Foundation under award DMR 0225180, using the
Macromolecular Diffraction at the CHESS (MacCHESS) facility, which is supported by award RR-01646 from the
National Institutes of Health, through its National Center for Research Resources.
Abbreviations
ALTZ
althiazide
AMPA
α-amino-3-hydroxy-5-methyl-4-isoxazole-propionic acid
CLTZ
chlorothiazide
CX614
pyrrolidino-1,3-oxazino benzo-1,4-dioxan-10-one
CTZ
cyclothiazide
FW
(S)-5-fluorowillardiine
flip and flop
alternatively spliced versions of AMPA receptors that vary in rates of
desensitization and sensitivity to allosteric modulators
iGluR
ionotropic glutamate receptor
GluA1-4
four subtypes of AMPA receptor
HCTZ
hydrochlorothiazide
HFMZ
hydroflumethiazide
IDRA-21
7-chloro-3-methyl-3,4-dihydro-2H-benzo[e][1,2,4]thiadiazine 1,1-dioxide
IPTG
isopropyl-β-D-thiogalactoside
LY404187
N-[2-(4′-cyanobiphenyl-4-yl)propyl]propane-2-sulfamide
PEPA
4-[2-(phenylsulphonylamino)ethylthio]-2,6,-difluorophenoxy acetamide
NMDA
N-methyl-D-aspartic acid
S1S2
extracellular ligand-binding domain of GluA2 and GluA3
SAR
structure-activity relationships
TCMZ
trichlormethiazide
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Figure 1.
(A) Comparison of glutamate-bound GluA2o S1S2 in the presence (blue) and the absence of
PEPA (green) in two orientations. Both orientations of PEPA are shown. Note that the
binding of PEPA results in a separation of the two components of the dimer (distance
between the Cα atoms of the threonine in the linker) by approximately 1.5 Å. (B) One
monomer of GluA2o S1S2 in the presence (blue) and the absence of PEPA (green) with one
orientation of PEPA shown. Both the J/K helices and the strand near S497 are displaced
upon binding PEPA. Also, the sidechains of S497 and S729 change rotameric states. (C)
Comparison of the water molecules at the dimer interface in the presence (tan spheres) and
the absence of PEPA (red spheres). PEPA is shown in both orientations. Despite the greater
separation of the dimer interface, a number of the ordered water molecules found in the
absence of PEPA are displaced by PEPA. The black circles delineate subsites of the
allosteric modulator binding site as described previously (31).
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Figure 2.
The PEPA binding site, emphasizing the important interactions, shown in two orientations.
(A) A view of the amide side of PEPA bound to GluA2 S1S2. The hydrogen bonding
network with the amide of PEPA is shown as dotted lines. The H-bond with the sidechain of
S729 is difficult to display in the orientation used in the figure. (B) A view of the phenyl
group of PEPA inserted into a hydrophobic pocket in GluA2 S1S2. (C) RMS plot showing
more variability in the J/K helices for the PEPA-bound structure than the unbound structure.
(D) J/K helix showing where differences in the two orientations were analyzed. The amide
of PEPA-N754 interaction (blue) maintains the position of the J helix in the absence of
PEPA (green) The J helix is displaced on the phenyl side of PEPA (red).
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Figure 3.
(A) Comparison of glutamate-bound GluA3o S1S2 in the presence (blue) and the absence of
PEPA (green) in two orientations. Both orientations of PEPA are shown. Note that the
binding of PEPA results in a separation of the two components of the dimer (distance
between the Cα atoms of the threonine in the linker) by approximately 2.5 Å. (B) One
monomer of GluA3o S1S2 in the presence (blue) and the absence of PEPA (green) with one
orientation of PEPA shown. Shown for comparison is the PEPA-bound form of GluA2o
(red). Both the J/K helices and the strand near S497 are displaced upon binding PEPA for
both GluA2o and GluA3o. Also, the sidechains of S497 and S729 are in different rotameric
states for GluA3o bound to PEPA compared with GluA3o in the absence of PEPA and
GluA2o bound to PEPA. Also, N754 is displaced in PEPA-bound GluA3o, such that only
one H-bond is possible with the amide of PEPA.
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Figure 4.
(A) Members of the full spanning class of allosteric modulators. The shape-highlighted
regions of the modulators illuminate key contact points to the specific binding pocket
residues and subsites (as labeled for PEPA). (B) Overlay of the full spanning modulator
structures. The structures were aligned at both sets of P494 and G731 residues. PEPA (gray)
occupies a similar arrangement of subsites as the dimeric biarylsulfonamide (PDB entry
3bbr, cyan, 30) and LY404187 (PDB entry 3kgc, magenta, 8). (C) The sulfonamide bridges
the two monomers in both PEPA and the dimeric biarylsulfonamide with the same
interactions to P494 and G731. (D) The hydrogen bond between the carbonyl of P494 and
the sulfonamide is maintained when the modulator is in a shifted position relative to the
peptide plane of K730 and G731 (green disk) located on the opposite monomer.
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Table 1
Structural Statistics
GluA2o (PEPA)
GluA3o (PEPA)
GluA3o
Space Group
P22121
P222
P222
Unit Cell (Å)
a=47.13 b=113.92 c=164.81
a=46.95 b=52.26 c=115.98
a=46.03 b=110.33 c=161.192
X-ray source
CHESS (A1)
CHESS (A1)
CHESS (A1)
Wavelength (Å)
0.977
0.977
0.977
Resolution (Å)
50–2.0 (2.03–2.00)
50–2.5 (2.54–2.00)
50–1.85 (1.88–1.85)
Measured reflections (#)
817961
62549
344340
Unique reflections (#)
70175
9141
69379
Data redundancy
6.9 (7.1)
6.0 (4.2)
4.7 (3.0)
Completeness (%)
99.9 (100.0)
99.5 (89.8)
96.5 (73.5)
Rsym (%)
11.4 (34.2)
13.0 (45.5)
7.5 (24.6)
I/σi
33.3 (7.1)
19.2 (2.5)
34.3 (3.3)
PDB ID
*
*
*
Current Model Refinement Statistics
Phasing
MR
MR
MR
Molecules/AU
3 (no NCS applied)
1
3 (no NCS applied)
Rwork/Rfree (%)
18.7/24.2
18.9/28.5
20.1/23.4
Free R test set size (#/%)
2000 (2.85)
914 (10.0)
2000 (2.88)
Number of protein atoms
5979
2030
6091
Number of heteroatoms
111
62
30
Rmsd bond length (Å)
0.011
0.015
0.009
Rmsd bond angles (°)
1.3
1.8
1.3
*to be submitted to RCSB Protein Data Bank
Biochemistry. Author manuscript; available in PMC 2011 April 7.
|
3M3N
|
Structure of a Longitudinal Actin Dimer Assembled by Tandem W Domains
|
Structure of a Longitudinal Actin Dimer Assembled by Tandem
W Domains – Implications for Actin Filament Nucleation
Grzegorz Rebowski1,§, Suk Namgoong1,§, Malgorzata Boczkowska1, Paul C. Leavis2, Jorge
Navaza3, and Roberto Dominguez1,*
1Department of Physiology, 3700 Hamilton Walk, University of Pennsylvania School of Medicine,
Philadelphia, PA 19104-6085, USA.
2Boston Biomedical Research Institute, Watertown, MA 02472-2899, USA.
3Institut de Biologie Structurale, F-38027 Grenoble, France
Abstract
Actin filament nucleators initiate polymerization in cells in a regulated manner. A common
architecture among these molecules consists of tandem W domains that recruit three to four actin
subunits to form a polymerization nucleus. We describe a low-resolution crystal structure of an
actin dimer assembled by tandem W domains, where the first W domain is crosslinked to Cys-374
of the actin subunit bound to it, whereas the last W domain is followed by the C-terminal pointed
end-capping helix of Tβ4. While the arrangement of actin subunits in the dimer resembles that of a
long-pitch helix of the actin filament, important differences are observed. These differences result
from steric hindrance of the W domain with inter-subunit contacts in the actin filament. We also
determined the structure of the first W domain of Vibrio parahaemolyticus VopL crosslinked to
actin Cys-374, and show it to be nearly identical to non-crosslinked W-actin structures. This result
validates the use of crosslinking as a tool for the study of actin nucleation complexes, whose
natural tendency to polymerize interferes with most structural methods. Combined with a
biochemical analysis of nucleation, the structures may explain why nucleators based on tandem W
domains with short inter-W linkers have relatively weak activity, cannot stay bound to filaments
after nucleation, and are unlikely to influence filament elongation. The findings may also explain
why Nucleation Promoting Factors of the Arp2/3 complex, which are related to tandem W domain
nucleators, are ejected from branch junctions after nucleation. We finally show that the simple
addition of the C-terminal pointed end-capping helix of Tβ4 to tandem W domains can change
their activity from actin filament nucleation to monomer sequestration.
Keywords
Actin nucleation; W domain; Nucleation Promoting Factors; Tβ4; actin monomer sequestration
© 2010 Elsevier Ltd. All rights reserved.
*Corresponding author (Phone: 215-573-4559; droberto@mail.med.upenn.edu).
§These authors contributed equally to this work
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PDB ACCESSION NUMBERS
Coordinates and structure factors were deposited under PDB ID 3M1F (WxActin) and 3M3N (3W-Actin).
SUPPLEMENTARY MATERIAL
Supplementary material is available online, including supplementary Material and Methods and Movies S1–S4.
NIH Public Access
Author Manuscript
J Mol Biol. Author manuscript; available in PMC 2011 October 15.
Published in final edited form as:
J Mol Biol. 2010 October 15; 403(1): 11–23. doi:10.1016/j.jmb.2010.08.040.
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INTRODUCTION
The nucleation of actin filaments in cells is kinetically unfavorable because of the instability
of polymerization intermediates (dimers, trimers and tetramers) and the actions of actin
monomer binding proteins such as profilin and thymosin-β4 (Tβ4) 1; 2. This creates an
opportunity for cells to use molecules known as actin filament nucleators to initiate the
formation of actin polymerization nuclei in a spatially and temporally controlled manner.
The actin filament can be described as either a single left-handed short-pitch helix, where
consecutive subunits are staggered with respect to one another by half a monomer length, or
two right-handed long-pitch helices of head-to-tail bound actin subunits 3; 4; 5. Different
nucleators work by different mechanisms, stabilizing small actin oligomers along either the
long- or the short-pitch helices of the actin filament 6; 7.
Most actin filament nucleators use the WASP-Homology 2 (WH2 or W) domain for
interaction with actin. The W domain has a short length (17–27aa) and is extremely
abundant and functionally versatile 7; 8; 9. The N-terminal portion of the W domain forms a
helix that binds in the hydrophobic (or target-binding) cleft 10 formed between subdomains
1 and 3 at the barbed end of the actin monomer 11; 12; 13. After this helix, the W domain
presents an extended region that is directed towards the pointed end of the actin monomer
(formed by subdomains 2 and 4 of actin). This region is variable in length and sequence, but
comprises the conserved four residue motif LKKT(V), which is critical for the interaction
with actin 11.
Filament nucleators are characterized by the presence of multiple actin-binding sites. The
simplest and most common architecture consists of tandem repeats of the W domain,
occurring in the proteins Spire 14, Cobl 15 and VopL/VopF 16; 17. The W domain also
participates in filament nucleation through the Nucleation Promoting Factors (NPFs) of the
Arp2/3 complex, which can have between one and three W domains 18; 19; 20. The muscle-
specific nucleator Lmod also contains one W domain 21. The nucleation activities of tandem
W domain-based nucleators vary widely. At least in part, the reason for these differences
may lie in the highly variable linkers between W domains. When the linkers are short, as in
the relatively weak nucleator Spire 14, only actin subunits along the long-pitch helix of the
actin filament can be connected. In contrast, the brain-enriched protein Cobl is a strong
nucleator, featuring three W domains with a long linker between its second and third W
domains, and is thought to stabilize a short-pitch actin trimer for nucleation 15. The
examples of Cobl, the Arp2/3 complex, and formins suggest that stabilization of a short-
pitch actin nucleus is a more effective way to promote polymerization than stabilization of a
long-pitch actin nucleus 6; 7. However, the structural bases for this observation are not well
understood.
In an attempt to understand the nucleation mechanism of tandem W domain-based
nucleators, we recently reported a solution study, using Small Angle X-ray Scattering
(SAXS), of an actin dimer and a trimer stabilized by tandem W domain constructs 22. These
complexes, referred to as 2W-Actin and 3W-Actin, and containing respectively two and
three W domains, were capped at the barbed end by structure-based crosslinking of the first
W domain to Cys-374 of the first actin subunit and at the pointed end by addition of the C-
terminal helix of Tβ4. Constructs 2W and 3W are based on the W domain repeat present in
the NPF protein N-WASP, which like Spire presents short inter-W linkers. The SAXS study
suggested that the actin subunits in the complexes adopted an elongated conformation
similar to that of the long-pitch helix of the actin filament. However, the resolution of this
study was insufficient to establish a direct comparison between the longitudinal contacts of
actin subunits in the complexes and in the actin filament.
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Here we report the crystal structure of 3W-Actin at 7 Å resolution. Only two actin subunits
are present in the structure, indicating that one of the actin subunits is released during
crystallization. Despite its low resolution, this structure, obtained by fitting high-resolution
structures of W-Actin complexes into the low-resolution data, offers a clearer picture of the
relative disposition of actin subunits bound to tandem W domains that are separated by
Spire-like short inter-W linkers. While the longitudinal arrangement of actin subunits in the
structure is somewhat related to that of the long-pitch helix of the actin filament 3; 4,
important differences are observed. These differences probably result from steric hindrance
of the W domain with inter-subunit contacts in the filament. The determination of the
structure of 3W-Actin was aided by determination of the 2.9 Å resolution crystal structure of
the first W domains of VopL 16 crosslinked to actin Cys-374 (hereafter referred to as
WxActin). The structure of WxActin is nearly undistinguishable from non-crosslinked W-
Actin structures determined previously 11; 12; 13, thus validating the use of crosslinking as
a tool to stabilize actin polymerization complexes for structural investigation. The structures,
and a biochemical analysis of nucleation, reveal important clues about the existing
disparities in the nucleation activities of tandem W domain-based nucleators.
RESULTS AND DISCUSSION
Crystal structure of crosslinked WxActin
In two previous studies, we reported low-resolution SAXS structures of actin nucleation
complexes formed by the Arp2/3 complex and tandem W domains 22; 23. Barbed end
polymerization in these studies was blocked by crosslinking of the W domain to Cys-374 of
the actin subunit located at the barbed end of the complexes. This approach was based on
analysis of the structures of various W-actin complexes 11; 12; 13, which placed the N-
terminus of the W domain within disulfide bond distance to actin Cys-374. In each case, a
Cys residue was introduced into the W domain at the most favorable position for
crosslinking to actin Cys-374. Here, this approach was used again to obtain the low-
resolution crystal structure of 3W-Actin. However, it remained unclear whether the crosslink
altered the structure of actin and/or the W domain in a significant way, which prompted us
to pursue the determination of a crosslinked WxActin structure. It later became apparent that
this structure also provided the best molecular replacement model for determination of the
structure of 3W-Actin.
After testing crystallization with various W domains, good diffracting crystals were obtained
of the crosslinked complex of actin with a synthetic peptide corresponding to the first W
domain (amino acids 130–160) of Vibrio parahemolyticus VopL. During synthesis, residue
Val-131 of this W domain was replaced by Cys and crosslinked to actin Cys-374 (Materials
and Methods). The crystal structure of WxActin was determined by molecular replacement
to 2.9 Å resolution (Fig. 1A and Table 1).
The structure of WxActin is very similar to those of non-crosslinked W-Actin complexes
determined with bound DNase I 11; 13 and that of Drosophila ciboulot bound to actin-
latrunculin A 12. Figure 1B shows a comparison of the structure of WxActin with that of the
non-crosslinked complex of actin with the W domain WASP (PDB code 2A3Z). The two
structures superimpose with r.m.s deviation of 0.66 Å for 358 equivalent Cα atoms. The
most important differences occur in regions that were visualized in one of the structures but
not the other, including the DNase I-binding loop (D-loop), the C-terminus of actin, and the
N-terminus of the W domain. The D-loop is disordered in most structures of actin, as well as
in the structure of WxActin described here, but forms an extended β-sheet with β-strands of
DNase I in the non-crosslinked structure. The C-terminus of actin is also disordered in most
crystal structures, except complexes with profilin, which interacts with the C-terminus of
actin 24; 25; 26. In the non-crosslinked W-Actin complex, the last 10 amino acids of actin
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(Gly-366 to Phe-375) are disordered and the W domain is only visualized starting from
WASP residue Arg-431 (corresponding to VopL Asn-132). In contrast, in the crosslinked
structure only the last amino acid of actin (Phe-375) is unresolved in the electron density
map, whereas the W domain of VopL is visualized from residue 130 to 151, i.e. the last nine
amino acids of the synthetic peptide were not resolved. The disulfide bond between actin
Cys-374 and VopL Cys-131 is visualized in the electron density map (inset in Fig. 1A),
although it is poorly defined compared to the rest of the structure.
The similarity of the structures suggests that the crosslinking approach used here and in
previous studies 22; 23 as a tool to cap the barbed end of actin polymerization nuclei for
structural investigation does not introduce significant structural distortions. Furthermore, as
we show next, the availability of the structure of WxActin aided the determination of the
structure of 3W-Actin.
Crystal structure of 3W-Actin
The solution SAXS study of 3W-Actin revealed an elongated molecule, consistent with the
presence of three actin subunits, somewhat similar to the long-pitch helix of the actin
filament 22. However, the nature of actin-actin contacts in the complex could not be
determined. We had suggested that subdomain 2 of actin could move slightly, which
combined with a helical conformation in the D-loop, would make the binding of tandem W
domains fully compatible with intersubunit contacts in the actin filament 3; 4. Other
investigators had suggested that the W domain would probably interfere with intersubunit
contacts in the filament 27; 28. Knowing which proposal is correct is important, because it
may shed light on the mechanism of nucleation, and possibly explain why tandem W
domain-based nucleators do not influence elongation the way formins do. It may also
answer important questions about differences in the activities of tandem W domain-based
nucleators, and the mechanism of action of NPFs of the Arp2/3 complex, which also contain
tandem W domains 7. Therefore, we set out to crystallize the complexes of 2W-Actin and
3W-Actin. While both complexes were crystallized readily, the crystals did not diffract the
X-rays. Additional search for conditions led to the identification of additives, such as RbCl
and polyvinylpyrrolidone K15, which improved diffraction somewhat. After several
attempts, the best result consisted of a rather complete and highly redundant X-ray dataset
collected from crystals of 3W-Actin to 7 Å resolution. While we initially considered not
reporting this structure, we later recognized that significant information could be obtained
by positioning high-resolution W-Actin structures into the unit cell of the 3W-Actin crystals
by molecular replacement. Because the individual structures are known at high-resolution,
this approach overcomes some of the typical limitations of low-resolution structures in
which the content of the unit cell is totally unknown. The limitations, however, are that
individual atomic positions cannot be refined and the inter-W linkers cannot be visualized.
Consistent with the design and mass measurements in solution of the complex of 3W-Actin
22, three copies of the W-Actin basic unit were expected in the asymmetric unit of the
crystal. The volume of the asymmetric unit was also compatible with it containing three
copies of the W-Actin unit (corresponding to a solvent content of 43%). However, weak
diffraction is typically consistent with a higher solvent content. Not surprisingly, the
molecular replacement solution, performed independently with the programs Phenix 29; 30
and AMoRe 31, located only two W-Actin complexes in the asymmetric unit, for a solvent
content of 62% (see Material and Methods and detailed description in Supplementary
Material). We do not understand why one of the actin molecules dissociates during
crystallization, although it could simply be that this molecule is bound loosely and is
therefore displaced by favorable crystal contacts. Analysis of the crystal packing
demonstrates why a third actin molecule was never found. Consecutive actin dimers are
stacked head-to-tail, forming a helix along the crystallographic c axis (Movie S1). Two such
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helices assemble tightly in anti-parallel fashion (see Movie S2). Each anti-parallel pair
comprises 24 actin subunits along the length of the c axis, which constitutes the basic
building block of the crystal lattice. Adjacent pairs of helices crossover twice in a repeat (or
helical turn), corresponding to the length of the c axis (see Movies S3 and S4), thus assuring
the connectivity of the crystal lattice and leaving no extra-space for the missing third actin
subunit (or rather 12 actin subunits, when the P6522 symmetry of the crystal is taken into
consideration).
Because of the limited resolution, we could not identify which of the actin subunits is lost
during crystallization, or whether the crystals consist solely of the actin subunit crosslinked
to the long 3W polypeptide. Note that any non-crosslinked actin dissociated from the
complex would be expected to polymerize during crystallization, and would therefore not be
present in the crystals. To address this question, a large number of crystals were collected,
washed multiple times in the crystallization solution by transferring them with a cryo-loop,
and then dissolved in water for analysis by non-reducing gel electrophoresis and mass
spectrometry (Materials and Methods). The results clearly illustrate that the crystals consist
of a 50/50 mixture of actin crosslinked to construct 3W and non-crosslinked actin (Fig. 2A).
Therefore, we conclude that one of the non-crosslinked actins was lost during crystallization
which, based on the arrangement of actin subunits in the asymmetric unit, is most likely that
bound to the last W domain.
The disposition of the actin subunits in the structure of 3W-Actin (Fig. 2B) is somewhat
similar to the longitudinal arrangement of actin subunits in the long-pitch helix of the actin
filament model 3; 4 (Fig. 2C). However, important differences are observed. To better
understand these differences, it is important to discuss what is currently known about
longitudinal contacts in the actin filament. Multiple crystal structures of actin show similar
longitudinal contacts between actin subunits (including both non-crystallographic dimers
and symmetry-related dimers), which are thought to mimic inter-subunit contacts in the actin
filament (Table 2). However, because of constraints imposed by crystal symmetry, these
dimers are unwound, i.e. they lack the natural twist of the actin filament. A detailed analysis
of these structures and their implications for our understanding of the actin filament has
been carried out by other investigators 32, and will not be repeated here. However, it is
important to compare the structure of 3W-Actin to both the actin filament model 3; 4 and the
longitudinal dimers observed in crystal structures, with the understanding that the structure
of 3W-Actin does not address the conformation of the actin filament per se, but rather the
mechanism of recruitment of actin subunits by tandem W domain proteins.
The dimers observed in crystal structures are generally similar and often crystallographically
isomorphous. Based on a superimposition of their structures, we have identified three
subgroups that diverge more significantly (represented by PDB entries 2FXU, 1Y64 and
2HMP) (Table 2). Compared to a long-pitch dimer of the actin filament model in which
consecutive subunits are rotated by ~27° 3; 4, these three subgroups present flat structures,
i.e. rotated counterclockwise with respect to the filament dimer by approximately −27° (Fig.
2D) (although the orientation of the axis of rotation is markedly different for entry 2HMP).
Remarkably, the longitudinal contacts between subdomains 4 and 3 of neighboring actin
subunits are well conserved in the three subgroups (Fig. 3). It thus appears that longitudinal
contacts between actin subunits in the filament have a strong tendency to reemerge as crystal
contacts in actin structures 32. It is important to note that these structures offer the most
accurate view of longitudinal contacts currently available 32, because the resolution of the
actin filament model 3; 4 is still insufficient to address specific atomic interactions.
Additional longitudinal contacts are thought to involve the D-loop in subdomain 2 33, which
is proposed to bind in the hydrophobic cleft between subdomains 1 and 3 of the actin
subunit immediately above it 4. However, the D-loop is disordered in all the crystal
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structures containing longitudinal actin dimers, and its conformation(s) and actual contacts
in the filament are unknown.
On the other hand, the rotation between the two actin subunits in the structure of 3W-Actin
is approximately −33°, i.e. −60° compared with a longitudinal dimer of the actin filament 3;
4 (Fig. 2D). As a result, the longitudinal contacts observed in other crystal dimers are
generally broken in the structure of 3W-Actin (Fig. 3), whereas the contacts involving
subdomain 2 are unresolved. Therefore, it appears that the presence of the W domain at the
interface between actin subunits breaks the natural tendency that actin has to preserve
filament-like longitudinal contacts in crystal structures, and induces a rotation between actin
subunits that is of similar magnitude but opposite direction to that of the filament (−33° vs
27°). These results are generally consistent with our previous SAXS studies 22, which
revealed an extended (pseudo long-pitch) arrangement of the actin subunits stabilized by
tandem W domains. However, the SAXS envelope lacked the resolution to distinguish
between the dimer observed here in the crystal structure of 3W-actin and a longitudinal
dimer of the actin filament model.
It is interesting to note that there is also a crystal contact in the structure of 3W-Actin
(between two adjacent dimers) that resembles the dimer of the asymmetric unit. This so-
called ‘crystal’ dimer differs even more significantly from both the actin filament and the
other actin dimers described above, due to an overall translation of ~8 Å between actin
subunits compared to the dimer of the asymmetric unit (Fig. 4). In the crystal dimer, the
crosslink with construct 3W is at the interface between actin subunits, which may explain
the added translation. However, it is significant that the actin subunits of both the non-
crystallographic and crystal dimers are rotated counterclockwise by about the same amount
compared to all the other actin dimers observed in crystal structures, suggesting that this is a
general constraint imposed by the W domain at the interface between actin subunits.
We conclude that while Spire-like tandem W domains can bring actin subunits into close
proximity for nucleation, the conformation of the polymerization nucleus that they form
differs significantly from that of the actin filament. This may explain their weak nucleation
activity as analyzed next.
Long-pitch nucleation by tandem W domains is suboptimal
The structural results prompted us to test the polymerization activity of construct 3W as
compared to those of the prototypical tandem W domain nucleator Spire, which stabilizes a
long-pitch nucleus, and the Arp2/3 complex, which forms a short-pitch nucleus. The
nucleation activity of Drosophila Spire 14 has been mapped to the fragment Spire366–482
comprising the four W domains (Fig. 5A), which was used in the current study. We used the
pyrene-actin polymerization assay to study the effect of Spire366–482 on the polymerization
of 2 µM actin (6% pyrene labeled) by monitoring the fluorescence increase resulting from
the incorporation of labeled actin monomers into the filament (Fig. 5B). At the concentration
of 25 nM, Spire366–482 had very little effect on actin polymerization (polymerization rate
1.0±0.2 nM/sec as compared to 0.8±0.1 nM/sec for actin alone), whereas the Arp2/3
complex activated by the WCA fragment of mouse N-WASP showed a major increase in
polymerization (polymerization rate 31.5±1 nM/sec). However, the nucleation activity of
Spire366–482 increased with concentration, becoming a stronger nucleator at 250 nM
(polymerization rate 4.8±0.2 nM/sec). The opposite effect was observed with construct 3W,
which had no effect on actin polymerization at the concentration of 25 nM, but inhibited
polymerization when used at 250 nM. This could be an indication that construct 3W, like
Tβ4, sequesters actin monomers (Fig. 5B).
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Tβ4 is a short 43-aa polypeptide related to the W domain 8; 9, but it contains an additional
helix at the C-terminus that binds atop actin subdomains 2 and 4 34 (Fig. 5A). As a result,
and despite the apparent simplicity of its helix-loop-helix design, Tβ4 has the ability to
block actin monomer addition to both the pointed and barbed ends of the actin filament,
making it an extremely effective monomer sequestering protein 35; 36. Some proteins
contain tandem repeats of the Tβ4 fold. Examples include, Acanthamoeba castellanii
actobindin 37, Drosophila melanogaster ciboulot 12 and Caenorhabditis elegans
tetrathymosin 38, which respectively contain two-and-a-half, three and four copies of the
Tβ4 fold. Contrary to tandem repeats of the W domain, that frequently mediate filament
nucleation 7, tandem Tβ4 proteins are characterized by their ability to sequester actin
monomers 37; 38. Therefore, we asked whether 3W, consisting of a tandem repeat of three
W domains followed by the C-terminal helix of Tβ4 (Fig. 5A), would sequester actin
monomers. A concentration-dependence analysis of steady-state actin polymerization
revealed that construct 3W sequesters actin monomers even more effectively than Tβ4 (Fig.
5C). We had previously shown, using analytical ultracentrifugation, light scattering and
native gel electrophoresis, that 3W binds three actin monomers in solution 22, which may
explain its stronger sequestering activity compared to Tβ4. Therefore, the effect of 3W on
actin polymerization is more closely related to that of tetrathymosin, which binds and
sequesters multiple actin monomers 38. Although actobindin and ciboulot also sequester
actin monomers, perhaps surprisingly they form 1:1 complexes with actin, indicating that
only one of their actin-binding sites is fully functional 12; 37. It thus appears that the simple
addition of the pointed end capping helix of Tβ4 to tandem W domains changes their
activity from nucleation, as in Spire 14, to monomer sequestration as in Tβ4 35; 36.
CONCLUSIONS
The crystal structure of crosslinked WxActin was found to be nearly undistinguishable from
those of non-crosslinked W-Actin complexes. We have used W domain crosslinking in this
work, as well as in two previous studies 22; 23. The finding that the structure is not altered
in a significant way by the crosslink suggests that this is a structurally sound approach that
can be used as a way to stabilize large polymerization complexes, which are intrinsically
dynamic, for structural investigation.
Various proteins contain tandem repeats of the W domain 7; 8; 9. While the W domain itself
presents well-conserved features (N-terminal helix and LKKT(V) motif), the linkers
between W domains are highly variable, and no single structure can be fully representative
of this large family of proteins. Irrespective of this variability, the inter-W linkers can be
sub-divided into two subgroups: short (as in Spire and N-WASP) and long (as in Cobl
linker-2). While 3W is a synthetic construct with no natural counterpart, it is based on the
tandem W repeat of N-WASP, and it therefore represents the short inter-W linker subgroup.
One general implication of the structure of its complex with actin is that the binding of the
W domain is intrinsically incompatible with inter-subunit contacts along the long-pitch helix
of the actin filament, which is contrary to what we had anticipated 3; 4. The structure of 3W-
Actin further suggests that the actin subunits recruited by tandem W domains with short
inter-W linkers are positioned in a way that resembles the long-pitch helix of the actin
filament, a conformation that would be expected to favor polymerization. However, due to
steric hindrance of the W domain, the contacts between actin subunits in these complexes
differ significantly from those of the actin filament. This may explain the weak nucleation
activity of Spire as compared to the Arp2/3 complex, formins, Cobl and Lmod, all proteins
that are thought to stabilize short-pitch actin nuclei to initiate polymerization. The
incompatibility of the W domain with longitudinal inter-subunit contacts in the filament also
implies that when the actin nucleus transitions into a filament and begins to elongate,
tandem W domain nucleators cannot stay bound to newly formed filaments, and would
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therefore be unlikely to influence elongation. Steric hindrance of the W domain may also be
a contributing factor in the release of NPFs of the Arp2/3 complex from branch junctions
once the branch filament begins to elongate (conformational changes within the Arp2/3
complex itself could be another factor). We finally found that the simple addition of the C-
terminal pointed end-capping helix of Tβ4 to tandem W domains can change their activity
from actin filament nucleation to monomer sequestration.
MATERIALS AND METHODS
Preparation of proteins and protein complexes
Detailed descriptions of the preparation and characterization of the complex of 3W-Actin 22
and the purification of the Arp2/3 complex from bovine brain and preparation of the WCA
fragment of mouse N-WASP 23 were reported previously. Actin was purified from rabbit
skeletal muscle 39. Tβ4 and the first W domain (amino acids Ser-130 to Ser-160) of Vibrio
parahemolyticus VopL (UniProt accession code: Q87GE5) were made as synthetic peptides,
and purified by reverse-phase chromatography. During peptide synthesis, an amino acid
substitution was made (Val-131->Cys) in the first W domain of VopL, a position chosen
based on analysis of the various W-Actin structures 11; 12; 13 as the most favorable for
crosslinking to actin Cys-374. The crosslinking reaction was performed by activation of the
W domain peptide with DTNB (5,5'-dithiobis(2-nitrobenzoic acid)), before mixing with
actin at an actin:W peptide ratio of 1:1.2. The crosslinked fraction was then separated by gel
filtration on a S200 column (Pfizer-Pharmacia) in 25 mM Tris-HCl pH 7.5, 100 mM NaCl,
0.2 mM CaCl2 and 0.2 mM ATP.
The fragment 366–482 of Drosophila Spire (Spire366–482), comprising the four W domains,
was amplified by PCR from cDNA purchased from Open Biosystems. The PCR product was
cloned between the NdeI and EcoRI sites of vector pTYB12 (New England BioLabs).
Protein expression was performed in BL21(DE3) cells (Invitrogen) grown in Terrific Broth
medium at 37°C until the OD600 was 1.0–1.2. Expression was induced with addition of 0.5
mM isopropylthio-β-D-galactoside for 5 h at 20°C. Cells were resuspended in chitin-column
equilibration buffer [20 mM HEPES (pH 7.5), 500 mM NaCl, 1 mM EDTA, and 100 µM
PMSF]. After purification on the chitin affinity column and release of the protein by DTT-
induced autocleavage of the intein, Spire366–482 was additionally purified on a reverse-phase
C18 column (0.1% trifluoroacetic acid and 0–90% acetonitrile), and then dialyzed
extensively against 25 mM Tris-HCl pH 7.5, 100 mM NaCl.
Crystallization of the complexes of 3W-Actin and WxActin
The complex of 3W-Actin (consisting of a tandem repeat of three W domains with three
actin subunits bound 22) was dialyzed against 20 mM HEPES pH 7.5, 100 mM NaCl, 0.2
mM CaCl2 and 0.2 mM ATP and concentrated to 15 mg/ml using an Amicon centrifugal
filter (Millipore). Needle-like crystals grow within hours, or even minutes, using the hanging
drop vapor diffusion method at 20°C, and from drops consisting of a 1:1 (v/v) mixture of
protein solution and a well solution containing 100 mM CAPS pH 10.0, and 24%
polyethylene glycol 3350. However, these crystals did not diffract the X-rays. Crystal
quality and diffraction were improved with addition of 10–100 mM RbCl or
polyvinylpyrrolidone K15 as additives. The crystals were flash-frozen in liquid nitrogen,
with addition of 20% glycerol as cryoprotectant. The crosslinked complex WxActin was
concentrated to 5 mg/ml, and crystallized using the hanging drop vapor diffusion method at
20°C from a well solution containing 0.2 M LiNO3 and 20% polyethylene glycol 3350.
The content of the crystals of 3W-Actin was analyzed by non-reducing gel electrophoresis
and mass spectrometry, using a Voyager DE Pro MALDI-TOF Mass Spectrometer (Applied
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Biosystems) and sinapinic acid as a matrix. For this analysis, multiple crystals were
collected and washed five times through the crystallization solution by transferring them
with a cryo-loop, and then dissolved in water.
Data collection and determination of the structures
X-ray datasets were collected from crystals of WxActin and 3W-Actin at the beamline 17-
BM of the IMCA-CAT facility of the Advance Photon Source (Argonne, IL). Data
indexation and scaling were carried out with the program HKL2000 (HKL Research, Inc.).
The crystals of 3W-Actin diffracted only to 7.0 Å resolution (Table 1). The data in the last
resolution shell (7.25 – 7.0 Å) is weak (I/σ 1.1) and only 34.3% complete. Yet, ~70% of the
data was obtained between 7.9 – 7.0 Å, with average I/σ of 2.1 and redundancy of 6. This
range includes 449 reflections (~20% of the total). Because of the limited resolution, special
emphasis was placed on obtaining a highly redundant dataset (the average redundancy is
20.5 for the entire dataset), which should minimize intensity errors.
The structure of WxActin was determined by molecular replacement, using as search model
the structure of actin complexed with the W domain of WASP (PDB code: 2A3Z).
Molecular replacement and refinement were carried out with the program Phenix 29, and
model building was performed with the program Coot 40.
The structure of 3W-Actin was determined by molecular replacement, using the stronger
data between 15 and 8 Å resolution, and independently with two different programs; Phaser
30 belonging to the Phenix package 29 and AMoRe 31. The two programs gave the same
solution. The likelihood-based scoring function (LLG) of the program Phenix is highly
sensitive to the quality of the search model 41. Several search models were tested, including
monomeric actin 42, complexes of W-Actin determined as ternary complexes with DNase I
11; 13, the complex of ciboulot-actin with bound latrunculin A 12, the structure of actin with
the C-terminal portion of Tβ4 34, and the structure of WxActin determined here. The best-
contrasted solution was obtained using the structure of WxActin as search model. Two
different models were prepared based on this structure, one consisting of the entire
crosslinked complex and one lacking the crosslinked portion (i.e. the last 5 amino acids of
actin and the first 3 amino acids of the W domain). These two models were positioned
independently using a multibody-body search, and clearly defined the locations of the first
(crosslinked) and the second (non-crosslinked) actin subunits of the dimer.
While Phenix was used in the automated mode, a more exhaustive search was performed
with the program AMoRe (details in Supplementary Material). AMoRe’s self-rotation
function gave a single prominent peak with correlation coefficient 0.62. Thus, while the
volume of the unit cell seemed to be compatible with the presence of three W-Actin
complexes in the asymmetric unit (corresponding to a Matthews’ coefficient Vm of 2.15 Å3/
Da and a solvent content of 43%), only two were found (for a Vm of 3.23 Å3/Da and a
solvent content of 62%). We tested many possible configurations in which the orientation of
one W-Actin complex was constrained with respect to the other according to the non-
crystallographic two-fold axis resulting from the self-rotation function. This gave a clearly
contrasted solution for two W-Actin complexes, where the correlation between calculated
and observed structure factor amplitudes was 0.66 (0.50 for the next peak that was not
contrasted above background). Because of the limited resolution, the only refinement
performed after molecular replacement was by rigid body, and using all the diffraction data
available.
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Actin Polymerization Assay
Pyrene-actin polymerization assays were carried out and analyzed as described 43, using a
Cary Eclipse fluorescence spectrophotometer (Varian). All the experiments were performed
at 20°C. Prior to data acquisition, 2 µM Mg-ATP-actin (6% pyrene-labeled) was mixed with
different concentrations of construct 3W (25 nM, 250 nM or 1 µM), Tβ4 (1 µM), and Spire
(25 nM, 250 nM) in F-buffer (10 mM Tris-HCl pH 7.5, 1 mM MgCl2, 50 mM KCl, 1 mM
EGTA, 0.02 mg/mL BSA, 0.2 mM ATP, 1 mM DTT, 0.1 mM NaN3). Note that the addition
of DTT prevents crosslinking of construct 3W to actin Cys-374 during the polymerization
assay. Polymerization rates were measured from the slope of the polymerization curve at
50% polymerization and converted to nM/sec assuming that the total concentration of
polymerizable actin monomers is 1.9 µM (2 µM – 0.1 µM, i.e. by subtracting the critical
concentration for actin monomer addition to the barbed-end from the total concentration of
actin) 43. Steady-state experiments with varying Tβ4 or 3W concentrations were carried out
under similar condition by allowing actin to polymerize for 16h.
Miscellaneous
The program DynDom 44 was used to calculate the relative rotation of actin subunits in
crystal structures of longitudinal actin dimers. Illustrations of the structures were prepared
with the program PyMol (DeLano Scientific LLC).
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
Supported by National Institutes of Health grant P01 HL086655. Use of IMCA-CAT beamline 17-BM was
supported by the Industrial Macromolecular Crystallography Association through a contract with Hauptman-
Woodward Medical Research Institute. The Advanced Photon Source was supported by Department of Energy
Contract W-31-109-Eng-38.
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FIG. 1.
Structure of WxActin. (A) Two perpendicular views of the structure of WxActin. The inset
shows the 2Fo-Fc electron density map (contoured at 1σ) in the region around the crosslink.
Although the crosslink was visualized, this is one of less well-defined regions of the map.
(B) Superimposition of the structures of WxActin (blue, actin; red, W domain) and the non-
crosslinked complex of actin with the W domain of WASP (pink, actin; yellow, W domain),
showing the similarity of the structures. (Two Column Figure).
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FIG. 2.
Structure of 3W-Actin. (A) Non-reducing gel electrophoresis and mass spectrometry
analysis indicate that the crystals of 3W-Actin consist of a 50/50 mixture of actin
crosslinked to construct 3W (expected mass 53,021 Da) and non-crosslinked actin. Actin is
also shown in the gel as a control. (B) Illustration of the actin dimer in the structure of 3W-
Actin. The linker between W domains was modeled. (C) Illustration of a longitudinal actin
dimer from the actin filament model 4. (D) Comparisons of the relative rotations between
actin subunits in the actin filament model (gray and magenta) and the structures of 3W-
Actin and three representative actin dimers observed in crystal structures (including non-
crystallographic and symmetry-related dimers, see also Table 2). For this comparison, the
structures were superimposed using as reference the lower actin subunit (gray), which is
only shown for the filament model. Note that compared to a long-pitch dimer of the actin
filament, in which subunits are rotated by ~27° (magenta arrow), there is a −60° rotation
between the two crystallographically independent actin subunits in the structure of 3W-
Actin. Other dimers observed in crystal structures tend to be flat due to symmetry
constraints and are therefore rotated −27° relative to a longitudinal dimer of the filament.
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The relative rotations between actin subunits were calculated with the program DynDom 44.
(Two Column Figure)
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FIG. 3.
Inter-subunit contacts in the structure of 3W-Actin compared to those of crystallographic
actin dimers. Insets show that longitudinal contacts between subdomains 4 and 3 of adjacent
actin subunits observed in various crystal structures (right) are mostly broken in the structure
of 3W-Actin (left). Representative distances between Cβ atoms are shown for reference.
(Two Column Figure)
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FIG. 4.
Comparison of the non-crystallographic and crystallographic dimers in the structure of 3W-
Actin. (A) Representation of two consecutive dimers related by crystal symmetry. (B)
Superimposition of the non-crystallographic (yellow-blue and red W domains) and
crystallographic (blue-yellow and magenta W domains) dimers. Note that despite their
general similarity, the crystallographic dimer differs more significantly from other actin
dimers observed in crystals structures (Table 2) and the actin filament model. While the
actin subunits in the crystallographic dimer are rotated ~13° relative to the non-
crystallographic dimer, which undoes part of the initial −60° rotation, there is also a
translation of ~8 Å, probably imposed by steric hindrance with the crosslinked W domain. It
is nonetheless significant that the actin subunits of both the non-crystallographic and
crystallographic dimers are rotated counterclockwise by about the same amount compared to
all the other dimers observed in crystal structures, which are generally unwound (see Fig. 2),
suggesting that this is a general property of the W domain at the interface between actin
subunits. (Two Column Figure)
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FIG. 5.
Different effects of Spire, 3W, and Tβ4 on actin polymerization. (A) Schematic diagram of
Tβ4, the four W domain region of Drosophila Spire, and construct 3W. Note that construct
3W consists of three W domains (occurring naturally in mouse N-WASP) separated by short
linkers (as in Spire) and the pointed end capping helix of Tβ4. This construct also contains a
Cys residue at the N-terminus that was crosslinked to actin Cys-374 for crystallization, but
the crosslink was reduced with DTT to measure the nucleation activity. (B) Time course of
polymerization of 2 µM Mg-ATP-actin (6% pyrene-labeled) alone (black) or in the presence
of different concentrations of Spire366–482 (different shades of blue), construct 3W (different
shades of green), Tβ4 (pink), and 25 nM Arp2/3 complex with 250 nM mouse N-WASP
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WCA (red). Each experiment was repeated at least three times. The polymerization rates are:
Actin (0.8±0.1 nM/sec), Spire (1.0±0.2 nM/sec at 25nM and 4.8±0.2 nM/sec at 250 nM),
Arp2/3 complex (31.5±1 nM/sec). (C) Steady-state concentration-dependence of actin
monomer sequestration by Tβ4 and 3W. (One Column Figure)
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Table 1
Crystallographic Data and Refinement Statistics
WxActin
3W-Actin
Diffraction data
Wavelength (Å)
1.0
1.0
Space group
P 212121
P 65 2 2
Unit cell a/b/c (Å)
66.6 / 76.4 / 86.1
100.7 / 100.7 / 458.8
Unit cell α/β/γ (°)
90.0 / 90.0 / 90.0
90.0 / 90.0 / 120.0
Resolution (Å)
50.0-2.89 (2.99–2.89)
50.0-7.0 (7.9–7.0)
Unique reflections
10207
2182
Completeness (%)
99.2 (92.5)
90.0 (70.1)
Redundancy
12.9 (6.4)
20.5 (6.0)
Rmergea (%)
16.8 (46.1)
8.6 (37.3)
I/σ
16.3 (1.8)
16.5 (2.1)
Refinement
Resolution (Å)
37.51–2.89
Atoms used in refinement
3058
Rfactorb (%)
21.2
Rfreec (%)
26.5
No atomic refinement was performed
Rmsd bond lengths (Å)
0.011
Rmsd bond angles (°)
1.910
Average B factors (Å2)
All atoms
62.90
Protein atoms
62.92
Solvent
58.57
Residues in Ramachandran plot
Most favored regions (%)
90.2
Other allowed regions (%)
9.8
PDB accession code
3M1F
3M3N
Values in parentheses correspond to highest resolution shell
aRmerge=Σhkl(I-<I>)/ΣI, where I and <I> are the observed and mean intensities of all the observations of reflection hkl, including its symmetry-
related equivalents
bRfactor=Σhkl∥Fobs| - |Fcalc∥/Σ|Fobs|, where Fobs and Fcalc are the observed and calculated structure factors of reflection hkl
cRfree, Rfactor calculated for a randomly selected subset of reflections (5%) that were not used in refinement
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Table 2
Structures of Longitudinal Actin Dimers.
PDB
actins
per AU
Description
Symmetry, Resolution (Å) and
Cell a, b, c (Å), α, β, γ (°)
References
3M3N
2
Dimer stabilized by tandem W
domains
P6522, 7.0
100.7, 100.7, 458.8, 90.0, 90.0, 120.0
This work
2HF3 2HF4
1
non-polymerizable actin mutant
(Ala-204Glu/Pro-243Lys)
C2, 1.8
199.7, 54.1, 39.6, 90.0, 93.2, 90.0
42
2ASM 2ASO
1
Complexes with marine
macrolides
C2, 1.6
171.2, 54.7, 40.7, 90.0, 96.0, 90.00
45; 46
2ASP
or
2FXU
C2, 1.35
60.1, 56.5, 101.7, 90.0, 94.6, 90.0
2A5X
1
Longitudinally crosslinked actin
dimer
C2, 2.49
207.4, 54.4, 36.2, 90.0, 98.6, 90.0
47
2Q1N 2Q31
2
Longitudinally crosslinked actin
dimer
P21, 2.7
108.1, 71.8, 54.8, 90.0, 104.7, 90.0
32
1Y64
1
Complex with formin homology 2
domain
C2, 3.05
232.0, 56.2, 100.9, 90.0 107.7, 90.0
48
2HMP
2
Non-polymerizable actin, cleaved
between Gly42 and Val43
P212121, 1.9
64, 198, 69.6, 90.0, 90.0, 90.0
49
J Mol Biol. Author manuscript; available in PMC 2011 October 15.
|
3M3R
|
Crystal structure of the M113F alpha-hemolysin mutant complexed with beta-cyclodextrin
|
Molecular bases of cyclodextrin adapter interactions
with engineered protein nanopores
Arijit Banerjeea, Ellina Mikhailovaa, Stephen Cheleya, Li-Qun Gub,2, Michelle Montoyac,3, Yasuo Nagaokaa,4,
Eric Gouauxd, and Hagan Bayleya,1
aDepartment of Chemistry, University of Oxford, Oxford, OX1 3TA, United Kingdom; bDepartment of Medical Biochemistry & Genetics, Texas A&M
University System Health Science Center, College Station, TX 77843-1114; cDepartment of Biochemistry and Molecular Biophysics and Howard Hughes
Medical Institute, Columbia University, New York, NY 10032; and dVollum Institute and Howard Hughes Medical Institute, Oregon Health and Science
University, Portland, OR 97239
Edited by Gregory A. Petsko, Brandeis University, Waltham, MA, and approved March 3, 2010 (received for review December 15, 2009)
Engineered protein pores have several potential applications in
biotechnology: as sensor elements in stochastic detection and
ultrarapid DNA sequencing, as nanoreactors to observe single-
molecule chemistry, and in the construction of nano- and micro-
devices. One important class of pores contains molecular adapters,
which provide internal binding sites for small molecules. Mutants
of the α-hemolysin (αHL) pore that bind the adapter β-cyclodextrin
(βCD) ∼104 times more tightly than the wild type have been ob-
tained. We now use single-channel electrical recording, protein en-
gineering including unnatural amino acid mutagenesis, and high-
resolution x-ray crystallography to provide definitive structural in-
formation on these engineered protein nanopores in unparalleled
detail.
alpha-hemolysin ∣single molecule ∣stochastic sensing ∣structure ∣
unnatural amino acid
M
any research groups have used protein engineering to
obtain enzymes and antibodies with new properties suited
for specific tasks (1–6). Fewer groups have taken on the difficult
problem of engineering membrane proteins (7). We have engi-
neered the α-hemolysin protein pore, mindful of several potential
applications in biotechnology, including its ability to act as a de-
tector in stochastic sensing (8) and ultrarapid DNA sequencing
(9), to serve as a nanoreactor for the observation of single-
molecule chemistry (10) and to act as a component for the con-
struction of nano- and microdevices (11).
An important breakthrough in this area, which enabled the sto-
chastic sensing of organic molecules including the detection of
DNA bases in the form of nucleoside monophosphates (12, 13),
was the discovery of internal molecular adapters, a form of non-
covalent protein modification (14). Most useful have been cyclo-
dextrin (CD) adapters, which have until now been used in the
absence of detailed structural information about how they work.
The present paper is a definitive investigation, which provides
such information through the application of a wide variety of
technical approaches: single-channel electrical recording, protein
engineering including unnatural amino acid mutagenesis, and
x-ray crystallography. The studies employing mutagenesis show
that the striking interactions seen in the crystal structures also
occur in individual pores in lipid bilayers.
We reveal that the tight-binding αHL mutants (15) M113N7
and M113F7 bind βCD in different orientations within the hep-
tameric pore. In the case of M113N7, the top (primary hydroxyls)
of the CD ring faces the trans entrance of the pore. In the case of
M113F7, the bottom (secondary hydroxyls) of the CD ring faces
the trans entrance, while the top of the ring is bonded to the pore
through remarkable CH-π interactions. Another tight-binding
mutant, M113V7, can bind the CD in both orientations. These
results illustrate the exquisite level of engineering that can be
achieved with protein nanopores, which is, for example, far be-
yond what is possible with solid-state pores. The work also pro-
vides information valuable for the design of new binding sites
within the lumen of the αHL pore or within other β-barrel pro-
teins. Our results will be of interest to others exploring the inter-
actions of CDs with the αHL pore (16, 17), including groups
involved in computational studies (18, 19). In addition CDs bind
to a variety of other pores, including porins (20, 21) and connex-
ins (22), and are being tested in vivo as blockers of the anthrax
protective antigen pore (23, 24). The CD adapter concept has
also been incorporated into other formats, e.g., with glass nano-
pores (25), and artificial pores based on CDs have been made by
several groups (26–28). Our work is pertinent to these studies.
Results
Kinetics and Thermodynamics of the Interactions of βCD with αHL
Pores Containing Met, Phe and Asn at Position 113. We showed earlier
that position 113 in the αHL pore (Fig. 1A) is critical for the bind-
ing of βCD (14). Subsequently, residue 113, which is Met in the
WT protein, was changed to each of the remaining 19 naturally
occurring amino acids by site-directed mutagenesis (15). We
found that 11 of these mutants, expressed as homoheptamers,
bound βCD with a similar affinity and with similar kinetics to
the WT homoheptamer. Two mutants (P, W) bound βCD about
10 times more strongly than the WT homoheptamer, while six of
them (V, H, Y, D, N, F) bound with high affinity, i.e., with a Kd
value 103 to 104 times lower than the WT.
Remarkably, the side chains of the latter six amino acids bear
little resemblance to one another, and this issue is addressed in the
present paper. We first examined the two amino acids with the
most disparate side chains (Fand N) by making heteromeric pores
containing WT (Met-113), M113F, and M113N subunits. Three
series of heteroheptamers were produced: WT7−nM113Nn,
WT7−nM113Fn, and M113F7−nM113Nn. The heteroheptamers
were separated by SDS-polyacrylamide gel electrophoresis aided
by an oligoaspartate (D8) tail on the first of the two types of sub-
unit (Fig. 1B) (29). All 21 combinations of WT, M113F, and
M113N subunits formed αHL pores that interacted with βCD as
shown by single-channel current recordings, which revealed the
extent of block by βCD (Fig. S1), the association and dissociation
Author contributions: A.B., S.C., E.G., and H.B. designed research; A.B., E.M., S.C., L.-Q.G.,
M.M., and Y.N. performed research; A.B., E.G., and H.B. analyzed data; and A.B. and
H.B. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
1To whom correspondence should be addressed. E-mail: hagan.bayley@chem.ox.ac.uk.
2Present address: Department of Biological Engineering and Dalton Cardiovascular
Research Center, University of Missouri, Columbia, MO 65211.
3Present address: Nature Structural & Molecular Biology, 75 Varick Street, 9th Floor, New
York NY 10013-1917.
4Present address: Department of Biotechnology, Faculty of Engineering, Kansai University,
3-3-35 Yamate-cho, Suita, Osaka 564-8680, Japan.
This article contains supporting information online at www.pnas.org/cgi/content/full/
0914229107/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.0914229107
PNAS ∣May 4, 2010 ∣vol. 107 ∣no. 18 ∣8165–8170
BIOCHEMISTRY
rate constants for βCD (kon and koff), and (from the latter) the
equilibrium dissociation constant for βCD (Kd ¼ koff∕kon) (15).
The kon values for βCD for the 21 combinations of subunits
were all similar at ∼5 × 105 M−1 s−1 (Fig. 1C, Upper). By contrast,
the koff values differed widely, ranging from ∼5 × 10−2 s−1 to
∼103 s−1. For WT7−nM113Nn and WT7−nM113Fn, the koff values
decreased as M113N or M113F subunits were added. In the case
of M113N, there was a steep drop in the value of koff after the
fifth subunit had been incorporated. In the case of M113F,
the decrease in the value of koff occurred less precipitously as the
M113F subunits were added (Fig. 1C, Lower). Intriguingly, with
M113F7−nM113Nn, koff first increased as M113N subunits were
added to M113F7 until n ¼ 4 (M113F3M113N4) and then de-
creased for larger values of n (Fig. 1C, Lower). We recognize that
there is more than one permutation of heteromers containing two
to five mutant subunits (Fig. 1B), but we have ignored this fact
here because no significant differences in the properties of indi-
vidual heteromers were observed. For example, 42 recordings
were made of WT5M113N2, which has three permutations.
Because, kon showed little variation with subunit composition,
the variation in Kd was similar to the variation in koff (Fig. 1C).
While these studies were in progress, the crystal structures of
βCD complexed to M113N7 (Fig. 2B) and M113F7 (Fig. 2C) were
solved (Table S1) (30). High-resolution structures could be
obtained because the CD and the αHL pore have the same C7
symmetry. In the case of M113N7, βCD is bound with the second-
ary hydroxyl face “upward” (Fig. 2B). In each glucose unit of the
βCD, the 2-hydroxyl is hydrogen bonded to the side-chain amide
of an Asn-113 (the residue introduced by mutagenesis) and the
3-hydroxyl is hydrogen bonded to the ϵ-amino group of Lys-147.
In the case of M113F7, two βCDs are bound to the αHL pore
(Fig. 2C). It is the top βCD in the structure that concerns us, be-
cause it is in contact with the Phe-113 residues introduced by mu-
tagenesis. It is immediately apparent that the top βCD in M113F7
is in the opposite orientation to the βCD in M113N7 with each
6-hydroxyl group in a CH-π bonding interaction (31–35) with a
Phe-113 side chain. The opposite orientations of the βCDs in
M113N7 and M113F7 immediately explain why heteromers
formed from similar numbers of M113N and M113F subunits
(e.g., M113N4M113F3) bind βCD weakly (see also Discussion).
Unnatural Amino Acid Mutagenesis. To further explore the range of
noncovalent interactions that are available when βCD binds to
the αHL pore, five unnatural amino acids (Fig. 3A and Fig. S2)
were incorporated at position 113, by using the in vitro nonsense
codon suppression method (36). In particular, we had noted that
M113V7 containing the β-branched Val binds βCD tightly (15),
and therefore we compared cyclopropylglycine (Cpg) and cyclo-
propylalanine (Cpa). We also further examined the means by
which M113F7 binds βCD tightly, by comparing the properties of
4-fluorophenylalanine (f1F), pentafluorophenylalanine (f5F),
and cyclohexylalanine (Cha) at position 113.
The five homomeric pores all produced single-channel cur-
rents with unitary conductance values in the range expected
for properly assembled heptamers (Fig. S3). All five bound βCD
(Fig. 3B, Level 2), either tightly (f1F, Cpg) or weakly (f5F, Cha,
Cpa) as described in detail below. During the long βCD binding
events, additional current spikes were seen (Fig. 3B). Similar
Fig. 1.
Binding of βCD by heteromeric αHL pores formed by WT, M113F and M113N subunits. (A) Crystal structure of WT-αHL (61) showing residue 113 (Met,
yellow). Left panel, side view and right panel, top view. (B) Separation of 35S-labeled αHL heteroheptamers by SDS-polyacrylamide electrophoresis. The
separation of the M113F7-nM113Nn heteromers is shown as detected by autoradiography of a dried gel. The M113F subunits carried a D8 tail. Lane 1,
M113N7; lane 2, M113F7−nM113Nn (the heteromers formed from several preparations made with differing ratios of M113F and M113N subunits were mixed
to give roughly equal amounts of each subunit combination); lane 3, M113F7. A diagram of the eight different combinations of subunits and their permuta-
tions is shown to the right of the autoradiogram. The various permutations are not separated by electrophoresis. (C) Kinetics of the interaction of βCD with
single heteromeric αHL pores as determined by bilayer recording. Values of kon were calculated by using kon ¼ 1∕ðτon½βCDÞ, where τon is the mean interevent
interval. Values of koff were determined by using koff ¼ 1∕τoff, where τoff is the mean dwell time of βCD in the pore. Values of Kd were calculated by using
Kd ¼ koff∕kon. Each point represents the mean s:d: for three or more determinations. Where they cannot be seen, the s.d. values lie within the symbol. Black
squares, WT7−nM113Nn; gray squares, M113F7−nM113Nn; empty squares, M113F7−nWTn.
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events had been observed previously with certain Met-113 repla-
cement mutants and may represent movement of the βCD at its
binding site (e.g., rotation about axes perpendicular to the C7
axis) (15). The additional current spikes were more prevalent
for M113V7 and M113Cpg7, which may take part in more con-
formationally labile interactions with βCD, compared with say
M113F7 (Fig. S4).
Interactions of βCD with Homoheptamers Bearing Aromatic Residues
at Position 113. To further understand the nature of the binding of
βCD to aromatic side chains, we examined the kinetics of βCD
binding to the homoheptamers containing f1F or f5F at position
113, M113f1F7 and M113f5F7 (Fig. 3C). For both mutants, the
value of kon was very similar to that of WT7, but the values of koff
and therefore Kd for M113f1F7 differed dramatically from WT7
and were close to the values for the tight-binding mutant M113F7
(Table S2A). By contrast, koff and Kd for M113f5F7 were similar
to the values for WT7 (Table S2A).
To determine whether M113f1F7 binds βCD in the same orien-
tation as M113F7 (Fig. 2C), we made heteromers of the M113f1F
subunit with M113N or M113F and examined M113F4M113f1F3
and M113N4M113f1F3. M113F4M113f1F3 binds βCD as strongly
as either M113F7 or M113f1F7, but M113N4M113f1F3 binds
βCD weakly with a similar affinity to WT7 (Fig. 3D and
Table S3). Therefore, it is reasonable to infer that M113F7
and M113f1F7 bind βCD in the same orientation with the 6-
hydroxyl groups of the CD in proximity to the aromatic rings
on the protein.
Cyclohexylalanine (Cha) was used to replace the aromatic side
chains with a roughly isosteric hydrophobic group. Again the va-
lue of kon for βCD was little changed, but koff for M113Cha7 had
an intermediate value of 42 6 s−1. Therefore, M113Cha7 binds
βCD more weakly than M113F7 but distinctly more strongly than
the WT7 pore (Table S2A and Fig. 3C).
Interactions of βCD with Homoheptamers Bearing Hydrophobic Resi-
dues at Position 113. M113V7 binds βCD very strongly, and there-
fore we compared αHL pores with Cpg or Cpa at position 113.
Cpg is roughly isosteric with Val, and like Val has a β-branched
side chain. Gratifyingly, M113Cpg7 has a kon value similar to the
other αHL pores, and koff and Kd values close to those of
M113V7 (Table S2B and Fig. 3E). Cyclopropylalanine (Cpa), with
an additional methylene group compared to Cpg, is roughly
isosteric with Leu, a weak binder, and M113Cpa7 also binds
βCD weakly with kon, koff and Kd values similar to those of
WT7 (Table S2B and Fig. 3E). M113I7 and M113T7, which are
β-branched, are also weak binders, but Ile and Thr are less closely
related to Val than Cpg.
To determine whether M113V7 binds βCD in the same orien-
tation as M113F7 or M113N7 (Fig. 2), we made heteromers of
M113V and the M113N or M113F subunits. M113V3M113F4,
M113V4M113F3, M113V3M113N4, and M113V4M113N3 were
examined in detail. All four heteroheptamers bound βCD more
weakly than M113V7, M113F7 or M113N7 (Fig. 3F and Table S4),
suggesting that Val at position 113 interacts with βCD strongly but
in a different manner to either Phe or Asn. Each heteromer
exhibited a range of Kd values, perhaps reflecting the various pos-
sible permutations of the two different subunits around the cen-
tral axis of the heptamer, although this heterogeneity was not
seen for heteromers made from WT, M113F and M113N (Fig. 1).
Discussion
Soon after we discovered that βCD binds to the WT-αHL pore for
around a millisecond, we found a mutant pore, M113N7, that re-
leases βCD ∼104 times more slowly (14). This prompted us to
examine all 19 mutants in which residue 113 is replaced by a nat-
ural amino acid, with the surprising result that a collection of ami-
no acids with structurally unrelated side chains (V, H, Y, D, N, F)
are tight binders (15). Here, we have examined the nature of the
binding interactions more closely by single-channel electrical re-
cording, protein engineering including unnatural amino acid mu-
tagenesis, and high-resolution x-ray crystallography, and we
provide the first definitive structural information on an engi-
neered protein nanopore.
We find that βCD can bind tightly to the αHL pore in three
different ways depending on the residue at 113, as exemplified
by Asn, Phe, and Val. Because Asn and Phe have quite different
side chains, we first compared the ability of M113N and M113F
subunits to take part in binding the CD. The examination of het-
eromeric proteins containing WT (Met-113), M113N and M113F
subunits showed that the replacement of WT subunits in WT7
with M113N or M113F subunits led to increased affinity for
βCD. The more M113N or M113F subunits that were added, the
tighter binding became. By contrast, when subunits in M113N7
were replaced with M113F subunits, binding became weaker,
reaching a minimum at three to four M113F subunits, and then
increasing in strength with five M113F subunits or more (Fig. 1C).
Parallel structural studies (30) revealed the basis of the “oppos-
ing” effects of the M113N and M113F subunits. βCD binds to
M113N7 in the opposite orientation to that in which it binds
to M113F7. In M113N7, the secondary hydroxyls in the βCD ring
are hydrogen bonded to Lys-147 and Asn-113 (Fig. 2A). By con-
trast, βCD interacts with M113F7 through its primary hydroxyl
face (Fig. 2B).
It seemed likely that M113V7, bound βCD in yet another way,
and this was examined by forming heteromers between M113V
and M113N or M113F. The presence of three or four subunits
of either M113N or M113F greatly decreases the affinity of
the pore for βCD (Fig. 3F), with an average koff of 7.3 × 101 s−1,
indicating that a third binding mode is indeed operating
Fig. 2.
X-ray structures of M113N and
M113F homoheptamers with βCD bound.
(A) Side view of heptameric αHL. βCD binds
in the blue highlighted region. (B) βCD
bound to M113N7 (dotted lines indicate hy-
drogen bonding). The side chains of Lys-147
are in pale brown and the side chains of Asn-
113 in yellow. (C) βCD bound to M113F7
(dotted lines indicate CH-π bonding). The
side chains of Phe-113 are in yellow. The sec-
ond βCD in the M113F7 · ðβCDÞ2 structure is
hydrogen bonded to the top βCD in a head-
to-head arrangement and has no apparent
interactions with the protein. For both (B)
and (C), four β strands were omitted from
the barrel to give a better view.
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(Table S4). In summary, the three groups of tight-binding mutants
comprise αHL pores incorporating, at position 113: (i) the hydro-
gen-bonding amino acids N, D (the latter would have to be largely
in the protonated form), and possibly H; (ii) the aromatics F, Y,
f1F, and possibly H, and more weakly W; (iii) the β-branched ami-
no acids V, Cpg. There may be yet other means by which CDs can
bind to the αHL pore. For example, we earlier found that hepta-
6-sulfato-βCD can bind tightly to αHL pores containing the
N139Q mutation (37). Presumably, this CD is bound at a site low-
er down in the β barrel in a fashion that includes hydrogen bond-
ing to the Gln at position 139. While the various mutants
exhibited widely different koff values, the value of kon was almost
invariant and averaged ∼2.3 × 105 M−1 s−1 (Table S2) (15). Ap-
parently, transport to the binding site is rate limiting, through
a route unaffected by mutagenesis.
koff increased precipitously with the addition of WTsubunits to
M113N7 (Fig. 1C). Crystal structures of M113N7 show that resi-
dues
111,
113,
and
147
are
reorganized
by
compari-
son with WT7 and then undergo a more limited rearrangement
when βCD binds (Fig. S5). For example, the side chain of
Lys-147 shifts position to form a bifurcated hydrogen bond with
a 3-hydroxyl group of βCD and the side chain carbonyl of an Asn-
113 (Fig. S6). Therefore, the side chains of residues 111, 113, and
147 might be in a variety of conformations in WT7−nM113Nn het-
eromers and offer less well preorganized binding sites for βCD
than they do in M113N7. Further, the intramolecular hydrogen
bonds of the secondary hydroxyls in βCD (38) must be disrupted
upon binding as both hydroxyls on each glucose ring form hydro-
gen bonds to the mutant subunits (Fig. 2B). Because the hydrogen
bonds that are broken in βCD are arranged in a circle, the break-
age of bonds involving a single glucose (three bonds in all) will be
relatively more disruptive than those involving adjoining glucose
residues or the entire circle. The overall binding cooperativity in
M113N7 could be attributed to enthalpic cooperativity outweigh-
ing entropic penalties to binding (39). Positive cooperativity has
been observed previously in fairly rigid model systems (40).
By contrast with M113N7, there is little movement of side
chains in ðM113FÞ7 by comparison with WT7 and little move-
ment, including Phe-113, upon binding βCD (Fig. S7A). Further,
the crystal structure of M113F7 · βCD suggests that each Phe re-
sidue interacts independently with the βCD through what appear
to be CH-π interactions (Fig. S7B). These interactions are ex-
pected to be weak and not strongly directional and hence offer
less enthalpic cooperativity, as supported by the B-factors (crys-
tallographic temperatures factors) at the primary βCD binding
site, which are between ∼40 and 50. Positive cooperativity is ob-
served, but it is less pronounced than in the case of M113N7
(Table S5). In the case of M113N7, the B-factors of the residues
that bind βCD are in the 20s implying that the βCD is more rigidly
held than it is in M113F7.
The binding of sugars to aromatic residues in proteins can in-
clude CH-π bonding (41) or OH-π bonding or a finely balanced
complement of both (42, 43). However, we have dismissed the
possibility of an OH-π interaction between Phe-113 and the
6-hydroxyl groups of βCD as the distance between the center
of the phenyl rings to the nearest hydroxyl oxygen is higher
(5.2 0.65 Å, n ¼ 7) than that expected for a favorable OH-π
interaction (33). While we propose that βCD binds to Phe-113
through a C-6 CH-π interaction (Fig. S7B), the distances between
the center of the Phe-113 ring and the nearest C-6 of βCD ob-
served in the M113F7 · βCD structure (4.66 0.24 Å, n ¼ 7)
are in the upper range of the expected distance for a strong inter-
action, which is ∼4.5 Å (33). The angle between the normal to the
aromatic rings and the line connecting the C-6 atoms to the aro-
matic midpoint is 8.0 5.6°, which is well within the expected
range (44). The measurements with M113f5F7 argue against a
hydrophobic interaction between Phe residues at position 113
and the βCD ring. In f5F, the hydrophobicity of the phenyl ring
is significantly increased (45) yet M113f5F7 binds βCD weakly,
like WT7 (Fig. 3C and Table S2A).
By contrast with F, f1F, Y and N, homomeric αHL pores with
f5F and W at position 113 bound βCD relatively weakly (Fig. 3C
and Table S2A). In the case of f5F, the powerful electron with-
drawing action of the five fluorine atoms leaves a highly increased
positive charge at the center of the ring (46, 47), mitigating
against a hydrogen-bonding interaction. The electron-rich Trp
Fig. 3.
Properties of pores containing natural and unnatural amino acid sub-
stitutions at position 113. The data were recorded at þ40 mV in 1.0 M NaCl,
10 mM sodium phosphate, pH 7.5. (A) Unnatural amino acids used in this
study: 4-fluorophenylalanine, f1F; pentafluorophenylalanine, f5F; cyclohex-
ylalanine, Cha; cyclopropylglycine, Cpg; cyclopropylalanine, Cpa. (B) Repre-
sentative
current
traces
from
single
homoheptameric
αHL
pores,
containing unnatural amino acids at position 113, in the presence of βCD.
βCD (40 μM final) was added to the trans chamber. Level 1, open pore current;
level 2, pore occupied by βCD. The broken line indicates zero current. (C) In-
teraction of βCD with homomeric αHL pores containing aromatic amino acids
at position 113. Kd values for the interaction between βCD and the αHL pore
were calculated by using Kd ¼ koff∕kon. Each column represents the mean
s:d: for 10 or more determinations: dark gray, natural amino acids; light gray,
unnatural amino acids. Data adapted from Gu and colleagues (15) are
marked (*). (D) Representative current traces from single-channel recordings
of βCD binding to M113F4M113f1F3 and M113N4M113f1F3. βCD (40 μM final)
was added to the trans chamber. The broken line indicates zero current. (E)
Interaction of βCD with homomeric αHL pores containing hydrophobic amino
acids at position 113. Kd values for the interaction between βCD and the αHL
pore were calculated by using Kd ¼ koff∕kon. Each column represents the
mean s:d: for ten or more determinations: dark gray, natural amino acids;
light gray, unnatural amino acids. Data adapted from Gu and colleagues (15)
are marked (*). (F) koff values for βCD from heteroheptamers formed with
M113F and M113V subunits and with M113N and M113V subunits. βCD
(40 μM final) was added to the trans chamber. The kon values for βCD for
all these mutants are similar, at ∼3 × 105 M−1 s−1. Empty square: average
koff values for the mutant (bar is s:d). Filled square: M113V3M113F4; filled
circle: M113V4M113F3; filled upright triangle: M113V3M113N4; filled in-
verted triangle: M113V4M113N3.
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Banerjee et al.
ring (44, 46, 47) should favor hydrogen bonding, but here we can-
not make a direct comparison with the crystal structure of
M113F7 as the indole ring is far larger than benzene. It is possible
that it cannot become oriented in the same manner and that it is
misaligned for hydrogen bonding.
Our experiments suggest that M113V7 and M113Cpg7 bind
βCD in a third way. In heteromers with M113V, both M113F
and M113N reduce the affinity of the pore for βCD suggesting that
neither the CH-π interaction with Phe-113 nor the hydrogen-
bonding interactions with Asn-113 and Lys-147 are compatible
with binding to Val. Close interactions of Val with glucose rings
have been noted previously (48). Therefore, we propose that the
Val side-chain interacts with the side of the glucose ring. This in-
teraction might occur in one or both orientations of the CD
ring (Fig. 4).
Conclusion
We provide structural information on engineered protein nano-
pores and describe three distinct ways in which βCD can bind
within the lumen of mutant αHL pores in atomic detail. Our re-
sults will be useful in several areas of basic science and biotech-
nology. By using host molecules lodged within the αHL pore,
host-guest interactions can be investigated in fine detail at the
single-molecule level (17, 49). The present work will now permit
us to examine binding events at a known face of a CD. The work
also provides information for designing new binding sites within
the lumen of the αHL pore (37) or within other β barrel proteins
(21, 50) and for using molecular design to devise ways in which to
covalently attach CDs within pores (13, 51). These areas impact
practical applications of nanopore technology including stochas-
tic sensing (8), single-molecule DNA sequencing (9, 12, 13, 52),
the use of nanoreactors for the observation of single-molecule
chemistry (10), and the construction of nano- and microdevices
(11, 53), as well as the design of CDs as therapeutic agents
(23, 24).
Methods
Full details of the experimental procedures can be found in SI Appendix.
Materials
L-Amino acids were obtained as follows: 4-fluorophenylalanine (f1F) (Fluka);
pentafluorophenylalanine (f5F) (PepTech Corp.); cyclopropylglycine (Cpg) (Ty-
ger); cyclopropylalanine (Cpa) (Tyger). 4-N-benzoyl-5′-O-(4,4′-dimethoxytri-
tyl)-2′-deoxycytidine-3′-[(2-cyanoethyl)-(N,N-diisopropyl)]-phosphoramidite
and bis(2-cyanoethyl)-N,N-diisopropylphosphoramidite for the synthesis of
pdCpA were purchased from Glen Research and Toronto Research Chemicals,
respectively.
Preparation
of
NVOC-Protected
Aminoacyl-pdCpA.
NVOC-protected
aminoacyl-pdCpAs were prepared as reported previously by reacting the
dinucleotide pdCpA with N-protected, carboxylic acid-activated, amino
acids (54–56).
Preparation of NVOC-Protected Aminoacyl-tRNA. NVOC-protected aminoacyl-
pdCpAs were ligated enzymatically with a truncated tRNA, prepared by using
methods described elsewhere (57, 58).
Genetic Constructs and Mutagenesis. All new αHL constructs were verified by
DNA sequencing. Details of each construct can be found in SI Appendix.
Synthesis, Assembly, and Purification of Mutant αHL pores. αHL monomers (WT
and mutants) were prepared in vitro by coupled transcription and translation
(IVTT) and assembled into homoheptamers on rabbit red blood cell
membranes followed by purification by SDS–PAGE as described earlier
(59). Heteroheptamers were prepared by mixing the two required DNAs
(one encoding an αHL with a D8 tail) before IVTT and then oligomerizing
the mixed translation products on rabbit red blood cell membranes. Pores
with the desired combinations of subunits were purified by SDS–PAGE (59).
Synthesis, Assembly, and Purification of αHL Mutants Containing Unnatural Ami-
no Acids. αHL polypeptides containing unnatural amino acids were synthe-
sized by IVTT in the presence of rabbit red blood cell membranes. The
plasmid with a stop codon (TAG) at position 113 was used. Deprotected ami-
noacyl-tRNAs (SI Appendix) were added to the IVTT mixtures. For heterohep-
tamers with subunits containing unnatural amino acids in combination with
M113N or M113F, monomers were first made, which were then coassembled
on rabbit red blood cell membranes and subsequently purified by SDS–PAGE.
Single-Channel Current Recordings in Planar Lipid Bilayers. (15, 60) Recordings
were made with 1.0 M NaCl, 10 mM sodium phosphate, pH 7.5, in both cham-
bers, at an applied potential of þ40 mV. Data were recorded at 22 2°C. The
bilayer was formed from 1,2-diphytanoyl-sn-glycero-phosphocholine (Avanti
Polar Lipids). Proteins were added to the cis chamber, and βCD to the trans
chamber. Single-channel currents were recorded with an Axopatch 200B
patch-clamp amplifier (Axon Instruments) and filtered at 2 kHz with a
built-in 4-pole low-pass Bessel Filter. The data were acquired at a sampling
rate of 10 kHz. For mutants that bind βCD strongly, the data were acquired
for at least 30 min and for weak-binding mutants for at least 10 min.
Kinetic Data Analysis. Current amplitude and dwell-time histograms were
made by using ClampFit 9.0. The mean dwell times, τoff, were determined
by fitting the dwell-time histograms to single exponentials. Values of kon
and koff were obtained by using the mean dwell times and mean interevent
intervals, as described previously (15, 60). This analysis assumes a binary in-
teraction, which was supported in all cases examined by the finding of only
one major blockade level and a single exponential distribution of dwell
times (τoff).
Fig. 4.
Molecular model showing the three classes of interaction between
the αHL pore and βCD identified in this work. The model identifies the region
of βCD responsible for each interaction (H atoms interacting with Phe-113 or
Asn-113 and Lys-147: gray). The first class of interaction is with aromatic
residues and involves the seven -CH2OH groups of the βCD. The second class
is typified by the interactions with Asn at position 113, which involve hydro-
gen-bonds to the secondary 2-hydroxyls of the βCD. Structural studies show
that this interaction is supported by hydrogen bonding between Lys-147 and
the secondary 3-hydroxyls of the βCD. Structural studies and experiments
with heteromers suggest that the βCD in M113F7 is in the opposite orienta-
tion to the βCD in M113N7, in support of the model shown here. As the inter-
action with Val is hydrophobic, it is not directional and βCD may not bind at
the same position inside the β barrel as it does in M113F7 or M113N7.
Banerjee et al.
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Protein Crystallography. Details can be found in SI Appendix. Protein Data
Bank: The coordinates and structure factors of the described structures have
been deposited with accession codes 3M2L ðM113F7Þ, 3M3R ðM113F7 · βCDÞ,
3M4D ðM113N7Þ, 3M4E ðM113N7 · βCDÞ.
ACKNOWLEDGMENTS. We thank Dennis Dougherty for the plasmid pTHG73.
This work was funded by a Royal Society Wolfson Research Merit Award
(to H.B.), the Medical Research Council (H.B.), the National Institutes of
Health (H.B.), and the Howard Hughes Medical Institute (E.G.).
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Banerjee et al.
|
3M3W
|
Crystal structure of mouse PACSIN3 BAR domain mutant
|
Rigidity of Wedge Loop in PACSIN 3 Protein Is a Key Factor in
Dictating Diameters of Tubules*□
S
Received for publication,March 4, 2012, and in revised form, April 26, 2012 Published, JBC Papers in Press,May 9, 2012, DOI 10.1074/jbc.M112.358960
Xiaoyun Bai‡§1, Geng Meng‡§1, Ming Luo¶, and Xiaofeng Zheng‡§2
From the ‡State Key Laboratory of Protein and Plant Gene Research, §Department of Biochemistry and Molecular Biology, School
of Life Sciences, Peking University, Beijing 100871, China and the ¶Department of Microbiology, University of Alabama at
Birmingham, Birmingham, Alabama 35294
Background: PACSINs participate in cellular membrane remodeling.
Results: PACSIN 3 F-BAR induces different tubules compared with PACSIN 1 and 2. Structures of PACSINs reveal a novel
wedge loop-mediated lateral interaction and different packing mode in PACSIN 3.
Conclusion: The rigidity of the wedge loop determines the angles between neighboring dimers and further dictates tubule
diameters.
Significance: The study provides novel insights into F-BAR domain-induced membrane deformation.
BAR (Bin/amphiphysin/Rvs) domain-containing proteins
participate in cellular membrane remodeling. The F-BAR pro-
teins normally generate low curvature tubules. However, in
the PACSIN subfamily, the F-BAR domain from PACSIN 1
and 2 can induce both high and low curvature tubules. We
found that unlike PACSIN 1 and 2, PACSIN 3 could only
induce low curvature tubules. To elucidate the key factors
that dictate the tubule curvature, crystal structures of all
three PACSIN F-BAR domains were determined. A novel type
of lateral interaction mediated by a wedge loop is observed
between the F-BAR neighboring dimers. Comparisons of the
structures of PACSIN 3 with PACSIN 1 and 2 indicate that the
wedge loop of PACSIN 3 is more rigid, which influences
the lateral interactions between assembled dimers. We fur-
ther identified the residues that affect the rigidity of the loop
by mutagenesis and determined the structures of two PAC-
SIN 3 wedge loop mutants. Our results suggest that the rigid-
ity-mediated conformations of the wedge loop correlate well
with the various crystal packing modes and membrane tubu-
lations. Thus, the rigidity of the wedge loop is a key factor in
dictating tubule diameters.
Cellular membrane deformation is important in the process
of cargo transportation and cell movement (1–5). Membrane
remodeling is induced by the packing of protein oligomers on
the negatively charged membrane surface (6–8). The Bin/am-
phiphysin/Rvs (BAR)3 domain proteins, including N-BAR
(N-terminal amphipathic helix-BAR), EFC/F-BAR (Fes/CIP4
homology-BAR), and IMD/IBAR (inverse-BAR) domain pro-
teins, are important for membrane remodeling in vesicle bud-
ding, membrane trafficking between intracellular compart-
ments, and cell division (6, 9–14). F-BAR domain proteins
contain a central -helix bundle that stabilizes the membrane
via the positively charged protein surface of homodimer and
bends the membrane into tubules with low curvature (8,
15–17). The crystal structures of FBP17 and CIP4 revealed that
F-BAR domain proteins form filaments through end-to-end
interactions between dimers in the crystal, and the diameters of
the induced tubules were proposed to be related to the intrinsic
large radial curvature of the F-BAR dimer (16). Cryo-EM stud-
ies of CIP4 and FBP17 showed that the F-BAR domain-induced
membrane tubulation is caused by the packing of the protein
helical lattice on the membrane surface via lateral and tip-to-tip
interactions of the F-BAR dimers (18).
PACSINs (also named syndapin) constitute a branch of the
F-BAR domain protein family, which are cytoplasmic proteins
involved in receptor-mediated endocytosis, synaptic vesicle
trafficking, and biogenesis of different cellular organelles (11,
19–26). PACSINs contain the N-terminal F-BAR domain and a
C-terminal mono-Src homology 3 domain. The PACSIN family
contains PACSIN 1, 2, and 3, which differ in their tissue distri-
butions. PACSIN 1 is detected primarily in neuron cells, and
PACSIN 3 is found in lung and muscle tissues, whereas PACSIN 2
is expressed ubiquitously (21, 27, 28). The F-BAR-mediated
membrane deformation of PACSIN 1 is autoinhibited by its Src
homology 3 domain (29). Interactions of PACSINs with other
proteins such as dynamin and the neutal Wiskott-Aldrich syn-
drome protein also play important roles in membrane remod-
eling activities (30).
In contrast to the typical F-BAR domain proteins that mainly
generate low curvature tubules (100 nm in diameter), PACSIN
* This work was supported by National High Technology and Development
Program of China Grant 2010CB911804, National Science Foundation of
China Grant 30930020, and International Centre for Genetic Engineering
and Biotechnology Project CRP/CHN09-01.
□
S This article contains supplemental Figs. S1–S7.
The atomic coordinates and structure factors (codes 3Q84, 3Q0K, 3QE6, 3M3W,
and 3SYV) have been deposited in the Protein Data Bank, Research Collabora-
tory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ
(http://www.rcsb.org/).
1 Both authors contributed equally to this work.
2 To whom correspondence should be addressed: School of Life Sciences,
Peking University, Beijing, 100871 China. Tel.: 86-10-6275-5712; Fax: 86-
10-6276-5913; E-mail: xiaofengz@pku.edu.cn.
3 The abbreviations used are: BAR protein, Bin/amphiphysin/Rvs domain pro-
tein; PACSIN, protein kinase C and casein kinase substrate in neurons pro-
tein; PDB, Protein Data Bank.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 287, NO. 26, pp. 22387–22396, June 22, 2012
© 2012 by The American Society for Biochemistry and Molecular Biology, Inc.
Published in the U.S.A.
JUNE 22, 2012•VOLUME 287•NUMBER 26
JOURNAL OF BIOLOGICAL CHEMISTRY 22387
1 and 2 can induce a wide range radius of membrane tubules
including not only the typical 100-nm tubule but also smaller
tubules with an average diameter of 10 nm. In addition, they
also facilitate tubule constrictions (31). Both crystal structures
of the F-BAR domain of PACSIN 1 and 2 reveal a unique wedge
loop, which was proposed to contribute to membrane insertion
and binding (31). A hinge in the distal end of the murine PACSIN
2 F-BAR domain was predicted to contribute to membrane cur-
vature sensing (32). However, the detailed mechanism of how
the F-BAR dimers of PACSIN induce various curvature tubu-
lations remains unclear.
To elucidate further the mechanism of membrane remodel-
ing by the F-BAR domain of PACSINs, we investigated the
membrane deformation by the PACSIN 3 F-BAR domain com-
pared with PACSIN 1 and PACSIN 2 F-BAR domains, by neg-
ative-stain electron microscopy. We further determined the
structures of the F-BAR domains from PACSIN 3, PACSIN 1,
PACSIN 2, and two PACSIN 3 loop mutants, E128A and
P121Q. We found that different from PACSIN 1 and 2, PACSIN
3 could only induce 100-nm tubules. Based on structure stud-
ies, we discovered that the more rigid PACSIN 3 wedge loop,
which is in a different structural conformation compared with
PACSIN 1 and 2, contributes to the distinct lattice formed by
the PACSIN 3 F-BAR domain in the crystal. The parallel lattice
of PACSIN 3 reveals a novel type of wedge loop-mediated lat-
eral interaction between neighboring dimers, and the lattice is
related to protein packing on the tubulated membrane. We fur-
ther identified the residues that determine the rigidity of the
wedge loop in the PACSIN 3 F-BAR domain. The mutations of
these residues change the structural conformation of the wedge
loop, which alters the wedge loop-mediated lateral interactions
between neighboring dimers and further influences the mem-
brane curvature. Therefore, the rigidity of the wedge loop in
PACSIN 3 is a key factor in dictating the diameters of tubules.
EXPERIMENTAL PROCEDURES
Protein Expression and Purification—The F-BAR domain of
PACSIN 1 (residues 1–344), PACSIN 1 mutants, the F-BAR
domain of PACSIN 2 (residues 1–372) and PACSIN 2 mutant
E130A, the F-BAR domain of mouse PACSIN 3 (residues
1–341, which has 94% amino acid sequence identity to human
PACSIN 3) and PACSIN 3 mutants, were cloned into pET28a
vector. All proteins except PACSIN 1 truncate 1–344 were
expressed in Escherichia coli BL21 (DE3) cells and purified on a
Ni2-HiTrap affinity column followed by a Superdex-75 col-
umn (GE Healthcare) (33).
PACSIN 1 truncate 1–344 was expressed in B834 cells which
were cultured in M9 minimal medium supplemented with 50
mg/liter kanamycin and 40 mg/liter selenomethionine, and
purified as described (33).
Protein Crystallization and Structural Determination—Ini-
tial crystallization conditions were screened using kits from
Hampton Research including PEG, Crystal Screen, Crystal
Screen 2, and Index. Crystals were optimized by the hanging-
drop vapor diffusion method. Drops were prepared by mixing 2
l of protein solution (8 mg/ml protein in 500 mM NaCl, 10 mM
HEPES, pH 7.5) with 2 l of reservoir solution and were equil-
ibrated against 500 l of reservoir solution at 293 K. A large
crystal of PACSIN 1 F-BAR (600 190 200 m) was
obtained within 2 weeks from 200 mM NH4H2PO4, 100 mM
HEPES, pH 7.5, 16% PEG 3350 (w/v), and 5% glycerol. The
crystal of PACSIN 2 F-BAR was obtained from 100 mM MgCl2,
100 mM sodium cacodylate, pH 6.5, 18% PEG 3350 (w/v) within
2 days. The crystal of PACSIN 3 F-BAR was obtained from 200
mM KSCN, 100 mM HEPES, pH 7.3, 100 mM CaCl2, 20% PEG
3350 (w/v) within 3 days. The crystal of PACSIN 3 E128A was
obtained from 200 mM CH3COONH4, 100 mM CaCl2, 16% PEG
3350 (w/v), and PACSIN 3 P121Q from 0.5 M ammonium sul-
fate, 0.1 M sodium citrate tribasic dehydrate, pH 5.6, 1.0 M lith-
ium sulfate monohydrate within 10 days. 15, 15, 25, 20, and 28%
glycerol was used as cryoprotectant for PACSIN 1, PACSIN 2,
PACSIN 3, PACSIN 3 E128A, and PACSIN 3 P121Q.
For crystal data collection, crystals were flash-cooled with a
nitrogen stream at 100 K, and x-ray diffraction data were col-
lected on Mar 345 image plate detector at SSRF. Data of PACSIN
1(1–344) were processed with HKL2000 and MLPHARE soft-
ware (34). Images of the F-BAR domains of PACSIN 1, PACSIN
2, PACSIN 3, PACSIN 3 mutant E128A, and PACSIN 3 P121Q
were integrated with Mosflm (35), and data were carried out by
Molecular Replacement by CCP4 (34) with PACSIN 1–344 as
the model, and finally refined by COOT (36).
Liposome Preparation—Lipids containing 80% DOPC and
20% DOPA (Avanti) were mixed and dissolved in chloroform.
The organic solvent was removed by evaporation under a
stream of nitrogen gas, followed by incubation for 2 h in a vac-
uum to ensure complete removal of solvent. Lipid films were
resuspended in HEPES buffer (10 mM HEPES, pH 7.4, 50 mM
NaCl, 0.2 mM EDTA) and subjected to 10 freeze-thaw cycles.
Large unilamellar vesicles were then formed by extrusion
through 100-nm nucleopore polycarbonate membranes. The
prepared liposomes were stored at 4 °C.
Liposome Sedimentation Assay—Protein (1 mg/ml) was
incubated with liposome (1 mg/ml) with 1:1 protein-lipid vol-
ume ratio for 20 min at room temperature. The protein-lipid
mixture was centrifuged at 140,000 g for 30 min at 4 °C in an
ultracentrifuge (Beckman TLA100 rotor). Supernatants were
then collected, and pellets were resuspended in 40 l of sample
buffer. Proteins in both fractions were subjected to SDS-PAGE,
stained with Coomassie Blue, and visualized by a Bio-Rad XRS
system.
Tubulation Assays—Protein (1 mg/ml) was incubated with
liposome (1 mg/ml) with 1:1 protein-lipid volume ratio for 5
min at room temperature. 6-l protein-liposome samples were
then spread onto freshly glow-discharged Formvar- and car-
bon-coated electron microscopy grids, stained with 2% uranyl
acetate for 1 min, and air dried at room temperature. The grid
was examined on a transmission electron microscope (FEI 200
kV) with the electron energy set to 120 kV.
Nanogold Labeling Tubulation Assays—Protein (1 mg/ml)
was incubated with liposome (1 mg/ml) with 1:1 protein-lipid
volume ratio for 5 min at room temperature. 6-l protein-lipo-
some samples were then spread onto freshly glow-discharged
Formvar- and carbon-coated electron microscopy grids and
stained with 5 nm Ni-nitrilotriacetic acid-nanogold (Nano-
probes) for 1 min at room temperature followed by staining
with 2% uranyl acetate for 1 min and air dried at room temper-
Key Factor in Dictating Diameters of Tubules
22388
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 287•NUMBER 26•JUNE 22, 2012
ature. The grid was examined on a transmission electron
microscope (FEI 200 kV) with electron energy set to 120 kV.
RESULTS
Membrane Tubulation Induced by F-BAR Domain of PAC-
SIN 3 Is Different from PACSIN 1 and 2—To investigate the
membrane deformation induced by PACSINs, the F-BAR
domains from PACSIN 1, 2, and PACSIN 3 were expressed in
E. coli and purified to near homogeneity. Liposome tubulation
induced by F-BAR domains was examined by negative stain
electron microscopy. Unlike other F-BAR domain-containing
proteins, tubules induced by PACSIN 1 and 2 have various
diameters, which is consistent with previous reports (31). These
tubules are mainly classified into three classes: the first class has
low curvature with a diameter of 100 nm, the second class has
intermediate curvature with a diameter of 50 nm, and the
third class has high curvature with a diameter of 10 nm (Fig.
1A). In addition, tubules with other diameters were also
observed. In contrast, the PACSIN 3 F-BAR domain could only
induce low curvature tubules of 100-nm diameter (Fig. 1A),
despite sharing 55 and 60% sequence identity with the F-BAR
domain of PACSIN 1 and 2, respectively.
Crystal Structure of PACSIN 3 F-BAR Domain Shows Novel
Type of Lateral Interaction Mediated by Wedge Loop—To elu-
cidate the molecular mechanism to explain the differences in
membrane tubulation between PACSIN 3 and PACSIN 1, 2, we
crystallized and determined the structures of the F-BAR
domain of PACSIN 3 (residues 1–304), as well as that of PAC-
SIN 1 (residues 14–308) and PACSIN 2 (residues 1–304) (sup-
plemental Fig. 1 and Table 1). The structure of the PACSIN 1
F-BAR domain was determined by single anomalous dispersion
phasing, using data collected with a crystal of selenomethio-
nine-derivatized protein, and the structures of PACSIN 2 and
PACSIN 3 F-BAR domains were solved by molecular replace-
ment using the initial structure of the PACSIN 1 F-BAR domain
as the searching model. The structures of PACSIN 1, PACSIN
2, and PACSIN 3 F-BAR domains were refined to resolutions of
2.9, 2.7, and 2.6 Å in the space groups C2, P21, and P21, respec-
tively (Table 1). The structures showed that PACSINs form a
dimer in the crystal and adopt a crescent shape with a six-helix
bundle core and two three-helix bundle arms extending from
the central body. Each monomer is composed of six -helices
that interact with the other monomer in an antiparallel man-
ner. Two neighboring helices 1 form the main central
dimerization interface that are flanked by two wings composed
of helix 3 and the curved region of helix 4. Helix 3 and a
longer helix 4 both extend the length of the monomer with
bending ends. Helix 2 and the shorter helices 5 and 6 gen-
erate the bundle center that is associated with helix 1 (Fig. 1B
FIGURE 1. PACSIN-induced liposome tubulation. A, negative stain EM images of liposomes and tubules induced by PACSIN 3 and PACSIN 1, two F-BAR
domains, and quantification of liposome tubulation in the same experiment. Liposomes were incubated with the purified F-BAR domain of PACSIN 1 (residues
1–344), PACSIN 2 (residues 1–372), and PACSIN 3 (residues 1–341). Insets show the higher magnifications of the tubules indicated by arrows. Scale bars, 50, 100,
and 500 nm, respectively. B, ribbon presentation of the F-BAR dimers of PASCIN 3. The two monomers interact with each other in an antiparallel manner. Each
monomer consists of six -helices named 1–6. One monomer is tinted in a different color for the six -helices; the other is colored in green. C, superimposition
of the F-BAR domain between PACSIN 3 (Protein Data Bank (PDB) code 3QE6) and FBP17 (PDB code 2EFL). Ribbon representation is shown. PACSIN 3 and FBP17
are colored in cyan and yellow, respectively. The arrow indicates the unique wedge loop in the PACSIN family proteins that is not found in any other F-BAR
domain structure.
Key Factor in Dictating Diameters of Tubules
JUNE 22, 2012•VOLUME 287•NUMBER 26
JOURNAL OF BIOLOGICAL CHEMISTRY 22389
and supplemental Fig. 1A). The structure-based sequence
alignment of the residues on the protein surface of the F-BAR
domains of PACSINs showed that the positively charged resi-
dues on the concave surface are highly conserved (supplemen-
tal Fig. 1, B and C). Consistent with the previously determined
structures of the F-BAR domain of PACSIN 1 and 2 (31), the
wedge loop between helices 2 and 3 is also present in PAC-
SIN 3, which is a distinct feature of the F-BAR domain of PAC-
SINs and has not been observed in other BAR domain-contain-
ing families (Fig. 1C).
Despite the structure similarity between the F-BAR dimer of
PACSIN 3 and those of PACSIN 1 and 2, we found that the
packing patterns of the neighboring dimers of these proteins
are strikingly different in the crystal: PACSIN 3 shows only a
parallel packing among all the different crystallization condi-
tions that have been examined, whereas PACSIN 1 and 2 show
different packing modes (supplemental Fig. 2).
We also found that the parallel packing of the PACSIN 3
F-BAR domain is similar to the packing lattice of the CIP4
F-BAR domain observed on the tubules by cryo-EM (18). How-
ever, a close-up view of the PACSIN 3 structure reveals a dis-
tinct type of lateral interaction between two neighboring
dimers (Fig. 2A), in addition to the typical CIP4-like tip-to-tip
interaction. In CIP4, the lateral interactions between the neigh-
boring dimers involve only the residues of neighboring helices,
whereas in PACSIN 3, the lateral interactions are between one
of the PACSIN-specific wedge loops with the helix 2 of the
neighboring dimer (Fig. 2, A, left, and B, upper). The other
wedge loop of the same dimer extends out into the membrane
direction (Fig. 2, A, right, and B, lower). This observation sug-
gests that the wedge loop might function in two ways: one loop
in the dimer is involved in the lateral interaction, and the other
loop in the dimer extends out and inserts into the membrane
bilayer.
Dual Roles of Wedge Loop in Mediating Protein Packing and
Membrane Binding—To confirm that the wedge loop is
involved in the interactions between the neighboring dimers,
we mutated the residues located in the lateral contacting area
(Fig. 2C, left). A total of 15 mutants of the five residues on the
contacting surface were constructed. The mutants were puri-
fied and examined by a liposome tubulation assay (Fig. 3A). The
binding of these mutants to liposomes was investigated by a
sedimentation assay (Fig. 3B). Mutations in the wedge loop
(H119E/A/Q, R127E/A/Q) or in the contacting area (E93R/
A/Q, E97R/A/Q and E100R/A/Q) abolished the liposome tubu-
lation activity completely, but retained the liposome binding
ability (Fig. 3 and supplemental Fig. 3B). This observation indi-
cates that the interaction between the wedge loop and its neigh-
boring dimer plays critical roles in protein packing on the mem-
brane to induce liposome tubulation.
We further examined the residues on the wedge loop that
may insert into the membrane bilayer. A series of mutants of
the wedge loop, V122L123, V122E/L123E, and V122R/L123R
of PACSIN 3 (Fig. 2C, right) were constructed in which the two
bulky hydrophobic residues at the base of the wedge loop were
either deleted or mutated. The liposome tubulation activities
and the binding of these mutants to liposomes were also inves-
tigated. Our results showed that the deletion of the wedge loop
(V122L123) or mutation of residues Val-122 and Leu-123 at
the center of the wedge loop to Glu or Arg (V122E/L123E,
V122R/L123R) totally abolished induction of tubules and
reduced liposome binding (Fig. 3B), which is consistent with
the observation in the site mutation constructs I125E or
M126E previously reported in PACSIN 1 (31). These results
indicate that the wedge loop binds liposomes through hydro-
phobic residues Val-122 and Leu-123 (or Ile-125/Met-126 in
PACSIN 1, and Met-124/Met-125 in PACSIN 2, supplemen-
tal Fig. 3B).
TABLE 1
Data collection and refinement statistics
Se-PACSIN
(residues 1–307)
PACSIN 2
(residues 1–304)
PACSIN 3
(residues 1–304)
PACSIN3E128A
(residues 1–302)
PACSIN3P121Q
(residues 6–299)
Diffraction data
Space group
C2
P21
P21
P21
P21
Unit cell parameters
(a, b, c) (Å)
85.3, 154.3, 215.4
31.58, 86.13, 353.80
46.9, 54.7, 193.7
47.5, 52.3, 196.5
120.570, 108.901, 222.319
90, 90.3, 90
90, 90, 90
90, 96.9, 90
90, 94.8, 90
90.00, 90.05, 90.00
Resolution range (Å)
30–2.8 (2.91–2.8)
30–2.6 (2.75–2.6)
30–2.6 (2.67–2.6)
15–2.6 (2.67–2.6)
50–3.1 (3.18–3.1)
Unique reflections
63,353
41,646
28,175
28,283
98,202
Completeness (%)
96.2 (87.43)
99.7 (85.9)
97.8 (90.8)
99.8 (93.7)
98.68 (98.02)
Mean I/(I)
16.65 (2.5)
14.5 (2.5)
35.4 (3.4)
14.2 (2.4)
18.3 (3.1)
Multiplicity
4.3
3.1
4.1
4
4.3
Refinement
Rwork/Rfree (%)a
22.1/29.4
20.77/29.2
22.7/27.4
23.2/28.1
27/33.6
Modeled chain: residues A
Non-H atoms (protein/water)
13,874/213
9,683//202
4,648/34
4,587/10
16,535/197
Root mean square deviations
Bonds (Å)/Angles (°)
0.017/1.692
0.015/1.593
0.022/2.041
0.011/1.297
0.0213/2.009
Ramachandran analysis
Favored
98.61
98.14
93.94
96.22
94.01
Allowed
1.33
1.68
5.81
3.78
5.14
Generously allowed
0.06
0.18
0.24
0
0.86
Disallowed
0
0
0
0
0
a R Fo Fc /Fo. The full-length PACSINs proteins were first crystallized but degraded from the C-terminal Src homology 3 domain. Then the truncation of Se-
PACSIN (residues 1–305), PACSIN 2 (residues 1–305), and PACSIN 3 (residues 1–302) was designed and crystallized.
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There is a possibility that the mutant proteins may be dis-
torted in structure, thus influencing the membrane tubulation
property. We therefore monitored the secondary structure of
all of the mutant proteins by a circular dichroism assay. Our
results showed that the secondary structures of these mutants
are similar to the wild-type protein (supplemental Fig. 3), indi-
cating that the lost of the tubulation activity is caused by the
elimination of protein-protein interaction, not by destroying
the structure of the protein.
To characterize further where the PACSIN proteins localize
in the tubulation process, 5-nm Ni2-nitrilotriacetic acid-
linked nanogold particles were used to label the His-tagged
PACSIN F-BAR proteins in tubulation assays (Fig. 3C). The
results showed that the nanogold particles were concentrated
on the surface of the tubules. This observation indicates that
tubules are induced by the PACSIN F-BAR proteins.
Rigidity of Wedge Loop Influences Lateral Interactions and
Dictates Diameter of Tubules—Superimposition of the wedge
loops among PACSIN 3 and PACSIN 1 and 2 revealed that the
wedge loop of PACSIN 3 has a different conformation. The
wedge loop of PACSIN 3 is wider and bends more toward
the inner side of the dimer compared with the relatively flat,
outward-splayed loops in PACSIN 1 and 2 (Fig. 4A, left). We
therefore hypothesized that the conformational difference of the
wedge loops would be an important factor in determination of the
interaction pattern between neighboring dimers and would con-
tribute further to the variations in membrane tubulation.
Structure-based sequence alignments of the wedge loop
region in PACSINs revealed that the predominant difference
between PACSIN 3 and PACSIN 1 or 2 is residue Pro-121
located in the center of the wedge loop of PACSIN 3, which
superimposes onto residue Gln-124 and Gln-123 in PACSIN 1
and 2, respectively (Fig. 4, B and C). The rigidity of the proline
residue may restrain the flexibility of the wedge loop in PACSIN
3 and affect the interaction pattern between neighboring
dimers and membrane tubulation.
To verify the above predictions, we mutated Pro-121 of
PACSIN 3 to glutamine, which is found in PACSIN 1 and 2, and
substituted the equivalent glutamine of PACSIN 1 and 2 with
proline. The effects of the mutant proteins on liposome tubu-
lation were investigated by negative stain EM. Consistent with
the hypothesis, replacement of proline with glutamine at posi-
tion 121 (P121Q) in PACSIN 3 led to the formation of a large
number of pseudopod-like small tubules, in addition to the
large tubules (Fig. 5A). These pseudopod-like tubules were
approximately 10 nm in diameter, and some are budded from
the surface of the large tubules. At the same time, replacement
of Gln with Pro in PACSIN 1 Q124P and PACSIN 2 Q123P
induced only one type of large tubules that are 100-nm in
diameter (Fig. 5B). Moreover, we determined the crystal struc-
ture of the PACSIN 3 P121Q mutant to 2.9 Å resolution (sup-
plemental Fig. 4 and Table 1). The structure revealed that the
mutation of proline to glutamine changed the orientation of the
wedge loop of PACSIN 3, and the wedge loop in the mutant
structure points outward and superimposes very well with the
wedge loop of native PACSIN 1 and 2 (Fig. 4A, middle). In
addition, the crystal packing of PACSIN 3 P121Q showed an
obvious change of the angle between the neighboring dimers,
which resembles that found in PACSIN 1 and 2 packing lattices
(supplemental Fig. 2). These results demonstrate that the rigid-
ity of the wedge loop in the PACSINs F-BAR domain correlates
with the conformational change of the wedge loop and further
FIGURE 2. Wedge loop plays dual roles in PACSIN 3 packing. A, novel type of loop-mediated lateral interaction found in PACSIN 3 F-BAR packing. The PACSIN
3 F-BAR domain packs in a parallel manner which connected by tip-to-tip interactions and the loop-mediated lateral interactions. B, close-up view of the two
wedgeloopsinthesamedimer.Oneofthewedgeloopsinteractswithitsadjacentdimer(upper).Theotherwedgeloopisexposedandextrudesintothetarget
membrane direction (lower). The right view is rotated by 90° clockwise relative to the left view. The two adjacent dimers are colored marine and yellow, and
presented as ribbon and surface, respectively. C, left image showing residues on the contacting surface between the wedge loop and the neighboring dimer.
Indicated residues are shown as side chain sticks. Right image shows the membrane binding wedge loop from residue His-119 to Glu-128 and is presented as
side chain sticks.
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affects lateral interactions of neighboring dimers and mem-
brane tubulation.
To investigate whether mutation of Pro-121 in PACSIN 3 to
glutamine affects the specific interactions between the wedge
loop and membrane, a sedimentation assay was performed. The
result showed that PACSIN 3 P121Q did not change the bind-
ing of protein to lipid bilayers (supplemental Fig. 3B). Con-
versely, the mutations of PACSIN3 Val-122 and Leu-123 to
charged residues, or deletion of them, resulted in the complete
loss of the membrane binding ability even though Pro-121 is
still intact (Fig. 3). These results indicate that Pro-121 is not
involved in membrane insertion. Therefore, the reason that
P121Q can induce the large type of tubules is not due to the
change of the wedge loop insertion.
Structure-based sequence alignments also suggest that there
is another residue, Glu-128 in PACSIN 3 (Glu-131 in PACSIN 1
FIGURE 3. Tubulation and sedimentation assays of PACSIN 3 F-BAR and mutants. A, negative-stained electron micrographs are shown. Liposomes were
incubated with PACSIN 3 and its mutant proteins and examined by EM. V122L123, V122E/L123E, V122R/L123R altered only membrane binding, whereas
R127E, H119E altered only the interactions between neighboring dimers. E93R, E97R, and E100R are mutations on the contacting surface of neighboring dimer
as shown in Fig. 2C, left. B, liposomes were incubated with PACSIN 3 and its mutant proteins and examined by sedimentation assays. Supernatant (S) and pellet
(P) fractions were analyzed by SDS-PAGE. Band in pellet represents the protein bound to liposome. Histograms with means S.E. (error bars) show the
quantified protein in supernatant and pellet with the total protein defined as 100%. C, liposomes were incubated with the His-tagged F-BAR proteins of
PACSINs followed by Ni-nitrilotriacetic acid-nanogold particles (5-nm diameter) treatment and then visualized by EM.
FIGURE 4. Comparisons of the wedge loop among PACSINs. A, structural alignments of the wedge loop among PACSIN 1, 2, 3 and PACSIN 3 mutants P121Q
and E128A. The wedge loop in PACSIN 3 points inward by 45° compared with PACSIN 2 and points inward by 30° compared with PACSIN 1 (left). Compared with
the wedge loop in the wild-type PACSIN 3 F-BAR domain, the wedge loop in the PACSIN 3 P121Q mutant swings outward and is superimposable with the
wedge loop of PACSIN 1 and 2 (middle); and the wedge loop in the PACSIN3 E128A mutant reoriented significantly by swinging 30° outward to resemble the
outward-splayed wedge loop in PACSIN 1 and PACSIN 2 (right). B, close-up view of the wedge loops of PACSIN 1, PACSIN 2, PACSIN 3, and PACSIN 3 E128A
mutant. All of the native structures of the PACSIN F-BAR domains include a conserved hydrogen bond between Glu and His at the base of the wedge loop
(Glu-131 and His-122 for PACSIN 1, Glu-130 and His-121 for PACSIN 2, and Glu-128 and His-119 for PACSIN 3), whereas this hydrogen bond is absent in the
PACSIN 3 E128A mutant. C, sequence alignments of the wedge loop among PACSINs from human and mouse using ClustalX. Instead of the conserved residue
Gln in PASCIN 1 and 2, it is a conformationally restrained Pro residue at position 121 in PACSIN 3. Asterisks and colons indicate homology sequence among
PACSIN 1 PACSIN 2 and PACSIN 3.
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JUNE 22, 2012•VOLUME 287•NUMBER 26
JOURNAL OF BIOLOGICAL CHEMISTRY 22393
and Glu-130 in PACSIN 2) that forms a hydrogen bond with the
mainchainofHis-119(His-122inPACSIN1andHis-121inPAC-
SIN2)(Fig.4B),mayalsobeimportantfortheconformationofthe
wedge loop. The crystal structure of the PACSIN 3 E128A mutant
determined here to 2.8 Å resolution (supplemental Fig. 4 and
Table 1) showed that substitution of Glu-128 with alanine elimi-
nated the hydrogen bond between Glu-128 and His-119 (Fig. 4B)
andresultedina significant conformational change in the wedge
loop. The E128A wedge loop of PACSIN 3 was found to resem-
ble the wedge loop of PACSIN 1 and 2 in an outward-splayed
pattern (Fig. 4A, right). Structural comparisons of the PACSIN
3 E128A loop with that of the wild type suggest that the hydro-
gen bond between Glu-128 and His-119 could function as a lock
to maintain the loop conformation. The PACSIN 3 E128A was
found to induce predominantly tubules with a diameter of 50
nm in addition to the typical 100-nm tubules induced by wild-
type PACSIN3 (Fig. 5C), which resemble the intermediate and
low curvature tubules induced by PACSIN 1 and 2. Similarly,
mutants E131A and E130A of PACSIN 1 and 2 also showed
more membrane constrictions budding from the tubules (sup-
plemental Fig. 5) and more high curvature tubules compared
with the wild-type PACSIN 1 and 2. These data suggest that, in
PACSINs, the hydrogen bond between Glu-128 (Glu-131 or
Glu-130) and His-119 (His-122 or His-121) helps to reinforce
the rigidity and orientation of the wedge loop and therefore
affects the curvature of the tubules.
DISCUSSION
Membrane curving by BAR domain proteins is an important
biological process involved in vesicle trafficking and subcellular
structure stabilization. Various tubules induced by the F-BAR
domain of PACSINs are important in membrane deformation
(20–25). It was suggested that the distinct wedge loop of PACSIN
1 and 2 F-BAR domains inserts into lipid bilayers of membrane
(31). However, there is no clear interpretation to explain why the
F-BAR domains of PACSIN 1 and 2 induce tubules with such
diverse diameters.
Here, we performed structural and biochemical studies on
the F-BAR domains from all three PACSINs and systematically
compared the differences of the F-BAR domains between PAC-
SIN 3 and 1, or 2 with respect to structure and membrane tubu-
lation activities. Our analyses revealed that the more rigid wedge
loop in the F-BAR domain of PACSIN 3 is responsible for its
unique feature of inducing only low curvature tubules compared
FIGURE 5. Key residues in determination of diameter of liposome tubules. Liposomes were incubated with different mutant proteins and examined by EM.
The numbers of the tubules were quantified on five independently prepared EM grids. Membrane constrictions are denoted by arrows. A, mutation in the
wedge loop of PACSIN 3 F-BAR P121Q has a profound influence on the diameters of the induced liposome tubules. B, substitution of residue Gln to Pro in the
wedge loop of PACSIN 1 and 2 only results in low curvature tubules as shown in PACSIN 3 F-BAR in Fig. 1C. C, mutation in the wedge loop of PACSIN 3 E128A
induces predominantly tubules with a diameter of 50 nm, in addition to the typical 100-nm tubules as induced by PACSIN 3 F-BAR.
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withPACSIN1and2.WhenthewedgeloopintheF-BARdomain
of PACSIN 1 and 2 was mutated to the more rigid structure as
found in PACSIN 3, these mutants induced only low curvature
tubules as the wild-type PACSIN 3. Conversely, mutants that
modified the PACSIN 3 wedge loop to resemble the wedge loop in
PACSIN 1 and 2 induced tubules with dimensions similar to those
found in the wild-type PACSIN 1 and 2.
The distal end of murine PACSIN 2 was proposed to sense the
membrane (32). Comparisons of the distal ends of wild-type PAC-
SIN 3 with PACSIN 1 and 2 show no obvious difference in all the
structures we determined (supplemental Fig. 6), suggesting that
the various phenotypes presented by PACSIN 1 and 2 F-BAR may
not be correlated with the intrinsic curvature of the dimer surface.
It was previously reported that during tubulation, F-BAR
proteins make tip-to-tip interactions and contacts between lat-
erally adjacent dimers (18). The diameters of the induced
tubules were proposed to be related to the intrinsic large radial
curvature of the F-BAR domain, such as CIP4 and FBP17 (16).
However, in PACSINs, the intrinsic curvature itself certainly
could not result in such diverse tubules with various diameters
because PACSIN 1 and 2 induce tubules different from those
induced by PACSIN 3 even though all three PACSINs almost
have the same intrinsic curvature. According to the crystal
packing pattern of PACSIN 3, we propose a potential mem-
brane tubulation model for PACSINs (supplemental Fig. 7): the
F-BAR domain is connected by tip-to-tip interactions and the
wedge loop-mediated lateral interactions to form filaments.
The filament of the F-BAR protein bends as a hinge motion at
the tip-to-tip interaction to various extents. The extent of the
motion is dependent on the angle between two adjacent dimers,
and this angle is determined by the conformation of the wedge
loop. This results in the filament winding spirally around a
cylindrical membrane. The larger the angle is between the two
dimers, the more the dimer bends the membrane; the more the
membrane is bent, and the smaller the diameter of the tubule is
(supplemental Fig. 7E). This model also shed light on under-
standing why PACSINs generate different size tubules in so
many biological processes, such as trans-Golgi network vesicle
formation, filopodia tips, and lamellipodia dynamics, micro-
spike formation, and caveola fission (22, 24, 37, 38).
In conclusion, the membrane tubulation by PACSIN 3 was
shown to be different from PACSIN 1 and 2, and these differ-
ences are due to different degrees of rigidity of the wedge loop.
We demonstrated that the rigidity of the wedge loop in the
PACSIN F-BAR domain is a key factor that determines the dif-
ferent angles between two neighboring dimers and dictates the
diameters of various tubules. Our study provides new insights
for understanding the mechanism of membrane deformation
by the PACSIN family proteins.
Acknowledgments—We thank Prof. Fuyu Yang and Dr. Kai Zhao
from Institute of Biophysics, Chinese Academy of Sciences, for lipid
preparation; Dr. Ning Gao at Tsinghua University for the nanogold
labeling assay; and Dr. Plomann at Stanford University School of
Medicine for providing the PACSIN 3 plasmid. X-ray diffraction data
collection was carried out at the Beijing Synchrotron Radiation Lab-
oratory and Shanghai Synchrotron Radiation Facility.
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|
3M3Y
|
RNA polymerase II elongation complex C
|
X-ray structure and mechanism of RNA polymerase II
stalled at an antineoplastic monofunctional
platinum-DNA adduct
Dong Wanga,b,1, Guangyu Zhuc, Xuhui Huangd, and Stephen J. Lippardc,1
aDepartment of Structural Biology, Stanford University School of Medicine, Stanford, CA 94305; bSkaggs School of Pharmacy and Pharmaceutical Sciences,
University of California, San Diego, La Jolla, CA 92093; cDepartment of Chemistry, Massachusetts Institute of Technology, Cambridge, MA 02139; and
dDepartment of Chemistry, Hong Kong University of Science and Technology, Clear Water Bay, Kowloon, Hong Kong, P.R. China
Contributed by Stephen J. Lippard, March 3, 2010 (sent for review February 9, 2010)
DNA is a major target of anticancer drugs. The resulting adducts
interfere with key cellular processes, such as transcription, to
trigger downstream events responsible for drug activity. cis-
Diammine(pyridine)chloroplatinum(II), cDPCP or pyriplatin, is a
monofunctional platinum(II) analogue of the widely used antican-
cer drug cisplatin having significant anticancer properties with a
different spectrum of activity. Its novel structure-activity properties
hold promise for overcoming drug resistance and improving the
spectrum of treatable cancers over those responsive to cisplatin.
However, the detailed molecular mechanism by which cells process
DNA modified by pyriplatin and related monofunctional complexes
is not at all understood. Here we report the structure of a transcri-
bing RNA polymerase II (pol II) complex stalled at a site-specific
monofunctional pyriplatin-DNA adduct in the active site. The re-
sults reveal a molecular mechanism of pol II transcription inhibition
and drug action that is dramatically different from transcription in-
hibition by cisplatin and UV-induced 1,2-intrastrand cross-links. Our
findings provide insight into structure-activity relationships that
may apply to the entire family of monofunctional DNA-damaging
agents and pave the way for rational improvement of monofunc-
tional platinum anticancer drugs.
anticancer ∣chemotherapy ∣DNA damage ∣pyriplatin ∣transcription
T
he DNA template for transcription is not only the site of in-
born errors of metabolism and of continuous attack by harm-
ful environmental agents, but it also represents a major target
for cancer therapy. Platinum-based anticancer drugs such as
cisplatin, cis-diamminedichloroplatinum(II), are widely used
and among the most effective antineoplastic treatments (1, 2).
Platinum-based drugs typically form bifunctional intra- or inter-
strand DNA cross-links by covalent bonding to the N7 positions of
two guanosine residues, triggering a variety of cellular processes,
including
transcription
inhibition
with
attendant
apoptosis
(1, 2). However, resistance and side effects can require with-
drawal of these drugs before they can effect a cure in certain types
of cancer (3).
In the effort to find new compounds that circumvent resis-
tance to conventional bifunctional platinum-based drugs, a class
of monofunctional platinum compounds were synthesized and
screened for anticancer activity (4–6). In contrast to other
inactive
monofunctional
platinum(II)
compounds
such
as
½PtðdienÞClþ and ½PtðNH3Þ3Clþ, cis-diammine(pyridine)chloro-
platinum(II) [cDPCP or “pyriplatin” (Fig. 1)] and related com-
plexes display significant anticancer properties and a different
spectrum of activity compared to conventional platinum-based
drugs. These features render them attractive candidates for treat-
ing cisplatin-refractory patients if the potency could be raised to
or beyond the level of that of cisplatin (4, 5, 7). Pyriplatin exhibits
unique chemical and biological properties, forming monofunc-
tional DNA adducts (Fig. 1 and Fig. S1) that can inhibit transcrip-
tion and better elude DNA repair (7). The x-ray crystal structure
of pyriplatin bound to a DNA duplex reveals substantially
different features than those of DNA adducts formed by conven-
tional, bifunctional platinum-based drugs. The overall DNA
duplex is much less distorted, with the pyridine ligand of the
cis-fPtðNH3Þ2ðpyÞg2þ moiety directed toward the 50-end of the
platinated strand. A hydrogen bond forms between the NH3
ligand trans to pyridine and O6 of the platinated guanosine
residue (7).
The detailed molecular mechanism by which cells process
DNA modified by monofunctional complexes such as pyriplatin
is not understood. Several important questions remain unan-
swered. By what process do monofunctional adducts block pol
II transcription? Does the mechanism differ from that of tran-
scription inhibition by 1,2- and 1,3-intrastrand cross-links that
comprise the major adducts of cisplatin? Why do pyriplatin
and its homologues, which violate the classical structure-activity
relationships (SARs) for active, bifunctional platinum drugs (8),
show such promise by comparison to related monofunctional
complexes like ½PtðNH3Þ3Clþ? Would knowledge of the struc-
ture of pyriplatin-modified DNA at its site(s) of biological action
inform the design of more potent analogues?
In the present work we take a combined biochemical and x-ray
structural approach to investigate the molecular mechanism of
pol II transcription inhibition by a site-specific monofunctional
platinum(II)-DNA adduct of pyriplatin. An unprecedented mo-
lecular mechanism for pol II transcription inhibition is revealed,
providing insight into structure-activity relationships that may ap-
ply to the entire family of monofunctional DNA-damaging
agents, whether or not they contain platinum.
Results
A Different Configuration of a Pyriplatin-DNA Adduct Accommodated
in the Pol II Active Site. To understand how a monofunctional
pyriplatin-DNA adduct is accommodated in the active site of
the transcribing pol II elongation complex, we designed and pre-
pared a DNA template containing a site-specific DNA lesion of
this complex, as described previously (7). A transcribing pol II
complex was then assembled in which the pyriplatin-DNA lesion
occupies the active (þ1) site (Complex B, Table 1). The crystal
structure of this complex reveals that the platinated nucleotide
is captured as a pol II complex in the post-translocation state,
in which the addition site is empty and ready for NTP loading
(Dashed Ring, Fig. 2A and Fig. S2). Fig. 2A reveals that the
Author contributions: D.W. and S.J.L. designed research; D.W., G.Z., and X.H. performed
research; D.W., G.Z., X.H., and S.J.L. analyzed data; and D.W., X.H., and S.J.L. wrote
the paper.
The authors declare no conflict of interest.
Data deposition: The atomic coordinates have been deposited in the Protein Data Bank,
www.pdb.org (PDB ID codes 3M4O and 3M3Y).
1To whom correspondence may be addressed. E-mail: dongwang@ucsd.edu or lippard@
mit.edu.
This article contains supporting information online at www.pnas.org/cgi/content/full/
1002565107/DCSupplemental.
9584–9589 ∣PNAS ∣May 25, 2010 ∣vol. 107 ∣no. 21
www.pnas.org/cgi/doi/10.1073/pnas.1002565107
positioning of the pyriplatin-damaged guanosine residue is lo-
cated above the bridge helix. This structure requires rotation
of the cis-fPtðNH3Þ2ðpyÞg2þ moiety and its bound guanosine re-
sidue into a different configuration compared to that adopted in
the pyriplatin-duplex DNA structure, in order to avoid a steric
clash with bridge helix (7). Fig. 2B depicts this comparison.
The rotation is energetically facilitated by the formation of hydro-
gen bonds between the ammine ligands on platinum with the
phosphodiester moiety of the backbone between positions þ1
and þ2, with concomitant loss of a hydrogen bond between O6
of the platinated guanosine residue and an ammine ligand. An
additional feature is that the pyridine group of the cis-
fPtðNH3Þ2ðpyÞg2þ fragment, which points downstream toward
the 50-direction of the template DNA, forms van der Waals inter-
actions with bridge helix residues Val 829 and Ala 832. The purine
base of the guanosine residue at position þ1 is displaced toward
the major groove of the RNA–DNA duplex by comparison with
structures having an undamaged base at this site in the post-trans-
location state (9–11).
Transcription Elongation Inhibited by a Pyriplatin–DNA Adduct. Be-
cause transcription inhibition is an important component in
the mechanism of action of platinum anticancer drugs (12–20),
we investigated the effect of a site-specific pyriplatin–DNA ad-
duct on the kinetics of pol II transcription elongation. We per-
formed an extension assay using platinated (Complex A,
Table 1) and unplatinated (Complex A0, control, Table 1) pol
II transcribing complexes having a 9mer RNA as primer. These
complexes were then incubated with a mixture of ATP, CTP, and
GTP. The RNA transcripts in A could be elongated from the 9mer
to the 11mer, stopping at a position corresponding to the Pt–
DNA lesion site observed in the pol II complex of the damaged
template DNA, whereas RNA transcripts in A0 were extended
much farther downstream on the undamaged template control
DNA (Fig. 3A). In order to avoid the possibility of misincorpora-
tion-induced transcription inhibition in this assay, we carried out
a similar extension assay using an RNA containing a 30-end CMP
matched against the damaged base (pol II complex C, 11mer)
(Table 1). A single matching GTP was incubated with this pol
II complex to test whether the enzyme could bypass the Pt–
DNA lesion. Consistent with the results of the previous assay,
RNA transcripts could not be extended beyond an 11mer in
the pol II complex with the damaged DNA template, whereas
RNA transcripts were efficiently extended farther downstream
along the undamaged DNA template (Fig. 3B). Similar extension
assay results were obtained using a chain-terminated GTP analo-
gue 30-dGTP or an RNA primer of different length (complex B,
10mer) (Table 1) (Fig. 3 C and D). Finally, to investigate whether
the presence of the damaged base affects the rate of NTP incor-
poration in a single round, we used complex B (10mer) and com-
plex C (11mer), incubating with CTP and 30-dGTP, respectively.
For CTP incorporation, RNA transcripts could be efficiently ex-
tended from the 10mer to the 11mer using both damaged and
nondamaged templates at a comparable rate (Fig. 3E), whereas
no obvious extension of RNA transcripts from the 11mer to a
12mer was observed on the damaged DNA template (Fig. 3C).
UTP failed to incorporate at the damaged template under the
same conditions (Fig. S3A). No obvious intrinsic cleavage was
observed for a pol II complex containing the 11mer RNA and
Pt-damaged DNA template in the presence of 20 mM Mg2þ
ion, suggesting that most of complex C (11mer) is not in the back-
tracked state (Fig. S3B) (21–23).
X-ray Structure of Pol II stalled at a Pyriplatin–DNA Adduct. To under-
stand the nature of the pol II complex stalled at the pyriplatin-
induced Pt–DNA adduct, we solved the x-ray crystal structure
Fig. 1.
Scheme depicting the formation of a monofunctional platinum-DNA
adduct by pyriplatin on double-stranded duplex DNA. The structure of the
pyriplatin-damaged DNA duplex used coordinates from the PDB (code
3CO3). The damaged and nondamaged DNA strands are shown in cyan
and green, respectively. The pyridine ligand and two ammine groups of
the cis-fPtðNH3Þ2ðpyÞg2þ moiety are depicted in magenta and blue, respec-
tively. The platinum atom and nitrogen atoms of the cis-fPtðNH3Þ2ðpyÞg2þ
moiety are highlighted in yellow and as a blue ball, respectively. The termini
of the DNA strands are labeled.
A
B
+1
-1
+1
-1
5’
3’
3’
5’
5’
3’
Non-template
DNA
Bridge
Helix
Bridge
Helix
Addition
Site
Addition
Site
5’
V829
A832
RNA
RNA
Template
DNA
Template
DNA
3’
Fig. 2.
Structure of a pol II transcribing complex encountering a site-specific
pyriplatin-dG adduct in DNA. (A) A site-specific pyriplatin-DNA adduct is ac-
commodated in the pol II active site. The view is a standard one, from the
“Rpb2 side,” as described elsewhere (9–11, 39). The RNA transcript, template
DNA strand, and nontemplate DNA strand are depicted in red, cyan, and
green, respectively. Parts of the bridge helix (Rpb1 825–848) are shown in
gray. The pyriplatin-damaged guanosine is colored magenta. The platinum
atom of the cis-fPtðNH3Þ2ðpyÞg2þ moiety is denoted as a yellow ball and
the two ammine groups are in blue. The dashed oval represents the empty
nucleotide addition site in the post-translocation state. The positions of the
RNA strand are labeled. (B) cis-fPtðNH3Þ2ðpyÞg2þ-dG in the pol II active site
adopts a different configuration in comparison with its conformation in
the
structure
of
pyriplatin-modified
duplex
DNA.
The
superimposed
geometry of the cis-fPtðNH3Þ2ðpyÞg2þ-guanosine unit from the DNA duplex
structure (3CO3) is shown in light blue. Side chains of Val 829 and Ala 832 are
depicted in orange. The remainder of the figure is the same as in A.
Wang et al.
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of the enzyme in complex with a platinated DNA using an RNA-
containing CTP matched against the damaged guanosine residue.
In this structure, pol II is in pre-translocation state, with the newly
added CMP still occupying the addition site without transloca-
tion. The platinated guanosine residue forms Watson–Crick base
pairs with the newly added CMP (Fig. 4 A and B and Fig. S4). The
cis-fPtðNH3Þ2ðpyÞg2þ moiety is surrounded by the bridge helix at
the bottom, part of the Rpb2 fork region (528–534) on the left
side, and the sugar-phosphate backbone connecting template
DNA positions þ1 and þ2 on the right side (Fig. 4B). Interest-
ingly, upon CMP incorporation, the cis-fPtðNH3Þ2ðpyÞg2þ moiety
adopts a different conformation. The pyridine group of this unit
now faces toward 30-direction of template DNA (Fig. 4 A and B).
The ammine group trans to pyridine is directed toward the bridge
helix and forms hydrogen bonds with main chain atoms of Ala 828
and the side chain of Thr 831 (Fig. 4B). The residues in the bridge
helix are highly conserved from yeast to humans. Because Thr 831
and Ala 828 are absolutely conserved between S. cerevisiae and
humans, the interactions we observe in the S. cerevisiae pol II
structure will also occur in human pol II.
These structural results provide important insights into the
transcription stalling process at a monofunctional pyriplatin–
DNA adduct. The adduct adopts a significantly different confor-
mation within the pol II active site compared to that in duplex
DNA (7). The present structural and biochemical evidence
reveals that pol II stalls after efficient incorporation of CTP
against the damaged guanosine residue. The conformation of
the pyriplatin–DNA adduct changes significantly upon incorpora-
tion of CTP. The modified guanosine rotates into the pol II active
site and serves as a template for base pairing with the matched
substrate, and the cis-fPtðNH3Þ2ðpyÞg2þ moiety is now directed
toward 30-end of the platinated DNA.
Table 1. RNA and DNA scaffold of pol II transcribing complexes
Complex A: (Damaged template 29mer with 9mer RNA)
RNA: 5′
AUGGAGAGG
3′
DNA: 3′ CTACCTCTCCTG*CCCCACCAATACCCATC 5′
DNA: 5′
GTGGTTATGGGTAG 3′
Complex A′: (Nondamaged template 29mer with 9mer RNA)
RNA: 5′
AUGGAGAGG
3′
DNA: 3′ CTACCTCTCCTGCCCCACCAATACCCATC
5′
DNA: 5′
GTGGTTATGGGTAG
3′
Complex B: (Damaged template 29mer with 10mer RNA)
RNA: 5′
AUGGAGAGGA 3′
DNA: 3′ CTACCTCTCCTG*CCCCACCAATACCCATC 5′
DNA: 5′
GTGGTTATGGGTAG 3′
Complex B′: (Nondamaged template 29mer with 10mer RNA)
RNA: 5′
AUGGAGAGGA 3′
DNA: 3′ CTACCTCTCCTGCCCCACCAATACCCATC
5′
DNA: 5′
GTGGTTATGGGTAG
3′
Complex C: (Damaged template 29mer with 11mer RNA)
RNA: 5′
AUGGAGAGGAC3′
DNA: 3′ CTACCTCTCCTG*CCCCACCAATACCCATC 5′
DNA: 5′
GTGGTTATGGGTAG 3′
Complex C′: (Non-damaged Template 29mer with 11mer RNA)
RNA: 5′
AUGGAGAGGAC3′
DNA: 3′ CTACCTCTCCTGCCCCACCAATACCCATC
5′
DNA: 5′
GTGGTTATGGGTAG
3′
G*: cDPCP-dG.
Fig. 3.
Pol II transcription elongation blocked by a site-specific pyriplatin-
DNA adduct. (A) In vitro transcription with preformed pol II elongation com-
plexes A and A0 incubated with a mixture of ATP, CTP, and GTP (25 μM each).
Time points were taken after 0, 0.5, 1, 2, 3, 4, 8, 16, 32, or 64 min incubation.
The RNA transcripts in lanes 1–10 were taken from reactions of the pol II com-
plex with a nondamaged DNA template, whereas the RNA transcripts in lanes
11–20 were taken from reactions of the pol II complex with a site-specifically
damaged DNA template. The stalled RNA transcript is indicated by a black
arrow (Right), and the extended RNA transcript is visible to the left. The
length and sequences of RNA transcripts are given at the left margin of
the gel. (B) In vitro transcription with preformed pol II elongation complexes
C and C’ incubated with 25 μM GTP. The remainder of gel is the same as in A.
(C) In vitro transcription with preformed pol II elongation complexes C and C’
incubated with 25 μM of 30-dGTP. The rest of gel is same as in A. (D) In vitro
transcription with preformed pol II elongation complexes B and B’ incubated
with a mixture of 25 μM CTP and 30-dGTP. Time points were taken after 0, 0.5,
1, 2, 4, 8, 16, 32, 64 min of incubation. The rest of gel is the same as in A. (E) In
vitro transcription with preformed pol II elongation complexes B and B’
incubated with 25 μM CTP. The remainder of the gel is the same as in A.
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Wang et al.
The result is that the RNA transcript fails to extend beyond the
site of damage, subsequent translocation and nucleotide addition
being strongly inhibited. Several factors contribute to such
translocation inhibition, including (i) stabilization of the initial
pre-translocation state by interaction between the platinated
guanosine and pol II residues (Fig. 4B); (ii) a high translocation
energy barrier; and (iii) an unfavorable subsequent post-translo-
cation state induced by the DNA lesion. Hydrogen bonding inter-
actions between an ammine group of the cis-fPtðNH3Þ2ðpyÞg2þ
moiety with bridge helix partially help to stabilize the initial pre-
translocation state (Fig. 4B). To address the factors ii and iii, we
modeled the pyriplatin-damaged guanosine residue at the −1 po-
sition to mimic the state following translocation of the pyriplatin-
modified guanosine from the þ1 to −1 position. The structure
clearly reveals that the cis-fPtðNH3Þ2ðpyÞg2þ moiety serves as
a strong steric block, narrowing the space between the DNA
nucleotide base (−1) and the bridge helix and preventing the
downstream undamaged nucleoside base on the DNA template
strand from rotating into the canonical þ1 position (Fig. 5A).
Moreover, the fact that the cis-fPtðNH3Þ2ðpyÞg2þ moiety at
the −1 position sterically clashes with the downstream nucleotide
base at the þ1 position suggests that this final state is unfavorable
(Fig. 5 A and B). In summary, our results indicate that pyriplatin–
DNA adducts inhibit pol II transcription elongation by prevent-
ing subsequent translocation and nucleotide addition beyond the
site of damage.
Discussion
Insights into Structure-Activity Relationships (SARs) for the Monofunc-
tional Platinum Drug Family. The original SARs pertaining to
bifunctional platinum compounds such as cisplatin (8) were for-
mulated to explain why anticancer activity appeared to require
neutral, cis-[PtA2X2] compositions, in which A is an amine ligand
and X is a monoanionic leaving group. These rules are clearly
violated by cationic, monofunctional platinum compounds such
as pyriplatin (4, 5). Other monofunctional platinum complexes,
including ½PtðdienÞClþ, ½PtðNH3Þ3Clþ, and trans-½PtðNH3Þ2
ðpyÞClþ, are inactive and do not arrest pol II transcription,
whereas the cis-fPtðNH3Þ2ðpyÞg2þ unit bound to guanosine
blocks pol II transcription and has significant anticancer proper-
ties in mice when administered as pyriplatin (4, 5, 8, 24–32).
The present structure of pol II in complex with DNA site-
specifically modified by pyriplatin provides unique insight into
SARs to be expected for monofunctional platinum drug candi-
dates. We constructed models of potential stalled transcription
complexes containing DNA modified by the following three
representative
units,
fPtðNH3Þ3g2þ, trans-fPtðNH3Þ2ðpyÞg2þ,
and cis-fPtðNH3Þ2ðpyÞg2þ bound to guanosine in DNA and posi-
tioned in either the −1 or þ1 site of pol II, in order to mimic the
A
B
+1
-1
+1
-1
Bridge Helix
Bridge
Helix
Rpb2 528-534
+1
5’
-1
3’
5’
3’
T831
A828
5’
3’
5’
3’
Non-template
DNA
5’
3’
+2
RNA
Template
DNA
Template
DNA
RNA
3.9 Å
3.9 Å
Fig. 4.
Structure of pol II transcribing complex stalled at a site-specific
pyriplatin-DNA adduct after CMP incorporation. (A) The newly incorporated
matched CMP is highlighted in yellow. Other colors are as in Fig. 2. Interac-
tions of the damaged nucleotide and pol II residues are highlighted in (B).
The view is taken roughly from an ∼90 degree clockwise rotation along
the RNA/DNA helix axis from A. Nitrogen and oxygen atoms are depicted
in blue and red, respectively. Hydrogen bonds between ammine group of
the cis-fPtðNH3Þ2ðpyÞg2þ moiety and bridge helix residues are shown as black
dashed lines. The loop of Rpb2 828–834 is shown in green.
X
A
+1
-1
-2
X
+1
-1
-2
RNA
Template
DNA
Bridge
Helix
Non-template
DNA
Addition
Site
3’
5’
5’
5’
3’
3’
+1
-1
-1
+1
+2
-2
Bridge Helix
Addition
Site
3’
3’
5’
Template
DNA
RNA
5’
3’
B
Fig. 5.
Pol II translocation following CMP incorporation is inhibited by a site-
specific pyriplatin-DNA adduct. (A) The cis-fPtðNH3Þ2ðpyÞg2þ-guanosine unit
is superimposed with a nucleoside in −1 position shown in magenta and as a
surface view. In the latter, the nitrogen and oxygen atoms are highlighted in
blue and red, respectively. CMP at the 30-end of RNA chain is highlighted in
yellow. The bridge helix is shown in gray as a surface view. The nucleosides at
the þ1 and þ2 position of the template DNA are drawn in wheat and orange,
respectively. The rotation of the downstream nucleoside base during trans-
location, from the þ2 position to the þ1 position, is blocked by the cis-
fPtðNH3Þ2ðpyÞg2þ moiety, as indicated. Other colors are as in Fig. 2. (B)
The cis-fPtðNH3Þ2ðpyÞg2þ moiety of pyriplatin-dG adduct modeled at −1 posi-
tion clashes with the base at the þ1 position. Colors are as in A, and the view
as in Fig. 4B.
Wang et al.
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post- and pre-translocation states, respectively (Fig. S1). For
each modeled structure, we rotated the platinum unit about
the Pt-N7 bond by 360° and computed van der Waals energies
arising from contacts between platinum ligands and the rest
of the pol II complex (Figs. S5–S9). The fPtðNH3Þ3g2þ and
trans-fPtðNH3Þ2ðpyÞg2þ moieties could be readily accommo-
dated within the pol II active site over wide energy minima.
The lack of a significant steric clash for these two groups, in either
the −1 or þ1 position of the pol II transcribing complex, indicates
the absence of a barrier to transcriptional bypass (Figs. S6–S9).
This finding agrees with experiment. In contrast, the energy bar-
rier is prohibitively high for cis-fPtðNH3Þ2ðpyÞg2þ platinated
DNA modeled at −1 position, which is consistent with its ability
to block pol II bypass and the failure of pol II to reach the sub-
sequent post-translocation state (Figs. S5 and S8). The presence
of a pyridine or other bulky group in the cis configuration is
important
for
restricting
the
rotation
range
of
the
cis-
fPtðNH3Þ2ðpyÞg2þ moiety and thus rendering it a strong steric
block to translocation. For a fPtðNH3Þ3g2þ or trans-fPtðNH3Þ2
ðpyÞg2þ adduct at the −1 position, such a steric clash can be
avoided by rotation about the Pt-N7 bond, facilitating subsequent
pol II translocation. These results are fully consistent with
previous biochemical studies revealing that the latter two
DNA adducts are inactive and fail to block transcription (5, 7,
12, 26–33).
A Unique Molecular Mechanism of Pol II Transcription Inhibition. The
stalling mechanism of monofunctional platinum drugs of the
pyriplatin family is dramatically different from transcription inhi-
bition by cisplatin and UV-induced 1,2-intrastrand cross-links.
For the latter two DNA-modifications, a translocation barrier
prevents delivery of damaged bases to the active site and/or leads
to misincorporation of NTPs against the damage site, respectively
(19, 34). Monofunctional platinum-damaged residues, on the
other hand, can cross over the bridge helix and be accommodated
in the pol II active site. For Pt–dG adducts, the correct CMP
nucleotide can be efficiently incorporated against the damaged
guanosine. It is blockage of the subsequent translocation from
this position after incorporation of the cytosine nucleotide that
leads to inhibition of the RNA polymerase, but only when a bulky
pyridine ligand is present in the cis coordination site.
In conclusion, we report here the structure of a pol II transcri-
bing complex stalled at a site-specific monofunctional DNA
adduct, revealing a unique mechanism of transcription inhibition
by this kind of genome damage. The results establish a basis for
SARs that govern the anticancer drug potential of monofunc-
tional platinum-based DNA-damaging agents. Specific inter-
actions between pol II active site residues and the platinum
ligands are revealed, providing a structural framework for
rational design of more potent monofunctional pyriplatin analo-
gues. Because the spectrum of activity of pyriplatin is dramatically
different from that of cisplatin against an extensive panel of can-
cer cell lines but with reduced potency (7), this information will
be valuable for increasing the anticancer drug potential of this
family of compounds based on pol II stalling with concomitant
induction of apoptosis.
Methods
Preparation of Pol II Transcribing Complexes. Ten-subunit S. cerevisiae pol II
was purified as described (35). RNA oligonucleotides were purchased from
Dharmacon and DNA oligonucleotides were obtained from IDT. cis-
½PtðNH3Þ2ðpyÞClCl was prepared by Ryan Todd at MIT. The site-specifically
platinated template DNA was obtained as described (7).
Pol II transcribing complexes were assembled with the use of synthetic oli-
gonucleotides (10). Briefly, DNA and RNA oligonucleotides were annealed
and mixed with pol II in 20 mM Tris (pH 7.5), 40 mM KCl, and 5 mM DTT.
The final mixture contained 2 μM pol II, 10 μM site-specific pyriplatin-
damaged template DNA strand, and 20 μM nontemplate DNA and RNA oli-
gonucleotides. The mixture was kept for 1 h at room temperature, and excess
oligonucleotides were removed by ultrafiltration. Crystals were obtained
from solutions containing 390 mM ðNH4Þ2HPO4∕NaH2PO4, pH 5.9–6.3,
50 mM dioxane, 10 mM DTT, and 9–11% PEG6000. Crystals of transcribing
complexes were transferred in a stepwise manner to cryobuffer as described
(10, 11). For the structure of the pol II complex with CTP incorporation, 10 mM
CTP was added to the cryobuffer (10, 11).
Crystal Structure Determination and Analysis. Diffraction data were collected
on beam line 11-1 at the Stanford Synchrotron Radiation Laboratory. Data
were processed in DENZO and SCALEPACK (HKL2000) (36). Model building
was performed with the program Coot (37), and refinement was done with
REFMAC with TLS (CCP4i) (Table S1). In the structure of pol II complex with a
CTP incorporation against damaged guanosine residue, we also observed
additional weaker density within the second channel in comparison to the
nucleoside residue at the þ1 position, which may correspond to nonspecific
binding of a second CTP molecule through the soaking process. All structure
models in the figures were superimposed with nucleoside residues near the
active site using PYMOL (38).
Transcription Assay. Transcription assays were performed essentially as de-
scribed (11). In a typical reaction, 32P-labeled RNA oligonucleotide (10 pmol)
was annealed with template DNA 29mer (20 pmol, damaged or nondamaged
template) and nontemplate DNA 14mer (20 pmol) in elongation buffer
(20 mM Tris-HCl, pH 7.5, 40 mM KCl, 0.5 mM MgCl2) in a final volume of
20 μL. An aliquot of the annealed RNA/DNA hybrid was incubated with a five-
fold excess of pol II (final concentration of pol II 1.1 μM, of RNA oligonucleo-
tide 0.22 μM, and of DNA oligonucleotides 0.44 μM) for 10 min at room tem-
perature. Equal volumes of the NTP mixture solution were added (final
concentrations 25 μM) and the mixture was then incubated for 0, 0.5, 1,
2, 3, 4, 8, 16, 32, or 64 min at room temperature before addition of stop solu-
tion (final concentrations 5 M urea, 44.5 mM Tris-HCl, 44.5 mM boric acid,
26 mM EDTA, pH 8.0, Xylene Cyanol and Bromophenol Blue dyes). RNA pro-
ducts were analyzed by PAGE in the presence of urea. Visualization and
quantification of products were performed with the use of a PhosphorIma-
ger (Molecular Dynamics).
Computer
Modeling
Analysis.
Three
representative
platinum
units,
fPtðNH3Þ3g2þ, trans-fPtðNH3Þ2ðpyÞg2þ, and cis-fPtðNH3Þ2ðpyÞg2þ bound to
guanosine in DNA and positioned in either the −1 or þ1 site of pol II were
modeled to mimic the post- and pre-translocation states, respectively. The
vdW interaction energies between the three ligands at different orientations
and the rest of the pol II complex were systematically computed and taken as
direct indicators of steric effects.
The structure of the cis-fPtðNH3Þ2ðpyÞg2þ fragment on DNA in pol II is
available from the current study. Initial configurations for the other two
units, fPtðNH3Þ3g2þ and trans-fPtðNH3Þ2ðpyÞg2þ, were obtained by modeling.
Briefly, the same configuration of pol II, DNA, and RNA as found in the struc-
ture containing cis-fPtðNH3Þ2ðpyÞg2þ was used for these two complexes. The
geometry of the fPtðNH3Þ3g2þ moiety was taken from a previous structure
where it binds to a B-DNA dodecamer (PDB ID: 5BNA) (39). Docking was
achieved by aligning the damaged guanosine base of the two structures.
Finally, the trans-ammine group in fPtðNH3Þ3g2þ was replaced with a pyridine
ligand, and the Pt-N bond length was appropriately adjusted to obtain the
structure for trans-fPtðNH3Þ2ðpyÞg2þ. The same procedure was used to
generate structures at both þ1 and −1 positions.
The vdW energies were computed for different configurations generated
by rotating about the Pt-N7 bond from −180° to 180° for each platinum modi-
fication (see Figs. S5–S7). The rotation angle (φ) was defined to be positive
when rotating in the anticlockwise direction. In the configuration with
φ ¼ 0°, the plane composed of two Pt-N bonds of the ligand which are per-
pendicular to the Pt-N7 bond was set to be parallel to the damaged guano-
sine base. We noticed that, for fPtðNH3Þ3g2þ and trans-fPtðNH3Þ2ðpyÞg2þ, two
trans ammine groups were accommodated at slightly different configura-
tions, with φ ¼ 0° due to the different local environment, which leads to
slightly different energies between conformations with φ and φ 180°.
Because the purpose of our modeling study is to identify major steric clashes
instead of accurately computing free energy changes associated with rota-
tion of the ligand, which requires extensive conformational sampling, we
performed a simple average of the two energies (E1ðφÞ and E2ðφ 180°Þ)
based on their Boltzmann weights (T 298 K), eq 1,
¯E ¼ ðe−βE1E1 þ e−βE2E2Þ∕ðe−βE1 þ e−βE2Þ
[1]
to get a better estimate of vdW energy profiles.
9588
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www.pnas.org/cgi/doi/10.1073/pnas.1002565107
Wang et al.
The GROMACS simulation package was used to compute vdW energies
between the ligands and the pol II complex (40). A 20-Å cutoff was adopted
for computing the vdW interactions. The AMBER03 force field was used
for the pol II complex including protein, RNA, and DNA (41). The vdW force
field (Leonard–Jones potential) parameters for ligands were generated from
the AMTECHAMBER module of the AMBER 9 package (42) using the general
AMBER force field (GAFF) (43) developed for rational drug design. Since
the Pt atom is not in direct contact with the pol II complex and does not con-
tribute significantly to any steric effects, we excluded it from our vdW energy
calculations.
ACKNOWLEDGMENTS. This research was supported by the National Institute of
General Medical Sciences (NIH Pathway to Independence Award GM085136
to D.W. and GM49985 to R.D. Kornberg) and by the National Cancer Institute
(Grant CA034992 to S.J.L.). Portions of the research were carried out at the
Stanford Synchrotron Radiation Laboratory, a national user facility operated
by Stanford University on behalf of the U.S. Department of Energy, Office of
Basic Energy Sciences. The SSRL Structural Molecular Biology Program is sup-
ported by the Department of Energy, Office of Biological and Environmental
Research, and by the National Institutes of Health, National Center for
Research Resources, Biomedical Technology Program, and the National Insti-
tute of General Medical Sciences.
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PNAS
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no. 21
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9589
BIOCHEMISTRY
|
3M41
|
Crystal structure of the mutant V182A of orotidine 5'-monophosphate decarboxylase from Methanobacterium thermoautotrophicum
|
Conformational Changes in Orotidine 5′-Monophosphate
Decarboxylase: “Remote” Residues that Stabilize the Active
Conformation†
B. McKay Wood‡, Tina L. Amyes§, Alexander A. Fedorov∥, Elena V. Fedorov∥, Andrew
Shabila‡, Steven C. Almo∥, John P. Richard§, and John A. Gerlt*,‡
‡Departments of Biochemistry and Chemistry, University of Illinois at Urbana-Champaign
§Department of Chemistry, University at Buffalo, Buffalo, NY 14260
∥Department of Biochemistry, Albert Einstein College of Medicine, Bronx, NY 10461
Abstract
The structural factors responsible for the extraordinary rate enhancement (~1017) of the reaction
catalyzed by orotidine 5′-monophosphate decarboxylase (OMPDC) have not been defined.
Catalysis requires a conformational change that closes an active site loop and “clamps” the orotate
base proximal to hydrogen-bonded networks that destabilize the substrate and stabilize the
intermediate. In the OMPDC from Methanobacter thermoautotrophicus, a “remote” structurally
conserved cluster of hydrophobic residues that includes Val 182 in the active site loop is
assembled in the closed, catalytically active conformation. Substitution of these residues with Ala
decreases kcat/Km with a minimal effect on kcat, providing evidence that the cluster stabilizes the
closed conformation. The intrinsic binding energies of the 5′-phosphate group of OMP for the
mutant enzymes are similar to that for the wild type, supporting this conclusion.
Orotidine 5′-monophosphate decarboxylase (OMPDC1) is one of Nature’s best catalysts: the
reaction occurs with a rate enhancement of ~1017 and a proficiency of ~1023 M (2). The
reaction coordinate includes a vinyl carbanion intermediate (Scheme 1) (3, 4).
Structures of OMPDCs complexed with UMP or 6-hydroxyUMP, a n intermediate analog
(5-9), reveal hydrogen-bonded networks proximal to 1) C6 of the pyrimidine (Asp 70-Lys
72-Asp 75 in the OMPDC from Methanobacter thermoautotrophicus; MtOMPDC); and 2)
O2, N3, and O4 of the base (Ser 127-Gln 185). These destabilize the substrate (9) and
stabilize the intermediate, although the structural strategy for the latter is uncertain.
The 5′-phosphate group binds in a conserved motif at the ends of the seventh and eighth β-
strands of the (β/α)8-barrel structure, with interactions to backbone NH groups as well as the
guanidinium group of a conserved Arg (Arg 203 in MtOMPDC). These interactions are
important for catalysis: 1) kcat/Km for OMP exceeds that for orotidine by a factor of ~1011
for the OMPDC from Saccharomyces cerevisiae (ScOMPDC) (10); 2) kcat/Km for OMP
†This work was supported by NIH Grants GM039754 to J.P.R. and GM065155 to J.A.G.
*To whom correspondence should be addressed: J.A.G.: telephone, (217) 244-7414; fax: (217) 244-6538; j-gerlt@uiuc.edu..
SUPPORTING INFORMATION AVAILABLE
Descriptions of the experimental procedures. This material is available free of charge via the Internet at http://pubs.acs.org.
1Abbreviations: OMP, orotidine 5′-monophosphate; OMPDC, OMP decarboxylase; MtOMPDC, OMPDC from Methanobacter
thermoautotrophicus; ScOMPDC, OMPDC from Saccharomyces cerevisiae; EO, 1-(β-D-erythrofuranosyl)-orotic acid; HPi,
phosphate dianion; IBE, intrinsic binding energy.
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 December 26.
Published in final edited form as:
Biochemistry. 2010 May 4; 49(17): 3514–3516. doi:10.1021/bi100443a.
NIH-PA Author Manuscript
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exceeds that for 1-(β-D-erythrofuranosyl)orotic acid (EO; 5′-truncated OMP analog) by
factors of 5.2 × 108 and 3.6 × 108 for ScOMPDC and MtOMPDC, respectively (11, 12); and
3 ) phosphite dianion ( H Pi) activates decarboxylation of EO by factors of 5.6 × 105 M−1
and 2.9 × 105 M−1 for ScOMPDC and MtOMPDC, respectively (11, 12). The “intrinsic
binding energy” [IBE; (13)] of the 5′-phosphate/HPi 1) increases the affinity for the
substrate (e.g., OMP vs. orotidine/EO); and 2) enables decarboxylation by juxtaposition of
the substrate with the active site hydrogen-bonded networks (substrate destabilization and
intermediate stabilization). How the IBE promotes catalysis is unknown but required to
understand the structural basis for the rate enhancement.
A loop located at the end of the seventh β-strand closes over the active site when OMP binds
(Figure 1). Although the active site loops differ in both length and sequence in divergent
OMPDCs (12), each includes a spatially conserved Gln (Gln 185 in MtOMPDC) hydrogen-
bonded to both the 5′-phosphate and O2 of the pyrimidine as well as a conserved Ser at the
end of the fifth β-strand that also is hydrogen-bonded to N3 of the pyrimidine (Ser 127). We
characterized the importance of these “clamp” residues in ScOMPDC (Gln 215-Ser 154)
using EO/HPi and confirmed that the 5′-phosphate/HPi “switch” is required to activate the
enzyme (14).
The structures of divergent OMPDCs reveal that OMP binding is always accompanied by a
conformational change (Figure 1). The most obvious component is closure of the active site
loop. However, the (β/α)8-barrel structure can be divided into two domains, one formed
from the second, third, fourth, and fifth β-strands (where the hydrogen-bonded Asp 70-Lys
72-Asp 75 motif and Ser 127 are located) and the second from the sixth, seventh, eighth, and
first β-strands (where the phosphate binding motif and the active site loop, including Gln
185, are located) (15). OMP binding reorients the domains, with the latter domain moving
toward the former, enforcing the orotate carboxylate group to be juxtaposed vis a vis Asp
70- Lys 72-Asp 75 and, also, allowing formation of the Ser 127-Gln 185 “clamp”. Thus, the
transition between the open and closed conformations is more complicated than “simple”
hinge motion of the loop on the rigid framework of the (β/α)8-barrel structure. In this Report
we identify “remote” residues involved in this conformational change and quantitate their
importance in promoting and stabilizing the catalytically competent form of the enzyme.
The active site loop of MtOMPDC, P180-G181-V182-G183-A184-Q185-G186-G187-D188,
is disordered in the absence of substrate but ordered and closed in its presence (Figure 1). In
the liganded structure two residues in the loop make contacts with the (β/α)8-barrel scaffold:
1) Gln 185 is hydrogen-bonded to Ser 127 (vide infra); and 2) Val 182 is embedded in a
hydrophobic cluster also formed by Ile 199, Val 201, and Ile 218. The conservation of this
hydrophobic cluster in all OMPDCs suggests its importance in a common catalytic strategy.
We probed this strategy by mutagenesis of these hydrophobic residues.
Ala substitutions for residues in the hydrophobic cluster cause substantial decreases in kcat/
Km for decarboxylation of OMP but have little effect on kcat (Table 1). We determined high
resolution X-ray structures (≤ 1.4 Å) for each mutant in the presence of 6-hydroxyUMP
(Figure 2). The liganded structures superimpose well with that of wild type, with only small
differences observed at the sites of the substitutions (panel A). The active sites are identical
to that of wild type (panel B), explaining the minimal impact on kcat.
The mutated residues are remote from the active site (Figures 1 and 2). Thus, the effects of
the substitutions on kcat/Km cannot be explained by altered direct interactions with the
substrate. Instead, the effects can be explained by decreased stabilities of the closed
conformation in which the substrate is destabilized (9) and the anionic intermediate is
stabilized. A consistent model (Scheme 2) is that an equilibrium of open (Eo) and closed
Wood et al.
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(Ec) conformations exists in the absence or presence of substrate (KC’ or KC; KC’ << 1). The
substitutions destabilize Ec (decrease KC and KC ’), so kcat/Km is decreased (the energy
difference between Eo + S and [Ec•S]‡ is increased). However, the invariance of kcat
establishes that the substitutions do not alter the reactivitiy of Ec•S (the energy difference
between Ec•S and [Ec•S]‡). The energy to form Ec•S from Eo + S is achieved from 1)
interactions of the 5′-phosphate group with its binding motif (IBE; vide infra), and 2) an
increased concentration of OMP to form Ec•S (KC/KS or, equivalently, KC’/KS’, although
the former is expected to be the relevant pathway).
We also used the two part EO•HPi substrate. The values of kcat/Km for EO are decreased
relative to that for wild type (Table 1); these can be explained by decreased populations of
Ec (Scheme 2), assuming that EO, without a 5′-phosphate group, is unable to promote the
transition from Eo to Ec. The values of kcat/Km for OMP and kcat/Km for EO allow
calculation of the IBE for the 5′-phosphate group of OMP (Table 1).
HPi activates the mutants as judged by the values of the third-order rate constant, (kcat/
Km)EO•HPi/Kd. The equivalent changes (ΔΔG‡) in kcat/Km for both OMP and EO and the
third-order rate constant indicate that all three measure the effects of the substitutions on the
values of KC.
The values of the IBEs for the 5′-phosphate group for the mutants are the same as that for
the wild type, establishing that decarboxylation in the Ec•S complex occurs with equivalent
amounts of ground state destabilization (9) and transition state stabilization (as also reflected
by the invariant values of kcat). The IBEs provide further support for the role of the “remote”
hydrophobic cluster in stabilizing Ec relative to Eo but do not directly participating in
catalysis.
Our experiments implicate a structurally conserved hydrophobic cluster, Val 182, Ile 199,
Val 201, and Ile 218 in MtOMPDC, in stabilizing the closed conformation required for
catalysis. Its identification provides evidence that structural elements distal from the active
site, in addition to the proximal active site loop that closes to “clamp” the substrate, are
required for OMPDC’s extraordinary catalytic efficiency and proficiency.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
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Figure 1.
Superposition of the active site of wild type MtOMPDC in the absence (cyan; disordered
loop depicted with the dotted line ) and presence ( grey ) of 6-hydroxyUMP. The carbons of
6-hydroxyUMP are highlighted in yellow; the carbons of Val 182 and Arg 203 in the
liganded structure are highlighted in orange.
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Figure 2.
Superpositions of the 6-hydroxyUMP-liganded structures of wild type MtOMPDC and the
single mutants in the hydrophobic cluster. Panel A, hydrophobic cluster; panel B, active
sites.
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Scheme 1.
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Scheme 2.
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Table 1
Kinetic Constants for OMP, EO and EO/HPi and Intrinsic Binding Energies of the 5′-Phosphate Group of OMP at pH 7.1 and 25 °C.
MtOMPDC
kcat
OMP
s−1
kcat/Km
OMP
M−1 s−1
ΔΔG‡
kcal/mola
kcat/Km
EO
M−1 s−1
ΔΔG‡
kcal/mola
(kcat/Km)/K
D
EO•HPib
M−2 s−1
ΔΔG‡
kcal/mola
5′-Phosphate
IBEc
kcal/mol
Wild type
4.6
2.9 × 106
8.7 × 10−3
2500
11.6 d
V182A
3.4
1.4 × 105
1.8
1.3 × 10−3
1.1
190
1.5
10.9
I199A
3.9
9.1 × 105
0.7
1.9 × 10−3
0.9
980
0.6
11.8
V201A
4.0
9.5 × 105
0.7
3.1 × 10−3
0.6
690
0.8
11.5
I218A
3.3
2.8 × 105
1.4
2.3 × 10−3
0.8
340
1.2
11.0
V182A/I199A
3.1
4.9 × 104
2.4
3.9 × 10−4
1.8
81
2.0
11.0
V182A/V201A
2.5
4.9 × 104
2.4
5.0 × 10−4
1.7
30
2.6
10.9
aCalculated from the ratio of the second-order or third-order rate constants for the wild type and mutant enzyme.
bThird-order rate constant for reaction of EO/HPi.
cTransition state stabilization by the 5′-phosphate group of OMP, calculated from the ratio of the values of kcat/Km for OMP and EO. IBEs for phosphite dianion can be calculated from the ratio of (kcat/
Km)/KD for EOHPi and kcat/Km for EO.
Biochemistry. Author manuscript; available in PMC 2011 December 26.
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3M43
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Crystal structure of the mutant I199A of orotidine 5'-monophosphate decarboxylase from Methanobacterium thermoautotrophicum
|
Conformational Changes in Orotidine 5′-Monophosphate
Decarboxylase: “Remote” Residues that Stabilize the Active
Conformation†
B. McKay Wood‡, Tina L. Amyes§, Alexander A. Fedorov∥, Elena V. Fedorov∥, Andrew
Shabila‡, Steven C. Almo∥, John P. Richard§, and John A. Gerlt*,‡
‡Departments of Biochemistry and Chemistry, University of Illinois at Urbana-Champaign
§Department of Chemistry, University at Buffalo, Buffalo, NY 14260
∥Department of Biochemistry, Albert Einstein College of Medicine, Bronx, NY 10461
Abstract
The structural factors responsible for the extraordinary rate enhancement (~1017) of the reaction
catalyzed by orotidine 5′-monophosphate decarboxylase (OMPDC) have not been defined.
Catalysis requires a conformational change that closes an active site loop and “clamps” the orotate
base proximal to hydrogen-bonded networks that destabilize the substrate and stabilize the
intermediate. In the OMPDC from Methanobacter thermoautotrophicus, a “remote” structurally
conserved cluster of hydrophobic residues that includes Val 182 in the active site loop is
assembled in the closed, catalytically active conformation. Substitution of these residues with Ala
decreases kcat/Km with a minimal effect on kcat, providing evidence that the cluster stabilizes the
closed conformation. The intrinsic binding energies of the 5′-phosphate group of OMP for the
mutant enzymes are similar to that for the wild type, supporting this conclusion.
Orotidine 5′-monophosphate decarboxylase (OMPDC1) is one of Nature’s best catalysts: the
reaction occurs with a rate enhancement of ~1017 and a proficiency of ~1023 M (2). The
reaction coordinate includes a vinyl carbanion intermediate (Scheme 1) (3, 4).
Structures of OMPDCs complexed with UMP or 6-hydroxyUMP, a n intermediate analog
(5-9), reveal hydrogen-bonded networks proximal to 1) C6 of the pyrimidine (Asp 70-Lys
72-Asp 75 in the OMPDC from Methanobacter thermoautotrophicus; MtOMPDC); and 2)
O2, N3, and O4 of the base (Ser 127-Gln 185). These destabilize the substrate (9) and
stabilize the intermediate, although the structural strategy for the latter is uncertain.
The 5′-phosphate group binds in a conserved motif at the ends of the seventh and eighth β-
strands of the (β/α)8-barrel structure, with interactions to backbone NH groups as well as the
guanidinium group of a conserved Arg (Arg 203 in MtOMPDC). These interactions are
important for catalysis: 1) kcat/Km for OMP exceeds that for orotidine by a factor of ~1011
for the OMPDC from Saccharomyces cerevisiae (ScOMPDC) (10); 2) kcat/Km for OMP
†This work was supported by NIH Grants GM039754 to J.P.R. and GM065155 to J.A.G.
*To whom correspondence should be addressed: J.A.G.: telephone, (217) 244-7414; fax: (217) 244-6538; j-gerlt@uiuc.edu..
SUPPORTING INFORMATION AVAILABLE
Descriptions of the experimental procedures. This material is available free of charge via the Internet at http://pubs.acs.org.
1Abbreviations: OMP, orotidine 5′-monophosphate; OMPDC, OMP decarboxylase; MtOMPDC, OMPDC from Methanobacter
thermoautotrophicus; ScOMPDC, OMPDC from Saccharomyces cerevisiae; EO, 1-(β-D-erythrofuranosyl)-orotic acid; HPi,
phosphate dianion; IBE, intrinsic binding energy.
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Published in final edited form as:
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exceeds that for 1-(β-D-erythrofuranosyl)orotic acid (EO; 5′-truncated OMP analog) by
factors of 5.2 × 108 and 3.6 × 108 for ScOMPDC and MtOMPDC, respectively (11, 12); and
3 ) phosphite dianion ( H Pi) activates decarboxylation of EO by factors of 5.6 × 105 M−1
and 2.9 × 105 M−1 for ScOMPDC and MtOMPDC, respectively (11, 12). The “intrinsic
binding energy” [IBE; (13)] of the 5′-phosphate/HPi 1) increases the affinity for the
substrate (e.g., OMP vs. orotidine/EO); and 2) enables decarboxylation by juxtaposition of
the substrate with the active site hydrogen-bonded networks (substrate destabilization and
intermediate stabilization). How the IBE promotes catalysis is unknown but required to
understand the structural basis for the rate enhancement.
A loop located at the end of the seventh β-strand closes over the active site when OMP binds
(Figure 1). Although the active site loops differ in both length and sequence in divergent
OMPDCs (12), each includes a spatially conserved Gln (Gln 185 in MtOMPDC) hydrogen-
bonded to both the 5′-phosphate and O2 of the pyrimidine as well as a conserved Ser at the
end of the fifth β-strand that also is hydrogen-bonded to N3 of the pyrimidine (Ser 127). We
characterized the importance of these “clamp” residues in ScOMPDC (Gln 215-Ser 154)
using EO/HPi and confirmed that the 5′-phosphate/HPi “switch” is required to activate the
enzyme (14).
The structures of divergent OMPDCs reveal that OMP binding is always accompanied by a
conformational change (Figure 1). The most obvious component is closure of the active site
loop. However, the (β/α)8-barrel structure can be divided into two domains, one formed
from the second, third, fourth, and fifth β-strands (where the hydrogen-bonded Asp 70-Lys
72-Asp 75 motif and Ser 127 are located) and the second from the sixth, seventh, eighth, and
first β-strands (where the phosphate binding motif and the active site loop, including Gln
185, are located) (15). OMP binding reorients the domains, with the latter domain moving
toward the former, enforcing the orotate carboxylate group to be juxtaposed vis a vis Asp
70- Lys 72-Asp 75 and, also, allowing formation of the Ser 127-Gln 185 “clamp”. Thus, the
transition between the open and closed conformations is more complicated than “simple”
hinge motion of the loop on the rigid framework of the (β/α)8-barrel structure. In this Report
we identify “remote” residues involved in this conformational change and quantitate their
importance in promoting and stabilizing the catalytically competent form of the enzyme.
The active site loop of MtOMPDC, P180-G181-V182-G183-A184-Q185-G186-G187-D188,
is disordered in the absence of substrate but ordered and closed in its presence (Figure 1). In
the liganded structure two residues in the loop make contacts with the (β/α)8-barrel scaffold:
1) Gln 185 is hydrogen-bonded to Ser 127 (vide infra); and 2) Val 182 is embedded in a
hydrophobic cluster also formed by Ile 199, Val 201, and Ile 218. The conservation of this
hydrophobic cluster in all OMPDCs suggests its importance in a common catalytic strategy.
We probed this strategy by mutagenesis of these hydrophobic residues.
Ala substitutions for residues in the hydrophobic cluster cause substantial decreases in kcat/
Km for decarboxylation of OMP but have little effect on kcat (Table 1). We determined high
resolution X-ray structures (≤ 1.4 Å) for each mutant in the presence of 6-hydroxyUMP
(Figure 2). The liganded structures superimpose well with that of wild type, with only small
differences observed at the sites of the substitutions (panel A). The active sites are identical
to that of wild type (panel B), explaining the minimal impact on kcat.
The mutated residues are remote from the active site (Figures 1 and 2). Thus, the effects of
the substitutions on kcat/Km cannot be explained by altered direct interactions with the
substrate. Instead, the effects can be explained by decreased stabilities of the closed
conformation in which the substrate is destabilized (9) and the anionic intermediate is
stabilized. A consistent model (Scheme 2) is that an equilibrium of open (Eo) and closed
Wood et al.
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(Ec) conformations exists in the absence or presence of substrate (KC’ or KC; KC’ << 1). The
substitutions destabilize Ec (decrease KC and KC ’), so kcat/Km is decreased (the energy
difference between Eo + S and [Ec•S]‡ is increased). However, the invariance of kcat
establishes that the substitutions do not alter the reactivitiy of Ec•S (the energy difference
between Ec•S and [Ec•S]‡). The energy to form Ec•S from Eo + S is achieved from 1)
interactions of the 5′-phosphate group with its binding motif (IBE; vide infra), and 2) an
increased concentration of OMP to form Ec•S (KC/KS or, equivalently, KC’/KS’, although
the former is expected to be the relevant pathway).
We also used the two part EO•HPi substrate. The values of kcat/Km for EO are decreased
relative to that for wild type (Table 1); these can be explained by decreased populations of
Ec (Scheme 2), assuming that EO, without a 5′-phosphate group, is unable to promote the
transition from Eo to Ec. The values of kcat/Km for OMP and kcat/Km for EO allow
calculation of the IBE for the 5′-phosphate group of OMP (Table 1).
HPi activates the mutants as judged by the values of the third-order rate constant, (kcat/
Km)EO•HPi/Kd. The equivalent changes (ΔΔG‡) in kcat/Km for both OMP and EO and the
third-order rate constant indicate that all three measure the effects of the substitutions on the
values of KC.
The values of the IBEs for the 5′-phosphate group for the mutants are the same as that for
the wild type, establishing that decarboxylation in the Ec•S complex occurs with equivalent
amounts of ground state destabilization (9) and transition state stabilization (as also reflected
by the invariant values of kcat). The IBEs provide further support for the role of the “remote”
hydrophobic cluster in stabilizing Ec relative to Eo but do not directly participating in
catalysis.
Our experiments implicate a structurally conserved hydrophobic cluster, Val 182, Ile 199,
Val 201, and Ile 218 in MtOMPDC, in stabilizing the closed conformation required for
catalysis. Its identification provides evidence that structural elements distal from the active
site, in addition to the proximal active site loop that closes to “clamp” the substrate, are
required for OMPDC’s extraordinary catalytic efficiency and proficiency.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
REFERENCES
(1). Wood BM, Chan KK, Amyes TL, Richard JP, Gerlt JA. Biochemistry. 2009; 48:5510–5517.
[PubMed: 19435313]
(2). Radzicka A, Wolfenden R. Science. 1995; 267:90–93. [PubMed: 7809611]
(3). Toth K, Amyes TL, Wood BM, Chan K, Gerlt JA, Richard JP. J. Am. Chem. Soc. 2007;
129:12946–12947. [PubMed: 17918849]
(4). Amyes TL, Wood BM, Chan K, Gerlt JA, Richard JP. J. Am. Chem. Soc. 2008; 130:1574–1575.
[PubMed: 18186641]
(5). Miller BG, Hassell AM, Wolfenden R, Milburn MV, Short SA. Proc. Natl. Acad. Sci. U A. 2000;
97:2011–2016.
(6). Appleby TC, Kinsland C, Begley TP, Ealick SE. Proc atl Acad Sci U S A. 2000; 97:2005–2010.
(7). Harris P, Navarro Poulsen JC, Jensen KF, Larsen S. Biochemistry. 2000; 39:4217–4224.
[PubMed: 10757968]
(8). Wu N, Mo Y, Gao J, Pai EF. Proc. Natl. Acad. Sci. U S A. 2000; 97:2017–2022. [PubMed:
10681441]
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(9). Chan KK, Wood BM, Fedorov AA, Fedorov EV, Imker HJ, Amyes TL, Richard JP, Almo SC,
Gerlt JA. Biochemistry. 2009; 48:5518–5531. [PubMed: 19435314]
(10). Sievers A, Wolfenden R. Bioorg. Chem. 2005; 33:45–52. [PubMed: 15668182]
(11). Amyes TL, Richard JP, Tait JJ. J. Am. Chem. Soc. 2005; 127:15708–15709. [PubMed:
16277505]
(12). Toth K, Amyes TL, Wood BM, Chan KK, Gerlt JA, Richard JP. Biochemistry. 2009; 48:8006–
8013. [PubMed: 19618917]
(13). Jencks WP. Adv. Enzymol. Rel. Areas. Mol. Biol. 1975; 43:219–410.
(14). Barnett SA, Amyes TL, Wood BM, Gerlt JA, Richard JP. Biochemistry. 2008; 47:7785–4487.
[PubMed: 18598058]
(15). Harris P, Poulsen JC, Jensen KF, Larsen S. J. Mol. Biol. 2002; 318:1019–1029. [PubMed:
12054799]
(16). Go MK, Amyes TL, Richard JP. Biochemistry. 2009; 48:5769–5778. [PubMed: 19425580]
Wood et al.
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Figure 1.
Superposition of the active site of wild type MtOMPDC in the absence (cyan; disordered
loop depicted with the dotted line ) and presence ( grey ) of 6-hydroxyUMP. The carbons of
6-hydroxyUMP are highlighted in yellow; the carbons of Val 182 and Arg 203 in the
liganded structure are highlighted in orange.
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Figure 2.
Superpositions of the 6-hydroxyUMP-liganded structures of wild type MtOMPDC and the
single mutants in the hydrophobic cluster. Panel A, hydrophobic cluster; panel B, active
sites.
Wood et al.
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Scheme 1.
Wood et al.
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Scheme 2.
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Wood et al.
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Table 1
Kinetic Constants for OMP, EO and EO/HPi and Intrinsic Binding Energies of the 5′-Phosphate Group of OMP at pH 7.1 and 25 °C.
MtOMPDC
kcat
OMP
s−1
kcat/Km
OMP
M−1 s−1
ΔΔG‡
kcal/mola
kcat/Km
EO
M−1 s−1
ΔΔG‡
kcal/mola
(kcat/Km)/K
D
EO•HPib
M−2 s−1
ΔΔG‡
kcal/mola
5′-Phosphate
IBEc
kcal/mol
Wild type
4.6
2.9 × 106
8.7 × 10−3
2500
11.6 d
V182A
3.4
1.4 × 105
1.8
1.3 × 10−3
1.1
190
1.5
10.9
I199A
3.9
9.1 × 105
0.7
1.9 × 10−3
0.9
980
0.6
11.8
V201A
4.0
9.5 × 105
0.7
3.1 × 10−3
0.6
690
0.8
11.5
I218A
3.3
2.8 × 105
1.4
2.3 × 10−3
0.8
340
1.2
11.0
V182A/I199A
3.1
4.9 × 104
2.4
3.9 × 10−4
1.8
81
2.0
11.0
V182A/V201A
2.5
4.9 × 104
2.4
5.0 × 10−4
1.7
30
2.6
10.9
aCalculated from the ratio of the second-order or third-order rate constants for the wild type and mutant enzyme.
bThird-order rate constant for reaction of EO/HPi.
cTransition state stabilization by the 5′-phosphate group of OMP, calculated from the ratio of the values of kcat/Km for OMP and EO. IBEs for phosphite dianion can be calculated from the ratio of (kcat/
Km)/KD for EOHPi and kcat/Km for EO.
Biochemistry. Author manuscript; available in PMC 2011 December 26.
|
3M44
|
Crystal structure of the mutant V201A of orotidine 5'-monophosphate decarboxylase from Methanobacterium thermoautotrophicum
|
Conformational Changes in Orotidine 5′-Monophosphate
Decarboxylase: “Remote” Residues that Stabilize the Active
Conformation†
B. McKay Wood‡, Tina L. Amyes§, Alexander A. Fedorov∥, Elena V. Fedorov∥, Andrew
Shabila‡, Steven C. Almo∥, John P. Richard§, and John A. Gerlt*,‡
‡Departments of Biochemistry and Chemistry, University of Illinois at Urbana-Champaign
§Department of Chemistry, University at Buffalo, Buffalo, NY 14260
∥Department of Biochemistry, Albert Einstein College of Medicine, Bronx, NY 10461
Abstract
The structural factors responsible for the extraordinary rate enhancement (~1017) of the reaction
catalyzed by orotidine 5′-monophosphate decarboxylase (OMPDC) have not been defined.
Catalysis requires a conformational change that closes an active site loop and “clamps” the orotate
base proximal to hydrogen-bonded networks that destabilize the substrate and stabilize the
intermediate. In the OMPDC from Methanobacter thermoautotrophicus, a “remote” structurally
conserved cluster of hydrophobic residues that includes Val 182 in the active site loop is
assembled in the closed, catalytically active conformation. Substitution of these residues with Ala
decreases kcat/Km with a minimal effect on kcat, providing evidence that the cluster stabilizes the
closed conformation. The intrinsic binding energies of the 5′-phosphate group of OMP for the
mutant enzymes are similar to that for the wild type, supporting this conclusion.
Orotidine 5′-monophosphate decarboxylase (OMPDC1) is one of Nature’s best catalysts: the
reaction occurs with a rate enhancement of ~1017 and a proficiency of ~1023 M (2). The
reaction coordinate includes a vinyl carbanion intermediate (Scheme 1) (3, 4).
Structures of OMPDCs complexed with UMP or 6-hydroxyUMP, a n intermediate analog
(5-9), reveal hydrogen-bonded networks proximal to 1) C6 of the pyrimidine (Asp 70-Lys
72-Asp 75 in the OMPDC from Methanobacter thermoautotrophicus; MtOMPDC); and 2)
O2, N3, and O4 of the base (Ser 127-Gln 185). These destabilize the substrate (9) and
stabilize the intermediate, although the structural strategy for the latter is uncertain.
The 5′-phosphate group binds in a conserved motif at the ends of the seventh and eighth β-
strands of the (β/α)8-barrel structure, with interactions to backbone NH groups as well as the
guanidinium group of a conserved Arg (Arg 203 in MtOMPDC). These interactions are
important for catalysis: 1) kcat/Km for OMP exceeds that for orotidine by a factor of ~1011
for the OMPDC from Saccharomyces cerevisiae (ScOMPDC) (10); 2) kcat/Km for OMP
†This work was supported by NIH Grants GM039754 to J.P.R. and GM065155 to J.A.G.
*To whom correspondence should be addressed: J.A.G.: telephone, (217) 244-7414; fax: (217) 244-6538; j-gerlt@uiuc.edu..
SUPPORTING INFORMATION AVAILABLE
Descriptions of the experimental procedures. This material is available free of charge via the Internet at http://pubs.acs.org.
1Abbreviations: OMP, orotidine 5′-monophosphate; OMPDC, OMP decarboxylase; MtOMPDC, OMPDC from Methanobacter
thermoautotrophicus; ScOMPDC, OMPDC from Saccharomyces cerevisiae; EO, 1-(β-D-erythrofuranosyl)-orotic acid; HPi,
phosphate dianion; IBE, intrinsic binding energy.
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 December 26.
Published in final edited form as:
Biochemistry. 2010 May 4; 49(17): 3514–3516. doi:10.1021/bi100443a.
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exceeds that for 1-(β-D-erythrofuranosyl)orotic acid (EO; 5′-truncated OMP analog) by
factors of 5.2 × 108 and 3.6 × 108 for ScOMPDC and MtOMPDC, respectively (11, 12); and
3 ) phosphite dianion ( H Pi) activates decarboxylation of EO by factors of 5.6 × 105 M−1
and 2.9 × 105 M−1 for ScOMPDC and MtOMPDC, respectively (11, 12). The “intrinsic
binding energy” [IBE; (13)] of the 5′-phosphate/HPi 1) increases the affinity for the
substrate (e.g., OMP vs. orotidine/EO); and 2) enables decarboxylation by juxtaposition of
the substrate with the active site hydrogen-bonded networks (substrate destabilization and
intermediate stabilization). How the IBE promotes catalysis is unknown but required to
understand the structural basis for the rate enhancement.
A loop located at the end of the seventh β-strand closes over the active site when OMP binds
(Figure 1). Although the active site loops differ in both length and sequence in divergent
OMPDCs (12), each includes a spatially conserved Gln (Gln 185 in MtOMPDC) hydrogen-
bonded to both the 5′-phosphate and O2 of the pyrimidine as well as a conserved Ser at the
end of the fifth β-strand that also is hydrogen-bonded to N3 of the pyrimidine (Ser 127). We
characterized the importance of these “clamp” residues in ScOMPDC (Gln 215-Ser 154)
using EO/HPi and confirmed that the 5′-phosphate/HPi “switch” is required to activate the
enzyme (14).
The structures of divergent OMPDCs reveal that OMP binding is always accompanied by a
conformational change (Figure 1). The most obvious component is closure of the active site
loop. However, the (β/α)8-barrel structure can be divided into two domains, one formed
from the second, third, fourth, and fifth β-strands (where the hydrogen-bonded Asp 70-Lys
72-Asp 75 motif and Ser 127 are located) and the second from the sixth, seventh, eighth, and
first β-strands (where the phosphate binding motif and the active site loop, including Gln
185, are located) (15). OMP binding reorients the domains, with the latter domain moving
toward the former, enforcing the orotate carboxylate group to be juxtaposed vis a vis Asp
70- Lys 72-Asp 75 and, also, allowing formation of the Ser 127-Gln 185 “clamp”. Thus, the
transition between the open and closed conformations is more complicated than “simple”
hinge motion of the loop on the rigid framework of the (β/α)8-barrel structure. In this Report
we identify “remote” residues involved in this conformational change and quantitate their
importance in promoting and stabilizing the catalytically competent form of the enzyme.
The active site loop of MtOMPDC, P180-G181-V182-G183-A184-Q185-G186-G187-D188,
is disordered in the absence of substrate but ordered and closed in its presence (Figure 1). In
the liganded structure two residues in the loop make contacts with the (β/α)8-barrel scaffold:
1) Gln 185 is hydrogen-bonded to Ser 127 (vide infra); and 2) Val 182 is embedded in a
hydrophobic cluster also formed by Ile 199, Val 201, and Ile 218. The conservation of this
hydrophobic cluster in all OMPDCs suggests its importance in a common catalytic strategy.
We probed this strategy by mutagenesis of these hydrophobic residues.
Ala substitutions for residues in the hydrophobic cluster cause substantial decreases in kcat/
Km for decarboxylation of OMP but have little effect on kcat (Table 1). We determined high
resolution X-ray structures (≤ 1.4 Å) for each mutant in the presence of 6-hydroxyUMP
(Figure 2). The liganded structures superimpose well with that of wild type, with only small
differences observed at the sites of the substitutions (panel A). The active sites are identical
to that of wild type (panel B), explaining the minimal impact on kcat.
The mutated residues are remote from the active site (Figures 1 and 2). Thus, the effects of
the substitutions on kcat/Km cannot be explained by altered direct interactions with the
substrate. Instead, the effects can be explained by decreased stabilities of the closed
conformation in which the substrate is destabilized (9) and the anionic intermediate is
stabilized. A consistent model (Scheme 2) is that an equilibrium of open (Eo) and closed
Wood et al.
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(Ec) conformations exists in the absence or presence of substrate (KC’ or KC; KC’ << 1). The
substitutions destabilize Ec (decrease KC and KC ’), so kcat/Km is decreased (the energy
difference between Eo + S and [Ec•S]‡ is increased). However, the invariance of kcat
establishes that the substitutions do not alter the reactivitiy of Ec•S (the energy difference
between Ec•S and [Ec•S]‡). The energy to form Ec•S from Eo + S is achieved from 1)
interactions of the 5′-phosphate group with its binding motif (IBE; vide infra), and 2) an
increased concentration of OMP to form Ec•S (KC/KS or, equivalently, KC’/KS’, although
the former is expected to be the relevant pathway).
We also used the two part EO•HPi substrate. The values of kcat/Km for EO are decreased
relative to that for wild type (Table 1); these can be explained by decreased populations of
Ec (Scheme 2), assuming that EO, without a 5′-phosphate group, is unable to promote the
transition from Eo to Ec. The values of kcat/Km for OMP and kcat/Km for EO allow
calculation of the IBE for the 5′-phosphate group of OMP (Table 1).
HPi activates the mutants as judged by the values of the third-order rate constant, (kcat/
Km)EO•HPi/Kd. The equivalent changes (ΔΔG‡) in kcat/Km for both OMP and EO and the
third-order rate constant indicate that all three measure the effects of the substitutions on the
values of KC.
The values of the IBEs for the 5′-phosphate group for the mutants are the same as that for
the wild type, establishing that decarboxylation in the Ec•S complex occurs with equivalent
amounts of ground state destabilization (9) and transition state stabilization (as also reflected
by the invariant values of kcat). The IBEs provide further support for the role of the “remote”
hydrophobic cluster in stabilizing Ec relative to Eo but do not directly participating in
catalysis.
Our experiments implicate a structurally conserved hydrophobic cluster, Val 182, Ile 199,
Val 201, and Ile 218 in MtOMPDC, in stabilizing the closed conformation required for
catalysis. Its identification provides evidence that structural elements distal from the active
site, in addition to the proximal active site loop that closes to “clamp” the substrate, are
required for OMPDC’s extraordinary catalytic efficiency and proficiency.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
REFERENCES
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Figure 1.
Superposition of the active site of wild type MtOMPDC in the absence (cyan; disordered
loop depicted with the dotted line ) and presence ( grey ) of 6-hydroxyUMP. The carbons of
6-hydroxyUMP are highlighted in yellow; the carbons of Val 182 and Arg 203 in the
liganded structure are highlighted in orange.
Wood et al.
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Figure 2.
Superpositions of the 6-hydroxyUMP-liganded structures of wild type MtOMPDC and the
single mutants in the hydrophobic cluster. Panel A, hydrophobic cluster; panel B, active
sites.
Wood et al.
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Scheme 1.
Wood et al.
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Scheme 2.
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Table 1
Kinetic Constants for OMP, EO and EO/HPi and Intrinsic Binding Energies of the 5′-Phosphate Group of OMP at pH 7.1 and 25 °C.
MtOMPDC
kcat
OMP
s−1
kcat/Km
OMP
M−1 s−1
ΔΔG‡
kcal/mola
kcat/Km
EO
M−1 s−1
ΔΔG‡
kcal/mola
(kcat/Km)/K
D
EO•HPib
M−2 s−1
ΔΔG‡
kcal/mola
5′-Phosphate
IBEc
kcal/mol
Wild type
4.6
2.9 × 106
8.7 × 10−3
2500
11.6 d
V182A
3.4
1.4 × 105
1.8
1.3 × 10−3
1.1
190
1.5
10.9
I199A
3.9
9.1 × 105
0.7
1.9 × 10−3
0.9
980
0.6
11.8
V201A
4.0
9.5 × 105
0.7
3.1 × 10−3
0.6
690
0.8
11.5
I218A
3.3
2.8 × 105
1.4
2.3 × 10−3
0.8
340
1.2
11.0
V182A/I199A
3.1
4.9 × 104
2.4
3.9 × 10−4
1.8
81
2.0
11.0
V182A/V201A
2.5
4.9 × 104
2.4
5.0 × 10−4
1.7
30
2.6
10.9
aCalculated from the ratio of the second-order or third-order rate constants for the wild type and mutant enzyme.
bThird-order rate constant for reaction of EO/HPi.
cTransition state stabilization by the 5′-phosphate group of OMP, calculated from the ratio of the values of kcat/Km for OMP and EO. IBEs for phosphite dianion can be calculated from the ratio of (kcat/
Km)/KD for EOHPi and kcat/Km for EO.
Biochemistry. Author manuscript; available in PMC 2011 December 26.
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3M45
|
Crystal structure of Ig1 domain of mouse SynCAM 2
|
N-Glycosylation at the SynCAM (Synaptic Cell Adhesion
Molecule) Immunoglobulin Interface Modulates Synaptic
Adhesion*□
S
Received for publication,March 8, 2010, and in revised form, August 3, 2010 Published, JBC Papers in Press,August 25, 2010, DOI 10.1074/jbc.M110.120865
Adam I. Fogel‡1, Yue Li‡, Joanna Giza‡, Qing Wang‡2, TuKiet T. Lam§, Yorgo Modis‡, and Thomas Biederer‡3
From the ‡Department of Molecular Biophysics and Biochemistry and the §W. M. Keck Foundation Biotechnology Resource
Laboratory, Yale University, New Haven, Connecticut 06520
Select adhesion molecules connect pre- and postsynaptic
membranes and organize developing synapses. The regulation
of these trans-synaptic interactions is an important neurobio-
logical question. We have previously shown that the synaptic
cell adhesion molecules (SynCAMs) 1 and 2 engage in homo-
and heterophilic interactions and bridge the synaptic cleft to
induce presynaptic terminals. Here, we demonstrate that site-
specific N-glycosylation impacts the structure and function of
adhesive SynCAM interactions. Through crystallographic anal-
ysis of SynCAM 2, we identified within the adhesive interface of
its Ig1 domain an N-glycan on residue Asn60. Structural model-
ing of the corresponding SynCAM 1 Ig1 domain indicates that
its glycosylation sites Asn70/Asn104 flank the binding interface
of this domain. Mass spectrometric and mutational studies con-
firm and characterize the modification of these three sites.
These site-specific N-glycans affect SynCAM adhesion yet act in
a differential manner. Although glycosylation of SynCAM 2 at
Asn60 reduces adhesion, N-glycans at Asn70/Asn104 of SynCAM
1 increase its interactions. The modification of SynCAM 1 with
sialic acids contributes to the glycan-dependent strengthening
of its binding. Functionally, N-glycosylation promotes the trans-
synaptic interactions of SynCAM 1 and is required for synapse
induction. These results demonstrate that N-glycosylation of
SynCAM proteins differentially affects their binding interface
and implicate post-translational modification as a mechanism
to regulate trans-synaptic adhesion.
Synapses in the central nervous system are highly specialized
sites of neuronal adhesion. They are morphologically defined
by a presynaptic terminal filled with synaptic vesicles, an
apposed postsynaptic specialization that contains neurotrans-
mitter receptors, and a synaptic cleft of 20-nm width that sep-
arates pre- and postsynaptic sites (1, 2). This cleft is filled with
proteinaceous material (3).
The proteins spanning the synaptic cleft not only tie pre- and
postsynaptic membranes together. Select synaptic surface mol-
ecules can also instruct the organization of nascent synapses
(4–6). This was first demonstrated for neuroligins, postsynap-
tic membrane proteins that bind the presynaptic neurexins
(7–9). Adhesion molecules of the Ig superfamily and proteins
containing leucine-rich repeats additionally mediate synaptic
differentiation (10–14). Similarly, receptor tyrosine kinases,
including EphB receptors, instruct synaptogenesis through
trans-synaptic signaling (15, 16). These synapse-inducing pro-
teins act in conjunction with N-cadherins that set the pace of
synaptic maturation (17, 18).
Among these synapse-organizing proteins, SynCAMs4 form
a family of four Ig superfamily members that are single-span-
ning membrane proteins with three extracellular Ig-like
domains (19). SynCAMs, also known as nectin-like molecules,
are prominently expressed throughout the brain and are
enriched in synaptic plasma membranes (10, 20). They are
N-glycosylated proteins, consistent with the presence of
multiple predicted N-glycosylation sites in their extracellu-
lar Ig domains (19, 20). SynCAMs 1, 2, and 3 can bind them-
selves through homophilic binding, but SynCAMs 1/2 and
3/4 preferentially engage each other in specific heterophilic
interactions (20, 21). SynCAM 1 and 2 form a trans-synaptic
adhesion complex and promote the number of functional
excitatory synapses (20).
Consistent with the critical importance of synapse organiza-
tion for brain functions, trans-synaptic adhesion molecules
need to be regulated. Mechanisms include the control of their
sorting by intracellular interactions as shown for neuroligins
(22) and alternative splicing within sequences encoding extra-
cellular domains, which specifies neurexin-neuroligin inter-
actions and has been analyzed at atomic resolution (23–26).
Post-translational modifications regulating synaptic adhesion
molecules are less understood, but a negative effect of glycosy-
lation on neuroligin 1 binding has been reported (27).
* Thisworkwassupported,inwholeorinpart,byNationalInstitutesofHealth
Grant R01 DA018928 (to T. B.). This work was also supported by a Bur-
roughs Wellcome Investigator in the Pathogenesis of Infectious Disease
grant (to Y. M.), National Institutes of Health Predoctoral Program in Cellu-
lar and Molecular Biology Grant T32 GM007223, and by National Institutes
of Health Grant P30 DA018343 from the National Institute on Drug
Abuse. Use of the National Synchrotron Light Source is supported by the
Offices of Biological and of Basic Energy Sciences of the U.S. Department of
Energy.
□
S The on-line version of this article (available at http://www.jbc.org) contains
supplemental text, Table S1, and Figs. S1–S3.
1 Present address: NINDS, National Institutes of Health, 35 Convent Dr.,
Bethesda, MD 20892.
2 Present address: Program in Neurobiology and Behavior, Columbia Univer-
sity, New York, NY 10032.
3 To whom correspondence should be addressed: 333 Cedar St., New Haven
CT06520.Tel.:203-785-5465;Fax:203-785-6404;E-mail:thomas.biederer@
yale.edu.
4 The abbreviations used are: SynCAM, synaptic cell adhesion molecule; GPI,
glycosylphosphatidylinositol; PNGase F, peptide:N-glycosidase F; CHAPS,
3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid; Pn,
postnatal day n; FT-ICR, Fourier transform ion cyclotron resonance.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 45, pp. 34864–34874, November 5, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
34864
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 45•NOVEMBER 5, 2010
Here, we address the molecular properties that underlie and
regulate SynCAM adhesion and function. Our crystallographic,
mass spectrometric, and biochemical analyses of SynCAM 1
and 2 demonstrate that they carry N-glycans adjacent to and
within the first Ig domain that provides their extracellular bind-
ing interface. Unexpectedly, the glycosylation of these two
SynCAM family members serves different roles. Although
N-glycans within the Ig1 binding interface of SynCAM 2 reduce
its binding, glycosylation at the SynCAM 1 Ig1 domain pro-
motes its adhesion. Consequently, the ability to N-glycosylate
SynCAM 1 increases its trans-synaptic interactions and syn-
apse inducing activity. Together, these results identify glycosy-
lation as a novel mechanism for positively and negatively regu-
lating the trans-synaptic SynCAM adhesion complex in the
brain.
EXPERIMENTAL PROCEDURES
Antibodies—Specific antibodies against SynCAM 1 (YUC8)
and SynCAM 2 (YU524) were described previously (20). For
immunostaining of SynCAMs 1–3, we utilized a pleio-SynCAM
antibody (T2412) raised against the SynCAM 1 C terminus that
equally recognizes this conserved sequence in full-length
SynCAM 2 and 3 (20) but not the GPI-anchored SynCAM con-
structs used in this study. Monoclonal antibodies to synapto-
physin (7.2) and GDI (81.2) were obtained from Synaptic Sys-
tems (Go¨ttingen, Germany), monoclonal antibodies to CASK
were from Millipore (Billerica, MA), and monoclonal antibod-
ies to FLAG (M2) were from Sigma. Monoclonal antibodies to
SV2 (developed by Kathleen Buckley) were obtained from the
Developmental Studies Hybridoma Bank maintained by the
University of Iowa.
Expression Vectors—A pCMV5 GPI vector backbone was
generated by amplifying the GPI targeting sequence from GPI-
VAMP2 (a gift from Dr. James Rothman, Department of Cell
Biology, Yale University) with 5 SalI and 3 BamHI sites and
subcloning into pCMV5. pCMV5 FLAG-GPI was generated
analogously, adding the FLAG epitope DYKDDDDK N-termi-
nal of the GPI anchoring sequence. Full-length SynCAM extra-
cellular sequences or sequences lacking select Ig domains were
amplified from pCMV IG9 vectors described previously (10)
and subcloned into pCMV5 GPI or pCMV5 FLAG-GPI vectors.
Point mutants were generated using site-directed PCR mu-
tagenesis (QuikChange; Stratagene, La Jolla, CA). Expression
vectors for full-length SynCAM 1 carrying an extracellular
FLAG epitope N-terminal of the transmembrane region and for
extracellular SynCAM sequences fused to a thrombin cleavage
and IgG1-Fc sequence were described previously (20). GFP was
expressed from pCAG GFP, a gift from Dr. Nenad Sestan
(Department of Neurobiology, Yale University).
Cell Culture—COS7 cells were maintained using standard
procedures and transfected with FuGENE 6 (Roche Applied
Science) for transient expression. HEK293 cell lines stably
expressing the SynCAM 1 or 2 extracellular domains were
selected in the presence of geneticin (American Bioanalytical,
Natick, MA) after transfection of HEK293 cells with the vector
pcDNA3.1 SynCAM 1 extracellular domain IgG1 linearized
with BglII. Dissociated cultures of hippocampal neurons were
prepared as described (28).
Expression and Purification of SynCAM 1 and SynCAM 2
Extracellular Sequences—The extracellular sequences of mouse
SynCAM 1 or SynCAM 2 were purified as described previously
(20). Briefly, HEK293 cell lines stably expressing the full-length
SynCAM 1 or SynCAM 2 extracellular sequences fused to
IgG1-Fc were grown in DMEM low glucose medium supple-
mented with 5% FBS and 50 mg/ml geneticin. The culture
supernatant was collected and replaced with fresh medium
every 72 h. 2 liters of culture supernatant were filtered, concen-
trated, and then applied to 2 ml of protein A-agarose resin
(Invitrogen) equilibrated in buffer A (50 mM Tris, pH 8.0, 150
mM NaCl, 2 mM -mercaptoethanol). SynCAM extracellular
domains were eluted from the resin by the addition of bovine
-thrombin protease (Hematologic Technologies, Essex Junc-
tion, VT) in a 1:300 molar ratio at 16 °C overnight to cleave the
resin-bound N-terminal human IgG1-Fc tag. The proteins were
further purified by size exclusion chromatography on a Super-
dex 200 column (GE Healthcare) in buffer A.
Isothermal Titration Calorimetry—The binding of SynCAM
1 to SynCAM 2 was studied by isothermal titration calorimetry
in 20 mM Tris, pH 8.0, 50 mM NaCl, at 25 °C, using an iTC200
system (MicroCal, Piscataway, NJ). The sample cell contained
the purified SynCAM 2 extracellular domain protein at 5 M,
and the syringe contained the SynCAM 1 extracellular domain
at 50 M, with the IgG1-Fc tags cleaved off. Typically, one initial
injection of 1.5 l and 19 serial injections of 2.0 l of SynCAM
1 were performed at 180-s intervals. The stirring speed was
maintained at 1000 rpm, and the reference power was kept
constant at 5 cal/s. The heat associated with each injection of
SynCAM 1 was integrated and plotted against the molar ratio of
SynCAM 1 to SynCAM 2. Thermodynamic parameters were
extracted from a curve fit to the data using the Origin 7.0 soft-
ware provided by MicroCal. The experiments were performed
in triplicate with excellent reproducibility (10% variation in
thermodynamic parameters).
Preparation of SynCAM Extracellular Domain Complexes—
SynCAM 2 was first expressed and applied to protein A-agarose
resin as described above. The protein was then eluted from the
resin with 0.2 M glycine, pH 3.0, and dialyzed into 20 mM Tris,
pH 8.0, 50 mM NaCl, 2 mM -mercaptoethanol. A 2-fold molar
excess of the purified SynCAM 1 extracellular domain with the
IgG1-Fc tag cleaved off was then added to obtain heteromeric
SynCAM 1-SynCAM 2 complexes in addition to the homo-
meric SynCAM 2 complexes present in this preparation. The
resulting mixture was incubated at 16 °C for 3 h and applied to
2 ml of protein A-agarose resin in buffer A. The SynCAM com-
plexes were eluted from the resin with thrombin protease
(1:300 molar ratio, 16 °C overnight) and were purified on a
Superdex 200 column in buffer A. To aid in the subsequent
crystallization step, the samples were partially deglycosy-
lated under native conditions with PNGase F and neuramin-
idase (New England Biolabs) at 37 °C for 48 h and separated
from the endoglycosidases and cleaved glycans on a Super-
dex 200 column in 50 mM Tris, pH 8.0, 50 mM NaCl, 2 mM
-mercaptoethanol.
Crystallization of the Ig1 Domain of SynCAM 2—Crystals
were grown at 20 °C using the hanging drop vapor diffusion
technique. The preparation of the SynCAM extracellular
Glycans Modulate SynCAM Adhesion
NOVEMBER 5, 2010•VOLUME 285•NUMBER 45
JOURNAL OF BIOLOGICAL CHEMISTRY 34865
domain complex was concentrated to 9.0 mg/ml in 20 mM Tris,
pH 8.0, 50 mM NaCl, 2 mM -mercaptoethanol. The protein
solution was mixed with an equal volume of well solution (0.1 M
HEPES, pH 6.5–7.0, 21% PEG5000 monomethyl ether). Irregu-
lar bulky crystals grew after 3 months. The crystals were tri-
clinic (space group P1) with unit cell dimensions a 42.8 Å,
b 50.5 Å, c 79.9 Å, 75.9°, 77.0°, and 65.2°. With
four molecules/asymmetric unit, the Matthews co-efficient VM
is 3.36 Å3 Da1, which corresponds to a solvent content of
63.4% (29). For data collection, the crystals were transferred to a
cryoprotectant containing 0.1 M HEPES, pH 6.8, 21% PEG5000
monomethyl ether and 18% glycerol (v/v) and immediately fro-
zen in liquid nitrogen.
Data Collection and Processing—Crystallographic data were
collected at 100 K on Beamline X29A of the National Synchro-
tron Light Source at Brookhaven National Laboratory. The data
were indexed, integrated, and scaled using the HKL2000 pro-
gram suite (30). The data collection statistics are summarized in
supplemental Table S1.
Structure Determination and Refinement—The crystal struc-
ture of SynCAM 2 Ig1 was determined by molecular replace-
ment using a monomer of human SynCAM 3 (nectin-like mol-
ecule 1) Ig1, Protein Data Bank Code 1Z9M (31), as the search
model in the program PHASER 2.1 (32). The details of structure
determination and refinement are described in the supplemen-
tal materials.
Glycopeptide Mapping—The purified SynCAM 1 extracellu-
lar domain with the IgG1-Fc cleaved off was digested with the
combined endoproteinases Lys C and trypsin. The samples
were C18 RP ZipTip-cleaned and desalted prior to collecting
MS data on a 9.4T Apex Qe FT-ICR MS instrument. Eluted
peptides were directly infused into the mass spectrometer via
nanoelectrospray at 250 nL/min into an Apollo II dual ion fun-
nel ESI source. The spray shield voltage was set at 3500, and a
4000-V potential was applied on the glass capillary end cap. The
instrument (running Compass Software with APEX control
acquisition component (v.1.2) is set up to acquire single free
induction decay signal (512,000) data with a mass range (m/z)
from 450 to 2000. Enrichment of glycopeptides was confirmed
using albumin, ovalbumin, -casein, and RNase B as standard
proteins. All of the data were processed utilizing DA analysis
software v. 3.4, online GlycoMod (Expasy), and MASCOT
search engine.
Glycan Profiling—Glycosidase treatment of the SynCAM 1
extracellular sequence with the IgG1-Fc cleaved off was per-
formed using the glycosidases PNGase F or endoglycosidase H.
Cleaved glycans were enriched with a Carbograph column, fol-
lowed by C18 RP ZipTip prior to direct infusion into a 9.4T
FT-ICR MS instrument. The electrospray source was config-
ured with a capillary (low flow) sprayer optimized for positive
mode. Ions were detected in the 450–2500 m/z with 512K data
points/MS scan. All of the data were processed utilizing DA
analysis software v.3.4, online GlycoMod (Expasy).
Tissue Preparation—Samples from rat brain regions were
prepared by rapid homogenization in 8 M urea. Protein concen-
trations were determined using the Pierce BCA assay.
ProteinDeglycosylation—Enzymaticdeglycosylationwasper-
formed using neuraminidase (sialidase; Roche Applied Science)
and PNGase F (New England Biolabs) according to the manu-
facturers’ instructions.
Affinity Chromatography—The SynCAM 1 extracellular do-
main was immobilized on protein A beads to serve as affinity
matrix. Rat forebrain proteins were solubilized with 1% CHAPS
(Roche Applied Science), and affinity chromatography and
quantitative immunoblotting were performed as described
(10, 20).
Surface Expression Control—COS7 cells expressing SynCAM
constructs tagged with an extracellular FLAG epitope were
fixed, labeled with anti-FLAG antibodies to detect surface-ex-
pressed epitopes (antibody M2; 1:1000), washed, and then per-
meabilized using 0.1% Triton X-100 to perform immuno-
staining for total SynCAM protein (antibody T2412; 1:1000).
The images were acquired on a Zeiss LSM 510 META laser
scanning confocal microscope.
Cell Overlay Experiments—Cell overlay assays were per-
formed as described (21). Briefly, COS7 cells were co-trans-
fected with expression vectors encoding extracellularly FLAG-
tagged SynCAM constructs and soluble GFP or GFP alone as
negative control. After 2 days, live cells were overlaid for 20 min
at 25 °C with the purified SynCAM 1 extracellular domain at 2
g/ml or the SynCAM 2 extracellular domain at 10 g/ml. The
IgG1-Fc fusion tag of these overlaid fusion proteins was directly
detected by including Alexa 546-conjugated protein A (6
g/ml; Invitrogen) in this step. Surface-expressed SynCAM
proteins were detected in these live cells by simultaneously add-
ing anti-FLAG (antibody M2; 1:1000) and secondary anti-
mouse antibodies conjugated to Alexa 488 (Invitrogen)
(1:1000). The medium was then replaced with DMEM without
phenol red, and the cells were immediately imaged with a
Hamamatsu Orca camera attached to a Nikon Eclipse
TE2000-U microscope. The signal of the secondary Alexa 488
antibody detecting anti-FLAG antibodies was used to define
regions of interest, within which the fluorescence from the
Alexa 546-conjugated protein A was measured and normalized
to the anti-FLAG signal. Signals were quantified using a custom
Matlab (MathWorks) script that is available upon request.
Mixed Co-culture Assay for Synapse Induction—Co-culture
assays were performed as described (28). Briefly, COS7 cells
co-expressing GPI-anchored SynCAM 1 constructs and soluble
GFP or GFP alone as negative control were seeded atop neurons
at 6–7 days in vitro. At 8–9 days in vitro, these mixed co-cul-
tures were fixed and immunostained for the presynaptic
marker SV2 and for neuronal SynCAM proteins with the anti-
body T2412. The images were acquired on a Zeiss LSM 510
META laser scanning confocal microscope. The surface area of
COS7 cells immunopositive for neuronal SynCAMs and SV2
was quantified using a Matlab script that is available upon
request. The images were collected blind to the synaptic marker
channel.
Miscellaneous Procedures—Sequence similarities were ana-
lyzed using the T-Coffee method (33). Amino acid numbers
refer to the position in the protein including the signal peptide.
Statistical analyses were performed using the two-tailed t test,
with statistical errors corresponding to the standard errors of
mean.
Glycans Modulate SynCAM Adhesion
34866
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 45•NOVEMBER 5, 2010
RESULTS
High Affinity Binding of SynCAM 1 to SynCAM 2 Requires the
Ig1 Domain—To define the molecular properties of SynCAM
interactions, we measured the affinity between the SynCAM 1
and SynCAM 2 extracellular sequences by isothermal titration
calorimetry. The resulting isotherm was consistent with a sin-
gle binding interface between the two proteins in a 1:1 complex,
with a tight apparent dissociation constant (Kd) of 78 nM (Fig.
1A). This Kd is very similar to the neuroligin 1/neurexin 1
interaction (34).
We next mapped this single bind-
ing interface within the three extra-
cellular Ig-like domains. Utilizing
constructs comprised of subsets of
SynCAM 1 Ig domains, we mea-
sured their adhesive interaction with
the SynCAM 2 extracellular domain
using a cell overlay approach (Fig.
1B). These experiments extended
previous affinity chromatography
studies (20) and allowed us to ana-
lyze SynCAM interactions as they
occur on the cell surface. We
expressed an array of SynCAM 1 Ig
constructs carrying an extracellular
FLAG
epitope
and
labeled
the
expressed proteins in live COS7
cells with anti-FLAG antibodies. To
quantify adhesive binding, we over-
laid these cells with the soluble
SynCAM 2 extracellular sequence
fused
to
IgG1-Fc
and
detected
retained protein using fluorophore-
labeled protein A. This signal was
divided by the fluorescence mea-
sured with anti-FLAG antibodies,
which normalized for each cell the
extent of SynCAM 2 retention to
the amount of its surface-expressed
SynCAM 1. All of the SynCAM 1 Ig
constructs were properly N-glyco-
sylated and sorted to the plasma
membrane, with the Ig1 domain
carrying N-glycans to the highest
apparent extent (supplemental Fig.
S1). These experiments showed that
the tandem Ig1 2 domains of
SynCAM 1 were sufficient for strong
binding (Fig. 1, B and C). Moreover,
the SynCAM 1 Ig1 domain was
required for binding because the
SynCAM 1 Ig2 3 construct did
not retain SynCAM 2. The SynCAM
1 Ig1 domain alone was sufficient
for SynCAM 2 binding, albeit at
a lower strength. This reduced
interaction of the SynCAM 1 Ig1
domain in the absence of the Ig2
and Ig3 domains is possibly due to a role of these domains in
conferring a steric orientation to SynCAM 1 Ig1 that is favor-
able for its interaction with SynCAM 2. Together, these
results show that the first Ig domain of SynCAM 1 provides
its binding interface.
Crystal Structure of the SynCAM 2 Ig1 Domain—Aiming to
characterize the extracellular SynCAM interactions at atomic
resolution, we performed crystallization trials of the SynCAM
1/2 extracellular domain complex. The crystal structure, which
was refined at 2.21 Å resolution (r 0.197, Rfree 0.245),
FIGURE1.ThefirstIgdomainmediatestightheterophilicbindingofSynCAM1toSynCAM2.A,isothermal
titration calorimetry analysis of the binding of the SynCAM 1 extracellular sequence to SynCAM 2. Left panel,
enthalpicheatreleasedat25 °CduringthetitrationoftheSynCAM1extracellularsequenceintotheisothermal
titration calorimetry cell containing the SynCAM 2 extracellular sequence. Right panel, integrated binding
isotherms of the titration and best fit to a single-site model. The best fit yielded a dissociation constant Kd
78.0 nM, enthalphy H 9.1 kcal/mol, and binding stoichiometry n 1. B, analysis of adhesive SynCAM
binding by cell overlay. COS7 cells expressed full-length SynCAM 1 or variants containing the indicated Ig
domains, with an extracellular FLAG epitope inserted proximal to the transmembrane region. These surface-
expressed proteins were detected in live cells by the addition of anti-FLAG antibodies and secondary antibod-
ies conjugated to Alexa 488 (top row, green in the merge). The cells were simultaneously overlaid with the
SynCAM 2 extracellular domain fused to IgG1-Fc together with protein A conjugated to Alexa 546 (second row,
red in the merge) to label the retained protein. The first Ig domain of SynCAM 1 is required for adhesive binding
to SynCAM 2 as depicted in the model below. C, quantification of the results in B. The results are expressed as
a protein A signal detecting retained SynCAM 2 normalized to the signal from the indicated anti-FLAG labeled
SynCAM 1 constructs expressed on COS7 cells. *, p 0.05; **, p 0.01; ***, p 0.001.
Glycans Modulate SynCAM Adhesion
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JOURNAL OF BIOLOGICAL CHEMISTRY 34867
showed that the crystals contained only the Ig1 domain of
SynCAM 2. This was consistent with the presence of a major
protein band at 17 kDa in these crystals (data not shown), cor-
responding to the size of one Ig domain. Upon closer examina-
tion of the drop that produced the crystals, fungal growth was
observed. This suggested that secreted fungal proteases may
have cleaved the SynCAM 1/2 extracellular domain complex,
allowing SynCAM 2 Ig1 to crystallize by itself. Several other
proteins have been crystallized as a result of either intentional
proteolytic cleavage or serendipitous cleavage by secreted fun-
gal proteases (35, 36). The crystal structure of the SynCAM 2
Ig1 monomer (residues 35–131) showed that it adopts an Ig-
like fold of the variable type (37) as
predicted by sequence analysis (19),
comprising two -sheets with nine
antiparallel -strands (denoted A to
G; Fig. 2A). Hydrophobic interac-
tions between the two sheets form
the core of the domain. A disulfide
bridge between Cys53 and Cys113
links -strands B and G, further sta-
bilizing the domain. Two N-linked
N-acetylglucosamine residues are
visible in the structure on Asn40 and
Asn60, respectively. The remainder
of these N-glycans had been re-
moved during sample preparation
to aid crystallization (see “Experi-
mental Procedures”).
Interestingly, SynCAM 2 Ig1
forms homodimers in the crystals,
with each asymmetric unit contain-
ing two dimers. The SynCAM 2
Ig1 homodimer has approximate
dimensions of 60 42 33 Å (Fig.
2A). Dimer formation buries a total
of 698 Å2 (11.5%) of solvent-accessi-
ble surface/monomer. The N and C
termini of each subunit in the dimer
are antiparallel, indicating that the
dimer corresponds to the trans-ad-
hesion complex. The dimer inter-
face is mostly hydrophobic (35% of
the residues are nonpolar) and
closely resembles the dimer inter-
face of the Ig1 domain of SynCAM
3, also known as nectin-like mole-
cule 1 (31), which is its closest struc-
tural homolog. SynCAM 3 also par-
ticipates in cell adhesion (20, 38).
The Ig1 domains of SynCAM 2 and
SynCAM 3 have high sequence
identity (63%) and structural simi-
larity (root mean square deviation,
0.7 Å over 96 equivalent C atoms).
However, although the SynCAM 3
structure lacks glycans, in SynCAM
2 the N-acetylglucosamine on Asn60
forms a weak intersubunit contact in the trans-dimer interface
of SynCAM 2 Ig1 (Fig. 2, A and B). Specifically, the carbonyl
oxygen atom in the acetyl moiety of the first residue of the
glycan is within 3.5 Å of the N atom of the Arg82 side chain in
the other monomer and of a structured water molecule located
in the dimer interface. The location of Asn60 at the dimer inter-
face leaves little room for a bulky glycan, however, suggesting
that full glycosylation at Asn60 may interfere with adhesive
dimer formation.
Homology Model of the SynCAM 1/2 Ig1 trans-Heterodimer—
Like SynCAM 2, SynCAM 1 engages in both homo- and het-
erophilic adhesion complexes (20, 21). The high sequence
C
N
N
A
B
C
C’
D
E
F
G
a1
B
C
A
C’
D E
F
G
a1
A
A
B
B
C
C
C’
C’
D
D
E
E
F
F
G
G
a1
a1
N60
R82
N60
R82
3.5 Å
3.5 Å
N60
N60
F92
F92
K96
K96
N60
N60
N60
N60
N40
N40
N40
N40
C
90o
N
N
C
C
FIGURE 2. Structure of the SynCAM Ig1 domain interface. A, crystallographic results show that the SynCAM
2 Ig1 domain forms a dimer, characterized by mostly hydrophobic interactions across a noncrystallographic
2-fold axis, as shown in this ribbon representation. The N and C termini are marked. The N-acetylglucosamine
residues on Asn40 and Asn60 are marked as spheres. B, stereodiagram of the SynCAM 2 Ig1 trans-homodimer
interface, with the hydrogen bonding interactions of the N-acetylglucosamine on Asn60 shown as dashed lines.
Side chains in the interfaces are shown in stick representation. Water molecules are highlighted as red spheres.
C, theoretical model of the trans-heterodimer interface of SynCAM 1 Ig1/SynCAM 2 Ig1 shown as stereodia-
gram. Residues in the interface are depicted in stick representation, with SynCAM 1 and SynCAM 2 in green and
magenta, respectively. Interfaces are displayed in the same orientation as in B. The N-linked glycan on Asn60 of
SynCAM 2 participates in the SynCAM 1 Ig1/SynCAM 2 Ig1 trans-heterodimer interface.
Glycans Modulate SynCAM Adhesion
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identity of 44% between the Ig1 domains of SynCAM 1 and
SynCAM 2 enabled us to build a homology model for SynCAM
1 Ig1 based on our crystal structure of SynCAM 2 Ig1. Our
model predicts that the trans-dimeric interface of the SynCAM
1 Ig1 homodimer is more hydrophobic than that of SynCAM 2
Ig1 (supplemental Fig. S2A). The residues that form additional
hydrophobic contacts in the SynCAM 1 model are Val76, Phe92,
and Pro94.
Although capable of homophilic binding, SynCAMs pref-
erentially assemble into specific heterophilic complexes, and
SynCAM 1 strongly binds SynCAM 2 (20, 21, 38–40). To
better understand dimerization specificities, we modeled
SynCAM 1 Ig1/SynCAM 2 Ig1 trans-heterodimers using the
homodimeric crystal structure of SynCAM 2 Ig1 as template.
Interestingly, a glycan-mediated contact occurs in the het-
erodimer model between the N-linked glycan on Asn60 of
SynCAM 2 and the side chain of Lys96 of SynCAM 1 (Fig.
2C). The Asn60 site of SynCAM 2 is not conserved in
SynCAM 1, and this glycan may contribute to regulating the
heterophilic binding of SynCAM 2
to SynCAM 1. Conversely, two
N-glycosylation sites of the Ig1
domain of SynCAM 1, Asn70 and
Asn104, are located on one face of
the
Ig1
domain,
in
the
loop
between strands B and C and in
the middle of strand E, respec-
tively
(supplemental
Fig.
S2B).
Residues 70 and 104 are both 20
Å from the trans-dimer interface
and face away from the interface.
N-Glycosylation of the SynCAM
1 Ig1 Domain—These crystallo-
graphic results map glycans to dif-
ferent surfaces of the Ig1 domain in
SynCAM 1 and SynCAM 2. To
examine SynCAM 1 N-glycosyla-
tion, we performed a mass spec-
trometry analysis of the SynCAM
1 extracellular sequence purified
from HEK293 cells, which glycosy-
late SynCAM 1 to the same appar-
ent extent as found in brain (20).
Using several different enzymes or
combinations thereof, we observed
a very high number of extracellular
SynCAM 1 peptides/glycopeptides,
which provide 70% sequence cov-
erage of the protein. Glycopeptide
mapping identified the asparagines
Asn70 and Asn104 in the first Ig1
domain as potential N-glycosylation
sites based on FT-ICR high mass
accuracy and GlycoMod prediction
(41) (Fig. 3). The deconvoluted mass
list of each spectrum was entered
into GlycoMod to predict possible
glycosylations sites along with their
potential glycan composition based on mass accuracy and the
consensus sequence for N-glycosylation (42). The GlycoMod
output identifies the Asn104 site as glycosylated (in the CNBr
trypsin and LysC trypsin digest conditions; Fig. 3, B and C),
with several glycopeptides showing sialylated glycan (NeuAc)
modifications at that site. N-Glycosylation at Asn70 was pre-
dicted from the mass list of SynCAM 1 digested with CNBr
trypsin (Fig. 3B). To obtain a profile of these N-glycan struc-
tures, we subjected SynCAM 1 to PNGase F or endoglycosidase
H to cleave the glycans. Glycan masses observed by FT-ICR
suggested the presence of Hex, HexNac, and NeuAc carbohy-
drates as predicted by GlycoMod (data not shown).
Complex Modification of SynCAM 1 and SynCAM 2 in the
Brain—SynCAM 1 and 2 are heavily glycosylated in the adult
brain (20). To obtain insight into the extent of post-transla-
tional SynCAM modification during early postnatal develop-
ment, when most synapses form, we analyzed SynCAM 1 and 2
in several rat brain regions (Fig. 4). SynCAM modification was
examined by immunoblotting prior to, during, and subsequent
FIGURE 3. Glycopeptide mapping and glycan profiling of the SynCAM 1 extracellular sequence.
Broadband FT-ICR MS mass spectrum of four different enzymatic digestion of the purified, glycosylated
SynCAM 1 extracellular sequence. 10 g of SynCAM 1 was utilized for each digestion with CNBr (A), CNBr
trypsin (B), Lys C trypsin (C), and protease type XIII (D). The asterisks indicate potential glycosylation
sites based on exact mass measurements and GlycoMod prediction (41). The inset in B shows an enlarged
region illustrating a predicted glycopeptide at m/z 1606.413 (3) that corresponds to a modification at
the Asn70 position. Note that internal calibrations were utilized to obtain mass accuracy of 5 ppm. The
inset in C shows an enlarged region with a glycopeptide that corresponds to the modified Asn104 residue
of SynCAM 1 at m/z 1341.616 (3).
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JOURNAL OF BIOLOGICAL CHEMISTRY 34869
to the peak of synapse formation in the rodent brain at postna-
tal day 4 (P4), P16, and P30, respectively (43). At all stages and in
all regions, the apparent molecular masses of SynCAM 1 and 2
proteins were notably higher than the 41–45 kDa predicted
from their open reading frames (19). Interestingly, SynCAM 1
modifications changed during development. At P4, it was
expressed as diverse species that ranged from 90 to 115 kDa. As
development progressed, SynCAM 1 was detected both as an
apparently uniform high molecular mass species of 100 kDa and
as multiple low molecular mass species of 70–85 kDa. These
changes in the modification of SynCAM 1 were accompanied
by a shift of its predominant expression from hindbrain to fore-
brain. This is consistent with roles of SynCAM 1 in synapse
formation, which progresses during brain development from
the hindbrain to the forebrain. Its binding partner SynCAM 2
also followed a developmental expression increase toward fore-
brain, yet SynCAM 2 was expressed as the same diverse species
at 62–76 kDa throughout. Other N-glycosylated proteins such
as synaptophysin also did not exhibit changes in their modifi-
cation (Fig. 4), consistent with the developmentally indepen-
dent glycosylation of other neuronal membrane proteins.
Modification of SynCAM 2 Ig1 at Asn60 Reduces Adhesion—
To perform a biochemical analysis of SynCAM 2 glycosylation,
we changed the asparagine at position 60 to glutamine, choos-
ing this substitution because it prevents N-glycosylation with-
out altering immunoglobulin folds (44). Consistent with the
conservative nature of this mutation, all Asn 3 Gln glycosyla-
tion mutants used in this study were sorted to the cell surface,
indicating proper folding (see below, Fig. 5, B and D, and sup-
plemental Fig. S3B). Furthermore, the slightly increased bulk of
the glutamine residue in the N60Q SynCAM 2 mutant can be
expected to be easily accommodated in the structures of its
homodimer as well as the heterodimer with SynCAM 1.
To focus our analysis on extracel-
lular interactions, we developed a
GPI-anchored SynCAM 2 construct
that
tethered
its
extracellular
sequence to the outer leaflet of the
plasma membrane. This construct
maintained a complex glycosylation
pattern comparable with that seen
for SynCAM 2 expressed in brain
(Fig. 5A, lanes 1 and 2). The GPI
construct of the SynCAM 2 N60Q
mutant, however, lacked the N-gly-
cosylated wild-type fractions above
55 kDa, consistent with selectively
reduced
glycosylation
(Fig.
5A,
lanes 3 and 4).
We next analyzed the role of
modifications at Asn60 for the ad-
hesive interactions of SynCAM 2
(Fig. 5, B–E). Using a cell overlay
approach with soluble proteins, we
expressed GPI-anchored SynCAM
2 carrying a FLAG epitope in COS7
cells while overlaying the cells with
the soluble extracellular sequence
of SynCAM 2. As described above, the COS7 cell expressed
protein was labeled with anti-FLAG antibodies and the overlaid
soluble protein with protein A. Notably, the absence of a glycan
at amino acid 60 of SynCAM 2 strongly increased its interaction
with the overlaid extracellular sequence of SynCAM 2, more
than doubling its homophilic retention by 125
31% (Fig. 5, B
and C). Similarly, the N60Q mutation increased the hetero-
philic binding of SynCAM 2 to overlaid SynCAM 1 by 61
10%
(Fig. 5, D and E). N-Glycosylation at Asn60 of SynCAM 2 Ig1
therefore restricts its adhesive binding to both SynCAM 2 and
SynCAM 1, possibly because of steric hindrance of N-glycans or
charge repulsion within the binding interface.
Positive Modulation of SynCAM 1 Adhesion by N-Linked
Modification of Its Ig1 Domain—To address whether N-glyco-
sylation within the first Ig domain of SynCAM 1 similarly reg-
ulates its adhesion, we generated a SynCAM 1 N70Q,N104Q
double mutant. These two sites were selected because our
structural models predicted them to flank the SynCAM 1 dimer
interface (supplemental Fig. S2B) and because our mass spec-
trometry data suggested that they were N-glycosylated (Fig. 3).
The SynCAM 1 N70Q,N104Q mutant migrated in immunob-
lots at a lower apparent molecular mass, consistent with
reduced N-glycosylation, and was correctly sorted to the
plasma membrane (supplemental Fig. S3). A SynCAM 1
N70Q,N104Q,N116Q triple mutant lacking all predicted
N-glycosylation sites in the Ig1 domain could not be analyzed
because it was not properly sorted to the cell surface (data not
shown).
Live cell overlay assays with these GPI-anchored constructs
showed that the lack of glycans at the Asn70/Asn104 sites of the
Ig1 domain reduced its homophilic binding to wild-type
SynCAM 1 by 51
16% (Fig. 6, A and B). Similarly, the
N70Q,N104Q mutations decreased the heterophilic binding of
FIGURE 4. SynCAM post-translational modifications are regionally and developmentally regulated in
the brain. The indicated brain regions were dissected from rats at P4, P16, or P30. Equal protein amounts of 30
g were analyzed by immunoblotting using the antibodies shown. SynCAM 1 and 2 exhibited distinct expres-
sion and modification patterns as described in the text. The N-glycosylated synaptic protein synaptophysin
and the scaffolding molecule CASK served as loading controls. The asterisks mark nonspecific bands.
Glycans Modulate SynCAM Adhesion
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SynCAM 1 to wild-type SynCAM 2 by 30
7%. This inhibitory
effect of the SynCAM 1 Ig1 N70Q,N104Q mutation on its
adhesive interactions contrasted with the increased binding
of the SynCAM 2 Ig1 N60Q mutant. We additionally per-
formed affinity chromatographies to analyze the effect of the
SynCAM 1 N70Q,N104Q mutation on the retention of Syn-
CAM 2 from brain (Fig. 6, C and D). Our results show that
the loss of these two N-glycosylation sites reduces hetero-
philic binding to SynCAM 2 by 30
3%, in agreement with
our cell overlay data.
FIGURE 5. SynCAM 2 glycosylation within the Ig1 interface at Asn60
reduces adhesive binding. A, immunoblot analysis of the GPI-anchored
SynCAM 2 extracellular sequence and its N60Q mutant expressed in COS7
cells. Lack of the Asn60 N-glycosylation site resulted in the absence of the
higher molecular mass glycoforms marked by asterisks. Deglycosylation with
PNGase F reduced both wild-type and mutant protein to the same apparent
molecular mass predicted for the unmodified protein. Constructs carried a
FLAG epitope for detection. B, loss of Asn60 glycosylation promotes
homophilic SynCAM 2 binding. COS7 cells expressing GPI-anchored SynCAM
2 or its N60Q mutant carrying an extracellular FLAG epitope (green) were
overlaid with the soluble extracellular domain of SynCAM 2 (red). Cells
expressing FLAG-tagged, GPI-anchored SynCAM 1 Ig2 3 served as a nega-
tive control. Construct expression and SynCAM 2 retention were detected as
described in Fig. 1B. C, quantification of the results in B. The results are
expressed as protein A signal detecting retained SynCAM 2 normalized to the
signalofCOS7surface-expressedSynCAMFLAGconstructs.COS7cellsexpress-
ing GPI-anchored SynCAM 1 Ig2 3 or GFP alone served as negative controls.
Signals are expressed relative to GFP negative control cells. Retention of
SynCAM 2 on cells expressing SynCAM 1 Ig2 3 was lower than on GFP-
expressing cells for unknown reasons (SynCAM 2, n 24 cells; N60Q, n 46;
SynCAM 1 Ig2 3, n 39; GFP 27). ***, p 0.001. D, loss of Asn60 glycosy-
lation in SynCAM 2 promotes its heterophilic binding to SynCAM 1. COS7 cells
expressing FLAG-tagged, GPI-anchored SynCAM 2, or its N60Q mutant
(green) were overlaid with the soluble extracellular domain of SynCAM 1 (red).
Construct expression and SynCAM 1 retention were detected as described in
Fig. 1B. E, quantification of the results in D was performed as described for C
(SynCAM 2, n 25 cells; N60Q, n 40; SynCAM 1 Ig2 3, n 28; GFP 28).
IB, immunoblot.
FIGURE 6. N-Glycosylation of SynCAM 1 at Ig1 sites Asn70/Asn104 pro-
motes adhesive binding. A, absence of Asn70/Asn104 glycosylation weakens
the homo- and heterophilic interactions of SynCAM 1. COS7 cells expressing
GPI-anchored SynCAM 1 or SynCAM 2 carrying an extracellular FLAG epitope
(green) were overlaid with the soluble extracellular sequence of wild-type
SynCAM 1 or its N70Q,N104Q glycosylation mutant (red). Cells expressing
solubleGFPservedasnegativecontrol.Constructexpressionandretentionof
soluble SynCAM 1 were detected as described in Fig. 1B. B, quantification of
the results in A. The results are expressed as fluorescence intensity of retained
SynCAM 1 normalized to the fluorescence intensity of COS7 surface-ex-
pressed SynCAMFLAG constructs. COS7 cells expressing GFP alone served as
negative controls. **, p 0.01; ***, p 0.001. C, lack of SynCAM 1 glycosyla-
tion at Asn70/Asn104 reduces binding to brain SynCAM 2. The extracellular
SynCAM 1 sequence or the N70Q,N104Q mutant were expressed in COS7
cellsasfusionswithIgG1-Fc,andequalamountswereimmobilizedonprotein
A beads. Retention of solubilized rat brain membrane proteins on the immo-
bilized proteins was analyzed by affinity chromatography. SDS eluates
obtained from two parallel affinity bindings are shown. SynCAM 2 signal was
detected by quantified immunoblotting. D, quantification of results obtained
as in C (n 3). E, sialic acid modification of SynCAM 1 promotes its hetero-
philic binding. The SynCAM 1 extracellular sequence was expressed and
immobilized as in C and treated without or with sialidase under native condi-
tions. Retention of membrane proteins from rat brain was analyzed by affinity
chromatography. Affinity matrices were first eluted with 800 mM potassium
acetate and then with SDS. SynCAM 1Ig lacking all three Ig domains served
as negative control, and the GDP dissociation inhibitor GDI and synaptophy-
sin served as controls for nonspecific binding. SynCAM 2 signal was detected
by quantified immunoblotting. FT, flow-through.
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JOURNAL OF BIOLOGICAL CHEMISTRY 34871
These observations raised the possibility that extracellular
SynCAM 1 interactions involve the participation of specific
carbohydrate types. Because SynCAM 1 is modified with sialic
and polysialic acid in brain (20, 45), which we confirmed in our
mass spectrometry analysis of the purified protein, we tested
whether sialic acids on SynCAM 1 contribute to its heterophilic
SynCAM 2 binding. Affinity chromatography of SynCAM 2
extracted from brain was performed on the extracellular
domain of SynCAM 1 that was either fully glycosylated (Fig. 6E,
lanes 1–4) or from which sialic acids had been removed enzy-
matically under native conditions (lanes 5–7). A construct lack-
ing all Ig domains served as a negative control (lanes 8–10). The
removal of sialic acids from SynCAM 1 reduced its retention of
SynCAM 2 by 34
9% (n 3). Although we did not determine
the sites of SynCAM 1 sialylation, this result indicates that
SynCAM 2 adhesion may involve specific interactions with
sialic acids on SynCAM 1. Alternatively, the negative charge of
sialic acids may mediate favorable electrostatic interactions
across the Ig1/Ig1 trans-interface, but the observation that the
SynCAM 1/2 interaction is resistant to high salt conditions
does not support this (Fig. 6E).
N-Glycosylation at the Asn70/Asn104 Sites of SynCAM 1 Ig1
Promotes Synapse Induction—Extending our structural and
biochemical analysis of SynCAM 1 N-glycosylation, we asked
whether modification at the Asn70/Asn104 sites of SynCAM 1
Ig1 alters its trans-synaptic interactions and synaptogenic func-
tion. We expressed the GPI-anchored SynCAM 1 extracellular
domain or the N70Q,N104Q mutant in COS7 cells and
co-cultured them with hippocampal neurons (Fig. 7A). We
then measured two activities, the ability of COS7-expressed
SynCAM 1 to recruit neuronal SynCAM proteins upon contact
and its induction of presynaptic specializations in contacted
neurons (28).
The recruitment of neuronal SynCAM proteins was deter-
mined by quantified immunostaining, taking advantage of the
fact that they can be selectively detected using an antibody that
does not recognize the GPI-anchored SynCAM constructs. As
expected, GPI-SynCAM 1 expressed in COS7 cells efficiently
recruited neuronal SynCAMs to contact sites (Fig. 7, A and B).
The N70Q,N104Q mutant, however, recruited neuronal
SynCAMs 24
11% less, consistent with a weakening of its
trans-synaptic adhesion (Fig. 7B). This weakened recruitment
of neuronal SynCAMs by SynCAM 1 N70Q,N104Q correlated
with its inability to induce presynaptic specializations in neu-
ronal co-cultures (Fig. 7C). The stronger effect of this mutation
on synapse induction than on SynCAM recruitment indicates
that a select threshold of trans-synaptic SynCAM clustering
may have to be met to induce synapses. The N60Q mutant of
SynCAM 2 did not further promote the synaptogenic activity of
SynCAM 2 in this mixed co-culture assay (data not shown),
presumably because the activity of the wild-type protein
already saturated the synapse-forming potential of neurons
under the overexpression conditions of this approach. Such sat-
uration may compromise the detection of positive modulatory
effects. Together, our functional studies demonstrate that
modification of the N-glycosylation sites Asn70/Asn104 of
SynCAM 1 Ig1 increases both trans-synaptic adhesion and its
synaptogenic activity.
DISCUSSION
Our biochemical, crystallographic, mass spectrometry, and
cell biological analyses characterize N-glycosylation as a modi-
fication that modulates SynCAM adhesion. Our results further
indicate roles of SynCAM 1 glycosylation in the regulation of
synapse induction. Four lines of evidence support these conclu-
sions. First, crystallographic results and structural modeling
show that N-glycosylation can occur within and adjacent to the
adhesive Ig1 interface of SynCAM 2 and SynCAM 1, respec-
tively. Second, the glycosylation of these sites within the Ig1
domain differentially affects SynCAM properties, reducing
the adhesion of SynCAM 2 while increasing the binding of
SynCAM 1. Third, the ability to glycosylate these sites increases
not only SynCAM 1 adhesion but also its synaptogenic activity.
Fourth, the post-translational modification of SynCAM 1 is
developmentally regulated in the brain, suggesting functional
roles in vivo.
The post-translational modification of synaptic adhesion
molecules could be an attractive mechanism to regulate them.
Indeed, an inhibitory effect of N-glycosylation on neuroligin 1
binding to neurexin 1 has been previously reported (27). How
can N-glycosylation differentially reduce SynCAM 2 Ig1 bind-
ing and promote SynCAM 1 adhesion? Our crystal structure of
SynCAM 2 Ig 1 and homology model of SynCAM 1/SynCAM 2
Ig1 show that these adhesive trans-dimer interfaces consist
mainly of hydrophobic interactions. The location of Asn60 at
FIGURE 7. Modification of SynCAM 1 at its N-glycosylation sites Asn70/
Asn104 increases trans-synaptic adhesion and synapse induction. A, wild-
type SynCAM 1 recruits neuronal SynCAMs and the presynaptic marker SV2 in
a mixed co-culture assay. COS7 cells co-expressing GFP with the GPI-an-
chored SynCAM 1 extracellular sequence or its N70Q,N104Q mutant were
seeded atop dissociated hippocampal cultures at 7 days in vitro. COS7 cells
expressing GFP alone served as negative control. Co-cultures were analyzed
at 11 days in vitro by immunostaining for neuronally expressed SynCAM pro-
teins (red) and the presynaptic vesicle marker SV2 (blue). GFP marked trans-
fected COS7 cells (green). Wild-type SynCAM 1 recruits and retains neuronal
SynCAMs, and SV2 puncta were detected atop COS7 cells expressing GPI-
anchored SynCAM 1. Cells expressing SynCAM 1 N70Q,N104Q exhibited less
SynCAM and no SV2 recruitment. B, quantification of the SynCAM recruit-
ment shown in A (SynCAM 1, n 30 cells; N70Q,N104Q, n 34; GFP, n 23;
apply also to C). *, p 0.05; **, p 0.01; ***, p 0.001. C, quantification of the
SV2 recruitment shown in A. n.s., not significant.
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VOLUME 285•NUMBER 45•NOVEMBER 5, 2010
this SynCAM 2 dimer interface leaves little room for a bulky
glycan in the crystal structure at this site. Glycans at Asn60 of
SynCAM 2 may therefore weaken the dominant hydrophobic
interactions at the dimer interface and reduce SynCAM 2 adhe-
sion (Fig. 8A).
In contrast, the glycans at Asn70 and Asn104 in SynCAM 1 do
not participate in Ig1 dimer contacts and face away from the
adhesive dimer interface. Why then is the N70Q,N104Q
mutant deficient in adhesive dimer formation? We consider it
possible that N-glycans of SynCAM 1 favor adhesive binding
through limiting the conformational space available to the pro-
tein or by inhibiting nonspecific protein clustering. Both mech-
anisms have been previously proposed for other Ig superfamily
adhesion proteins (46). Specifically, the glycans on Asn70 and
Asn104 may bias or restrict the relative orientations of the
SynCAM 1 Ig1 domain to favor adhesive dimer formation, for
example by limiting the conformational space available to the
Ig1 domain (Fig. 8B).
Our results complement a body of studies characterizing the
role of glycosylation for Ig superfamily members. These studies
have established that carbohydrates can modulate homophilic
adhesion and function, such as shown for L1 and NCAM, and
that specific carbohydrate structures on Ig proteins can regu-
late extracellular interactions as demonstrated for polysialy-
lated NCAM (47–49). Interestingly, a fraction of SynCAM 1
also carries polysialic acids, making it only the second protein
next to NCAM that exhibits this modification in the brain (45).
This polysialylation of SynCAM 1 occurs at the third N-glyco-
sylation site, which was not analyzed in our study, and may
serve as an additional mechanism regulating adhesive strength.
Sialic acids can also specify protein interactions as shown for
the Siglec family of Ig-like lectins (50, 51). However, SynCAMs
do not conform to conserved sequence motif in Siglecs (52) and
appear unlikely to belong to this protein family. The potential
roles of carbohydrates in binding specificity and carbohydrate-
carbohydrate interactions (53) can now be addressed in future
studies of adhesive SynCAM recognition.
The significant developmental changes in the post-transla-
tional modification of SynCAM 1 indicate that specific, pres-
ently unknown glycosyltransferases modify it in the brain. In
contrast, only a minor fraction of SynCAM 2 may undergo
regulated carbohydrate modification. Functionally, this differ-
ential glycosylation could modulate SynCAM interactions
between neuronal populations, refining the potential for adhe-
sive coding provided by the distinct SynCAM gene expression
patterns (20, 21). The modification of SynCAMs with glycans
may not only adjust their synaptic adhesive strength during
brain development. Glycosylation could also change the struc-
tural organization of SynCAM complexes in the synaptic cleft,
analogous to the role of N-glycans in patterning the trans-ad-
hesion arrays formed by L1 (54). Future studies will determine
whether glycans on residues other than those analyzed here
further modulate SynCAM structure and function, including
the O-glycans at the stalk of the SynCAM 1 extracellular
domain (19).
With respect to the roles of modulated adhesion, it is inter-
esting to note that the glycosylation sites Asn60 of SynCAM 2
and Asn70 of SynCAM 1 are evolutionarily conserved between
human and murine orthologs and that the Asn104 site of mam-
malian SynCAM 1 is even present in the avian and fish
orthologs (19). This indicates that the ability to modify these
sites in SynCAM Ig1 domains is functionally relevant.
Together, N-glycosylation alters the adhesive interactions and
synapse-inducing functions of SynCAMs, demonstrating that
this modification modulates trans-synaptic SynCAM interac-
tions. Our findings support the notion that glycosylation plays
important roles in synaptic surface interactions (55, 56).
Acknowledgments—We thank the members of the Biederer and Modis
laboratories for helpful discussions. We also thank Edward Voss
(W. M. Keck Foundation Biotechnology Resource Laboratory) and
Michael Easterling (Bruker Daltonics, Inc.) for running some of the
samples on the FT-ICR MS. We thank Howard Robinson, Annie He´r-
oux, and other staff at the X25 and X29A beamlines of the National
Synchrotron Light Source at Brookhaven National Laboratory.
FIGURE 8. Model of differential SynCAM modulation by N-glycosylation
of the first Ig domain. A, N-glycans may reduce SynCAM 2 adhesion through
steric hindrance within the Ig1 binding interface. B, in contrast, N-glycans
facing away from the SynCAM 1 Ig1 domain may restrict its conformational
freedom and position it toward binding. Note that we do not exclude addi-
tional interactions between SynCAM Ig domains. Dark gray, SynCAM 1; light
gray, SynCAM 2.
Glycans Modulate SynCAM Adhesion
NOVEMBER 5, 2010•VOLUME 285•NUMBER 45
JOURNAL OF BIOLOGICAL CHEMISTRY 34873
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Glycans Modulate SynCAM Adhesion
34874
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 45•NOVEMBER 5, 2010
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3M47
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Crystal structure of the mutant I218A of orotidine 5'-monophosphate decarboxylase from Methanobacterium thermoautotrophicum
|
Conformational Changes in Orotidine 5′-Monophosphate
Decarboxylase: “Remote” Residues that Stabilize the Active
Conformation†
B. McKay Wood‡, Tina L. Amyes§, Alexander A. Fedorov∥, Elena V. Fedorov∥, Andrew
Shabila‡, Steven C. Almo∥, John P. Richard§, and John A. Gerlt*,‡
‡Departments of Biochemistry and Chemistry, University of Illinois at Urbana-Champaign
§Department of Chemistry, University at Buffalo, Buffalo, NY 14260
∥Department of Biochemistry, Albert Einstein College of Medicine, Bronx, NY 10461
Abstract
The structural factors responsible for the extraordinary rate enhancement (~1017) of the reaction
catalyzed by orotidine 5′-monophosphate decarboxylase (OMPDC) have not been defined.
Catalysis requires a conformational change that closes an active site loop and “clamps” the orotate
base proximal to hydrogen-bonded networks that destabilize the substrate and stabilize the
intermediate. In the OMPDC from Methanobacter thermoautotrophicus, a “remote” structurally
conserved cluster of hydrophobic residues that includes Val 182 in the active site loop is
assembled in the closed, catalytically active conformation. Substitution of these residues with Ala
decreases kcat/Km with a minimal effect on kcat, providing evidence that the cluster stabilizes the
closed conformation. The intrinsic binding energies of the 5′-phosphate group of OMP for the
mutant enzymes are similar to that for the wild type, supporting this conclusion.
Orotidine 5′-monophosphate decarboxylase (OMPDC1) is one of Nature’s best catalysts: the
reaction occurs with a rate enhancement of ~1017 and a proficiency of ~1023 M (2). The
reaction coordinate includes a vinyl carbanion intermediate (Scheme 1) (3, 4).
Structures of OMPDCs complexed with UMP or 6-hydroxyUMP, a n intermediate analog
(5-9), reveal hydrogen-bonded networks proximal to 1) C6 of the pyrimidine (Asp 70-Lys
72-Asp 75 in the OMPDC from Methanobacter thermoautotrophicus; MtOMPDC); and 2)
O2, N3, and O4 of the base (Ser 127-Gln 185). These destabilize the substrate (9) and
stabilize the intermediate, although the structural strategy for the latter is uncertain.
The 5′-phosphate group binds in a conserved motif at the ends of the seventh and eighth β-
strands of the (β/α)8-barrel structure, with interactions to backbone NH groups as well as the
guanidinium group of a conserved Arg (Arg 203 in MtOMPDC). These interactions are
important for catalysis: 1) kcat/Km for OMP exceeds that for orotidine by a factor of ~1011
for the OMPDC from Saccharomyces cerevisiae (ScOMPDC) (10); 2) kcat/Km for OMP
†This work was supported by NIH Grants GM039754 to J.P.R. and GM065155 to J.A.G.
*To whom correspondence should be addressed: J.A.G.: telephone, (217) 244-7414; fax: (217) 244-6538; j-gerlt@uiuc.edu..
SUPPORTING INFORMATION AVAILABLE
Descriptions of the experimental procedures. This material is available free of charge via the Internet at http://pubs.acs.org.
1Abbreviations: OMP, orotidine 5′-monophosphate; OMPDC, OMP decarboxylase; MtOMPDC, OMPDC from Methanobacter
thermoautotrophicus; ScOMPDC, OMPDC from Saccharomyces cerevisiae; EO, 1-(β-D-erythrofuranosyl)-orotic acid; HPi,
phosphate dianion; IBE, intrinsic binding energy.
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 December 26.
Published in final edited form as:
Biochemistry. 2010 May 4; 49(17): 3514–3516. doi:10.1021/bi100443a.
NIH-PA Author Manuscript
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exceeds that for 1-(β-D-erythrofuranosyl)orotic acid (EO; 5′-truncated OMP analog) by
factors of 5.2 × 108 and 3.6 × 108 for ScOMPDC and MtOMPDC, respectively (11, 12); and
3 ) phosphite dianion ( H Pi) activates decarboxylation of EO by factors of 5.6 × 105 M−1
and 2.9 × 105 M−1 for ScOMPDC and MtOMPDC, respectively (11, 12). The “intrinsic
binding energy” [IBE; (13)] of the 5′-phosphate/HPi 1) increases the affinity for the
substrate (e.g., OMP vs. orotidine/EO); and 2) enables decarboxylation by juxtaposition of
the substrate with the active site hydrogen-bonded networks (substrate destabilization and
intermediate stabilization). How the IBE promotes catalysis is unknown but required to
understand the structural basis for the rate enhancement.
A loop located at the end of the seventh β-strand closes over the active site when OMP binds
(Figure 1). Although the active site loops differ in both length and sequence in divergent
OMPDCs (12), each includes a spatially conserved Gln (Gln 185 in MtOMPDC) hydrogen-
bonded to both the 5′-phosphate and O2 of the pyrimidine as well as a conserved Ser at the
end of the fifth β-strand that also is hydrogen-bonded to N3 of the pyrimidine (Ser 127). We
characterized the importance of these “clamp” residues in ScOMPDC (Gln 215-Ser 154)
using EO/HPi and confirmed that the 5′-phosphate/HPi “switch” is required to activate the
enzyme (14).
The structures of divergent OMPDCs reveal that OMP binding is always accompanied by a
conformational change (Figure 1). The most obvious component is closure of the active site
loop. However, the (β/α)8-barrel structure can be divided into two domains, one formed
from the second, third, fourth, and fifth β-strands (where the hydrogen-bonded Asp 70-Lys
72-Asp 75 motif and Ser 127 are located) and the second from the sixth, seventh, eighth, and
first β-strands (where the phosphate binding motif and the active site loop, including Gln
185, are located) (15). OMP binding reorients the domains, with the latter domain moving
toward the former, enforcing the orotate carboxylate group to be juxtaposed vis a vis Asp
70- Lys 72-Asp 75 and, also, allowing formation of the Ser 127-Gln 185 “clamp”. Thus, the
transition between the open and closed conformations is more complicated than “simple”
hinge motion of the loop on the rigid framework of the (β/α)8-barrel structure. In this Report
we identify “remote” residues involved in this conformational change and quantitate their
importance in promoting and stabilizing the catalytically competent form of the enzyme.
The active site loop of MtOMPDC, P180-G181-V182-G183-A184-Q185-G186-G187-D188,
is disordered in the absence of substrate but ordered and closed in its presence (Figure 1). In
the liganded structure two residues in the loop make contacts with the (β/α)8-barrel scaffold:
1) Gln 185 is hydrogen-bonded to Ser 127 (vide infra); and 2) Val 182 is embedded in a
hydrophobic cluster also formed by Ile 199, Val 201, and Ile 218. The conservation of this
hydrophobic cluster in all OMPDCs suggests its importance in a common catalytic strategy.
We probed this strategy by mutagenesis of these hydrophobic residues.
Ala substitutions for residues in the hydrophobic cluster cause substantial decreases in kcat/
Km for decarboxylation of OMP but have little effect on kcat (Table 1). We determined high
resolution X-ray structures (≤ 1.4 Å) for each mutant in the presence of 6-hydroxyUMP
(Figure 2). The liganded structures superimpose well with that of wild type, with only small
differences observed at the sites of the substitutions (panel A). The active sites are identical
to that of wild type (panel B), explaining the minimal impact on kcat.
The mutated residues are remote from the active site (Figures 1 and 2). Thus, the effects of
the substitutions on kcat/Km cannot be explained by altered direct interactions with the
substrate. Instead, the effects can be explained by decreased stabilities of the closed
conformation in which the substrate is destabilized (9) and the anionic intermediate is
stabilized. A consistent model (Scheme 2) is that an equilibrium of open (Eo) and closed
Wood et al.
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(Ec) conformations exists in the absence or presence of substrate (KC’ or KC; KC’ << 1). The
substitutions destabilize Ec (decrease KC and KC ’), so kcat/Km is decreased (the energy
difference between Eo + S and [Ec•S]‡ is increased). However, the invariance of kcat
establishes that the substitutions do not alter the reactivitiy of Ec•S (the energy difference
between Ec•S and [Ec•S]‡). The energy to form Ec•S from Eo + S is achieved from 1)
interactions of the 5′-phosphate group with its binding motif (IBE; vide infra), and 2) an
increased concentration of OMP to form Ec•S (KC/KS or, equivalently, KC’/KS’, although
the former is expected to be the relevant pathway).
We also used the two part EO•HPi substrate. The values of kcat/Km for EO are decreased
relative to that for wild type (Table 1); these can be explained by decreased populations of
Ec (Scheme 2), assuming that EO, without a 5′-phosphate group, is unable to promote the
transition from Eo to Ec. The values of kcat/Km for OMP and kcat/Km for EO allow
calculation of the IBE for the 5′-phosphate group of OMP (Table 1).
HPi activates the mutants as judged by the values of the third-order rate constant, (kcat/
Km)EO•HPi/Kd. The equivalent changes (ΔΔG‡) in kcat/Km for both OMP and EO and the
third-order rate constant indicate that all three measure the effects of the substitutions on the
values of KC.
The values of the IBEs for the 5′-phosphate group for the mutants are the same as that for
the wild type, establishing that decarboxylation in the Ec•S complex occurs with equivalent
amounts of ground state destabilization (9) and transition state stabilization (as also reflected
by the invariant values of kcat). The IBEs provide further support for the role of the “remote”
hydrophobic cluster in stabilizing Ec relative to Eo but do not directly participating in
catalysis.
Our experiments implicate a structurally conserved hydrophobic cluster, Val 182, Ile 199,
Val 201, and Ile 218 in MtOMPDC, in stabilizing the closed conformation required for
catalysis. Its identification provides evidence that structural elements distal from the active
site, in addition to the proximal active site loop that closes to “clamp” the substrate, are
required for OMPDC’s extraordinary catalytic efficiency and proficiency.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
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Figure 1.
Superposition of the active site of wild type MtOMPDC in the absence (cyan; disordered
loop depicted with the dotted line ) and presence ( grey ) of 6-hydroxyUMP. The carbons of
6-hydroxyUMP are highlighted in yellow; the carbons of Val 182 and Arg 203 in the
liganded structure are highlighted in orange.
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Figure 2.
Superpositions of the 6-hydroxyUMP-liganded structures of wild type MtOMPDC and the
single mutants in the hydrophobic cluster. Panel A, hydrophobic cluster; panel B, active
sites.
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Scheme 1.
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Scheme 2.
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Wood et al.
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Table 1
Kinetic Constants for OMP, EO and EO/HPi and Intrinsic Binding Energies of the 5′-Phosphate Group of OMP at pH 7.1 and 25 °C.
MtOMPDC
kcat
OMP
s−1
kcat/Km
OMP
M−1 s−1
ΔΔG‡
kcal/mola
kcat/Km
EO
M−1 s−1
ΔΔG‡
kcal/mola
(kcat/Km)/K
D
EO•HPib
M−2 s−1
ΔΔG‡
kcal/mola
5′-Phosphate
IBEc
kcal/mol
Wild type
4.6
2.9 × 106
8.7 × 10−3
2500
11.6 d
V182A
3.4
1.4 × 105
1.8
1.3 × 10−3
1.1
190
1.5
10.9
I199A
3.9
9.1 × 105
0.7
1.9 × 10−3
0.9
980
0.6
11.8
V201A
4.0
9.5 × 105
0.7
3.1 × 10−3
0.6
690
0.8
11.5
I218A
3.3
2.8 × 105
1.4
2.3 × 10−3
0.8
340
1.2
11.0
V182A/I199A
3.1
4.9 × 104
2.4
3.9 × 10−4
1.8
81
2.0
11.0
V182A/V201A
2.5
4.9 × 104
2.4
5.0 × 10−4
1.7
30
2.6
10.9
aCalculated from the ratio of the second-order or third-order rate constants for the wild type and mutant enzyme.
bThird-order rate constant for reaction of EO/HPi.
cTransition state stabilization by the 5′-phosphate group of OMP, calculated from the ratio of the values of kcat/Km for OMP and EO. IBEs for phosphite dianion can be calculated from the ratio of (kcat/
Km)/KD for EOHPi and kcat/Km for EO.
Biochemistry. Author manuscript; available in PMC 2011 December 26.
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3M4A
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Crystal structure of a bacterial topoisomerase IB in complex with DNA reveals a secondary DNA binding site
|
Crystal structure of a bacterial topoisomerase IB in complex with
DNA reveals a secondary DNA binding site
Asmita Patel1, Lyudmila Yakovleva2, Stewart Shuman2,3, and Alfonso Mondragón1,3
1 Department of Biochemistry, Molecular Biology and Cell Biology, Northwestern University, 2205
Tech Drive, Evanston, Illinois 60208
2 Molecular Biology Program, Sloan-Kettering Institute, 1275 York Avenue, New York, New York
10065
Summary
Type IB DNA topoisomerases (TopIB) are enzymes that relax supercoils by cleaving and resealing
one strand of duplex DNA within a protein clamp that embraces a DNA segment. A longstanding
conundrum concerns the capacity of TopIB enzymes to stabilize intramolecular duplex DNA
crossovers and, in the case of poxvirus TopIB, form protein-DNA synaptic filaments. Here we report
a structure of D. radiodurans TopIB in complex with a 12-bp duplex DNA that demonstrates a
secondary DNA binding site located on the C-terminal domain. It comprises a distinctive interface
with one strand of the DNA duplex and is conserved in all TopIB enzymes. Modeling of a TopIB
with both DNA sites suggests that the secondary site could account for DNA crossover binding,
nucleation of DNA synapsis, and generation of a filamentous plectoneme. In support of this,
mutations of the secondary site eliminate synaptic plectoneme formation without affecting DNA
cleavage or supercoil relaxation.
Introduction
Type IB DNA topoisomerases (TopIB) are encoded in the genomes of all eukarya, several
eukaryal viruses (poxviruses and mimivirus; Benarroch et al., 2006), many bacteria (Krogh
and Shuman, 2002), and several archaea (Forterre et al., 2007). They play important roles in
relaxing supercoils generated during DNA replication and transcription. TopIB enzymes
accomplish this task by repeatedly breaking and rejoining one strand of the DNA duplex
through a covalent DNA-(3′-phosphotyrosyl)-enzyme intermediate (Corbett and Berger,
2004). Within the covalent TopIB–DNA complex, the noncovalently held 5′-OH DNA segment
swivels about the protein-DNA nick before being religated to the 3′-phosphate of the covalently
held strand. The number of supercoils removed by TopIB per cleavage-religation cycle follows
an exponential distribution that depends on the torque stored in the supercoiled DNA and
friction at the protein-DNA interface during the swivel (Koster et al., 2005).
Crystal structures of DNA-bound cellular and poxvirus TopIB enzymes captured at sequential
steps along the reaction pathway (pre-cleavage, transition-state, and post-cleavage covalent
complex) have fully illuminated the DNA-protein interactions and reaction chemistry (Davies
3Corresponding authors: A.M. Phone: 847-491-7726, Fax: 847-467-6489, a-mondragon@northwestern.edu. S.S. Phone: 212-639-7145,
Fax: 212-772-8410, s-shuman@ski.mskcc.org.
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Author Manuscript
Structure. Author manuscript; available in PMC 2011 June 9.
Published in final edited form as:
Structure. 2010 June 9; 18(6): 725–733. doi:10.1016/j.str.2010.03.007.
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et al., 2006; Perry et al., 2006, 2010; Redinbo et al., 1998). These structures, together with
biochemical studies (Krogh and Shuman, 2000; Tian et al., 2005), revealed how nucleophilic
attack of the active-site tyrosine hydroxyl on the DNA phosphodiester bond is catalyzed by
two arginines, a lysine and a histidine that stabilize the pentacoordinate transition-state and
expel the 5′-OH leaving strand. The crystal structures also underscored how all TopIB enzymes
envelop the duplex DNA cleavage site by forming a C-shaped protein clamp, wherein a C-
terminal catalytic domain engages the DNA minor groove at and surrounding the scissile
phosphodiester while an N-terminal domain module engages the DNA major groove on the
face of the duplex opposite the cleavage site.
Comparison of the structures of DNA-bound poxvirus TopIB (Perry et al., 2006) and the free
apoenzyme (Cheng et al., 1998) highlighted that the active site is not preassembled prior to
DNA binding. Indeed, in the poxvirus apoenzyme, three of the five catalytic residues (the lysine
and arginine general acids and the tyrosine nucleophile) are either disordered or out of position
to perform transesterification chemistry. The formation of a catalytically competent active site
entails multiple conformational switches and disordered-to-ordered transitions within the
catalytic domain that are triggered by recognition of specific nucleobases and backbone
phosphates of the consensus 5′-CCCTT↓/3′-GGGAA cleavage site for poxvirus TopIB (Perry
et al., 2006; Tian et al., 2004a; Tian et al., 2004b; Yakovleva et al., 2006).
Many bacterial species encode a TopIB that resembles the poxvirus and mimivirus enzymes
with respect to their small size, primary structures, and bipartite domain organization (Krogh
and Shuman, 2002). It is speculated that horizontal transfer of TopIB genes among bacteria
and eukaryal viruses occurred during their cohabitation in a unicellular eukaryal host, e.g.,
amoebae (Benarroch et al., 2006). The bacterial TopIB clade is exemplified by Deinococcus
radiodurans TopIB (DraTopIB), the only member that has been characterized biochemically
and structurally (Krogh and Shuman, 2002; Patel et al., 2006). Although the crystal structures
and active sites of poxvirus TopIB and DraTopIB are quite similar (more so to each other than
to the much larger eukaryal cellular TopIB enzymes), three features of DraTopIB stand out in
comparison to the poxvirus TopIB: (i) DraTopIB does not transesterify at the poxvirus 5′-
CCCTT↓ cleavage site, (ii) the five catalytic amino acids of DraTopIB are pre-assembled in
the apoenzyme crystal structure, and (iii) a segment of the DraTopIB catalytic domain flanking
a catalytic arginine that is disordered in the apoenzyme crystal structure corresponds to the
“specificity helix” of poxvirus TopIB that is critical for DNA site recognition and cleavage
(Patel et al., 2006; Perry et al., 2006; Yakovleva et al., 2008).
It has long been suspected that the catalytic DNA binding mode seen in the available TopIB-
DNA crystal structures might not be the only means by which TopIB interacts with DNA. In
a pioneering study, Zechiedrich and Osheroff observed by electron microscopy that
mammalian TopIB prefers to bind to relaxed circular and linear plasmid DNA molecules at
the nodes created by the crossing of two duplex helices (Zechiedrich and Osheroff, 1990). They
suggested that TopIB might initially bind to one DNA segment and then capture a second DNA
segment at a distant site in the same plasmid molecule. Later, Madden et al. reported that the
Y723F active site mutant of human TopIB binds preferentially to positively or negatively
supercoiled plasmid DNA compared to relaxed DNA molecules (Madden et al., 1995).
Crossover recognition provides a “topology sensor” and a potential means to direct TopIB
action to plectonemic DNAs. Electron microscopy has also been used to visualize complexes
formed by poxvirus TopIB on plasmid DNAs (Shuman et al., 1997). The poxvirus TopIB
formed intramolecular loop structures in which non-contiguous DNA segments were synapsed
at protein-containing nodes or within filamentous protein stems. The formation of filaments
along the DNA suggested that poxvirus TopIB binds DNA cooperatively. At high TopIB
concentrations, the DNA appeared to be “zipped up” within the protein filaments such that the
duplex was folded back on itself. Formation of loops and filaments was also observed with an
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active site mutant, TopIB-Phe274. The zipped-up poxvirus TopIB-DNA complexes formed on
relaxed DNA were shown to be plectonemic supercoils, in which the two duplexes
encompassed by the protein filaments are interwound in a right-handed helix (Shuman et al.,
1997). Thus, TopIB binding to DNA directly imposes a higher order DNA structure. Further
insights to the DNA binding properties of poxvirus TopIB were gained by applying atomic
force microscopy (AFM) to the problem (Moreno-Herrero et al., 2005). AFM verified that
poxvirus TopIB formed nodes and filaments on linear or nicked-circular DNAs by
intramolecular synapsis of two distant DNA segments. Measuring the filament length as a
function of TopIB concentration showed that synapsis is a highly cooperative process. The
congruence of the EM and AFM studies suggested that TopIB-mediated DNA synapsis might
contribute to organization of the 200-kbp vaccinia genome into a higher order structure
conducive to transcription within virus cores.
A key question is how TopIB bridges distant DNA sites. Is it via protein-protein interactions
between two DNA-bound TopIB molecules? Or can a single molecule of TopIB bind
simultaneously to two DNA segments? These two simple physical mechanisms to account for
synapsis by the poxvirus TopIB are depicted in Fig. 1 as models A and B, respectively. In
model A, protein-protein interactions between TopIB molecules provide the “glue” for
synapsis of two TopIB-DNA filaments. (The model arbitrarily depicts the interaction between
the C-terminal catalytic domains of DNA-bound protomers; it could just as well entail N-
domain/N-domain interactions or N-domain/C-domain contacts.) In model B, the synapsed
DNA duplex is captured at a putative secondary DNA binding site on the TopIB protomer.
Similarly, the binding of TopIB to DNA crossovers can be explained by either TopIB-TopIB
interactions or simultaneous occupancy of two DNA binding sites on TopIB.
Here we report the crystal structure of the bacterial DraTopIB enzyme in a complex with DNA.
The structure demonstrates a secondary DNA binding site located on the surface of the C-
terminal domain. The secondary DNA site is ~30 Å from the catalytic DNA site and comprises
an extensive network of direct and water-mediated hydrogen bonds from the enzyme to one
strand of the DNA duplex. The secondary site appears to be conserved in the poxvirus and
eukaryal cellular TopIB enzymes. A model of the poxvirus TopIB enzyme with both DNA
sites filled suggests how second site capture might account for DNA crossover binding,
nucleation of DNA synapsis, and plectonemic supercoiling within the synaptic filament. We
provide biochemical evidence in support of this model by showing that mutations in the
putative secondary DNA binding site of poxvirus TopIB affect the generation of plectonemic
supercoils, but not supercoil relaxation.
Results and Discussion
Structure of a DraTopIB-DNA complex
Crystals of DraTopIB were grown in the presence of a 12-bp DNA duplex (see Materials and
Methods) that is not a substrate for cleavage by the enzyme. The crystals belong to space group
C2, unlike the DraTopIB apoenzyme (Patel et al., 2006), which crystallized in space group
P21. The structure of the DraTopIB-DNA complex was solved by Molecular Replacement
using the structure of the DraTopIB apoenzyme catalytic domain as a search molecule and
refined at 1.65 Å resolution to Rwork/Rfree of 0.193/0.222 (Table I). In the crystal lattice, the
short DNA duplexes are stacked end-to-end to form a pseudo-continuous helix to which
DraTopIB is bound. In prior TopIB-DNA structures, the DNA fits into an interdomain cleft,
the domains form a circumferential clamp around the duplex, and the active site residues in
the C-domain directly coordinate the scissile phosphodiester (Fig. 2A). By contrast, in the
DraTopIB-DNA complex, the N- and C-domains are splayed apart (not shown) and the
catalytic DNA binding site is unoccupied (Fig. 2B). The N-domain (aa 1–90) is partially
disordered and makes no contacts to the 12-bp DNA ligand. The C-domain (aa 91–346) is fully
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ordered and the DNA is docked on the basal surface far from the active site (Fig. 2B). The C-
domains of the bacterial and viral TopIB enzymes comprise two globular lobes (lobes 1 and
2) (Fig. 2). Alignment of the C-domains of the poxvirus TopIB and DraTopIB DNA complexes
reveals that the DraTopIB-DNA structure heralds the existence and position of a bona fide
secondary DNA-binding site on TopIB. The secondary DNA binding site is located within lobe
1 (Fig. 2B).
DNA binding triggers folding of the “specificity helix”
Comparison of the C-domains of the DraTopIB apoenzyme and the DNA-bound DraTopIB
reveals both subtle and profound changes correlated with DNA binding. The C-domains of the
two structures superimpose with a rmsd of 1.07 Å for all main chain atoms. However, when
the isolated lobes 1 and 2 are superimposed separately, the rmsd values are only 0.87 Å and
0.76 Å, respectively. Thus, whereas the individual lobes are nearly identical in the apoenzyme
and DNA-bound DraTopIB, the DNA elicited a subtle reorientation of the lobes with respect
to each other. The five catalytic amino acids (Arg137, Lys174, Arg239, Asn280 and Tyr289)
are preassembled into an active site in the DNA complex (Fig. 2B) and are located in the same
positions as in the apoenzyme. Yet, none of the five catalytic residues make contact with DNA
in the crystal lattice.
An important insight from the poxvirus TopIB-DNA cocrystal (Perry et al., 2006) was the
identification of a “specificity helix” (131FGKMKYLKENETVG144) that binds the DNA
target site in the major groove (Fig. 2B) and makes atomic contacts to nucleobases and
phosphate oxygens that are important for cleavage site recognition and DNA transesterification
(Yakovleva et al., 2008). This specificity helix is conserved among poxvirus and mimivirus
TopIB enzymes and is a distinctive secondary structure feature of the viral/bacterial TopIB
clade that is absent in human TopIB (Redinbo et al., 1998). The specificity helix of poxvirus
TopIB is protease-sensitive and disordered in the apoenzyme, but adopts a defined secondary
structure and becomes protease-resistant when the poxvirus TopIB is in the DNA-bound state
(Cheng et al., 1998; Perry et al., 2006; Sekiguchi and Shuman, 1995). Tight docking of the
specificity helix in the major groove 5′ of the scissile phosphate aids in placing the catalytic
Arg130 residue in the active site.
The DraTopIB equivalent of the poxvirus specificity helix is 138VGSDIYARQHKTYG151.
Structure probing of free DraTopIB by limited proteolysis delineated a single trypsin-sensitive
site within this segment between Arg145 and Gln146 (Krogh and Shuman, 2002). Moreover,
the 139GSDIYARQHK148 peptide is disordered in the crystal structure of apo-DraTopIB (Patel
et al., 2006). By contrast, we find presently that in the DraTopIB-DNA cocrystal, the previously
disordered peptide segment forms a well-ordered α-helix that mimics the specificity helix of
poxvirus TopIB (Fig. 2B). In the poxvirus TopIB, the specificity helix penetrates deeply into
the DNA major groove, where it makes multiple side chain and main-chain contacts to the
DNA phosphates and bases (Fig. S1A). The equivalent α-helix in DraTopIB makes a single
(nonspecific) contact to a backbone phosphate in a symmetry-related 12-mer DNA and one
contact to a nucleobase. We attribute these limited contacts to crystal packing (Fig. S1B). The
symmetry related 12-mer DNA is not situated in the active site.
The structures of DraTopIB and poxvirus TopIB suggest that several mechanisms exist to
trigger the folding of the specificity helix. In poxvirus TopIB, this occurs as a direct response
to binding of the cleavage recognition sequence in the catalytic DNA site. In the case of
DraTopIB, the equivalent conformation switch occurs either as: (i) a nonspecific response to
duplex DNA that does not trigger catalysis, or (ii) an indirect, perhaps allosteric, response to
occupancy of the secondary DNA binding site. Because the catalytic Arg137 is already poised
in the active site in the DraTopIB apoenzyme, we presume that the induced folding of the
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specificity helix is critical for recognition of the target site(s) for DNA transesterification,
which (though clearly different from that of the poxvirus TopIB) are presently uncharted.
Architecture of the secondary DNA binding site
The secondary DNA binding site is located entirely within lobe 1 of the C-domain of DraTopIB.
The DNA interface comprises a helix-loop-helix (aa 106–129) that contacts one strand of the
12mer duplex from the minor groove side and another α-helix (aa 186–194) that contacts the
same strand from the major groove side (Fig. 3A). The component peptide elements form a
groove on the basal surface of the C-domain into which only one strand of the DNA duplex
fits. Most of the atomic contacts entail H-bond donation from protein side chain (Thr110,
Lys122, Arg129, Thr187, Asn191, Lys194) or main chain (Gly117) atoms to three consecutive
DNA phosphates of one DNA strand (Fig. 3B). In addition, Arg116 and Arg186 contact
different guanine bases in the same DNA strand (Fig. 3A,B). The “footprint” of the secondary
DNA site covers 4 nucleotides (Fig. 3B) and consists of 11 direct hydrogen bonds from protein
to DNA (Fig. 3A). In addition, there are at least 8 water-mediated hydrogen bonds from protein
to DNA (not shown). The interface area between the protein and DNA is 584 Å2, significantly
larger than the 380 Å2 for the interface area between DNA and the specificity helix. The
interface area is small in comparison with that of the poxvirus Top IB with DNA (1517 Å2),
but this is to be expected for a non-sequence-specific secondary site with a relatively small
footprint. Thus, we regard the DNA interactions at the secondary site of DraTopIB as too
extensive to ascribe to incidental lattice contacts, unlike the case of the DraTopIB specificity
helix discussed above that makes few contacts with the DNA.
The significance of the secondary DNA binding site defined by the new DraTopIB structure
is underscored by its conservation in the poxvirus and eukaryal TopIB. Superposition of the
viral and human enzymes on DraTopIB shows that the component α-helices and loops of the
secondary DNA site are preserved in all three TopIBs, affording a similar groove to
accommodate one strand of duplex DNA (Fig. 4). Also, many of the basic and hydrophilic side
chains that contact the DNA in DraTopIB (Fig. 4A) are either conserved in the poxvirus and
human enzymes or substituted by a related side chain in a similar spatial position (Fig. 4B and
C). The available structural and phylogenetic information suggest that the secondary DNA site
is not a unique feature of DraTopIB. Whether other TopIB enzymes can bind DNA using this
site remains to be determined (see below). A recent study implicates a cluster of four lysines
on the surface of human TopIB as contributory to the preferential binding of TopIB to
supercoiled DNA (Yang et al., 2009); these lysines (underlined in Fig. 4C) are located within
the putative human TopIB equivalent of the DraTopIB secondary DNA binding site identified
presently and three of them are conserved in poxviral and/or bacterial TopIB. These
observations lend further credence to a secondary DNA site common to type IB enzymes.
To gain a sense of whether the secondary site might be important for DNA relaxation and/or
DNA site recognition by poxvirus TopIB, we engineered three alanine-cluster mutants that
eliminated the putative equivalents of the secondary DNA-binding side chains. One cluster
(N103A-K104A-K107A-K108A-Y115A; or 5xAla) targeted five amino acids in the helix-
loop-helix that engages from the minor groove; the second cluster (R181A-K184A-K188A; or
3xAla) targeted three residues in the α-helix that binds from the major groove. A third cluster
(N103A-K104A-K107A-K108A-Y115A-R181A-K184A-K188A; or 8xAla) simultaneously
changed all eight amino acids to alanine. The mutated poxvirus TopIBs were produced in E.
coli with a C-terminal His6-tag and purified from soluble bacterial lysates by Ni-agarose and
phosphocellulose chromatography (Yakovleva et al., 2008) in parallel with wild-type TopIB.
The rate of relaxation of supercoiled plasmid DNA by the 3xAla cluster mutant was
indistinguishable from the wild-type TopIB, whereas the 5xAla and 8xAla cluster mutants
relaxed at about half and one-third of the wild-type rate, respectively (Fig. 5A). The rates of
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single-turnover cleavage of a short duplex DNA “suicide substrate” containing a single 5′-
CCCTT↓ cleavage site for poxvirus TopIB were identical for the wild-type, 3xAla, 5xAla, and
8xAla cluster mutant proteins (not shown). We surmise that individual constituents or grouped
subsets of the imputed secondary DNA site of poxvirus are not crucial for topoisomerase
activity under the in vitro conditions we employ routinely to assess mutational effects.
Implications for TopIB-mediated DNA synapsis
The present discovery of a secondary DNA binding site on DraTopIB lends support for Model
B in Figure 1, in which DNA synapsis by poxvirus TopIB reflects capture of a second DNA
duplex by DNA-bound proteins. Instructive clues to the process of synapsis were gained by
docking the 12-mer DNA duplex bound to DraTopIB into the structure of the poxvirus TopIB
bound covalently to its CCCTT target site in duplex DNA as a vanadate transition state mimetic
(Perry et al., 2010). This was performed by superimposing the DNA-bound TopIB structures
and then subtracting the DraTopIB protein. The resulting model of poxvirus TopIB with the
primary and secondary DNA sites occupied in shown in Fig. 6, in three different orientations.
The image in Fig. 6C highlights the C-shaped clamp formed by the N- and C-domains around
the DNA duplex engaged in the catalytic site. Fig. 6B highlights the groove between the α-
helices of the secondary DNA site at the base of the C-domain into which one of the DNA
strands fits. The primary and secondary DNA duplexes are oriented similarly, but separated
by ~30 Å. The view in Fig. 6A illustrates that the DNAs in the two sites do not have a parallel
trajectory; rather their paths cross in this and other planes. Indeed, the angular difference
between the paths of the two helices could explain the plectonemic winding of one duplex
abound the other that occurs within the synaptic filaments formed by poxvirus TopIB on
initially relaxed circular DNAs (Shuman et al., 1997). A similar model built using human
TopIB (not shown) also shows the possible formation of a complex with two DNA binding
sites, despite the much larger size of the eukaryotic enzyme and the presence of additional
protein domains. This binding mode could explain the way eukaryotic TopIB binds to nodes
in positively or negatively supercoiled DNA.
The secondary binding site is required for synaptic plectoneme formation by poxvirus TopIB
The structure of DraTopIB in complex with DNA at a secondary binding site provides a
blueprint for functional probing of the basis for DNA condensation and synapsis by poxvirus
TopIB, especially the plausibility of the model depicted in Fig. 6. Thus, we queried whether
the cluster mutations in the secondary site of vaccinia TopIB affect the formation of
plectonemic DNA braids, a key biochemical manifestation of TopIB-mediated synapsis
(Shuman et al., 1997). To perform this analysis, we modified the wild-type TopIB by changing
the Tyr274 nucleophile to phenylalanine, a maneuver that abolishes transesterification without
affecting noncovalent DNA binding or intramolecular synapsis (Shuman et al., 1997). As
shown in Fig. 5B, the incubation of relaxed plasmid DNA circles with stoichiometric amounts
of the Phe274 protein introduced torsional strain, which, after relaxation by catalytic amounts
of wild-type TopIB, resulted in the acquisition of up to 8 negative supercoils. As discussed
previously (Shuman et al., 1997), this reaction indicates that the TopIB-DNA synaptic complex
is a plectonemic supercoil in which the two duplexes encompassed by the protein filament are
interwound in a right-handed helix (Shuman et al., 1997). When the same F274 change was
introduced into the 5xAla, 3xAla and 8xAla cluster mutants, we found that they were uniformly
unable to promote such plectonemic braiding (Fig. 5B). We surmise that the poxvirus
equivalent of the secondary DNA site is essential for intramolecular synapsis and its topological
sequelae. Our results are consistent with model B in Fig. 1 as the basis for TopIB-mediated
synapsis.
In summary, we have demonstrated the existence, structure, and functional relevance of a
secondary DNA binding site on bacterial and viral TopIB. This work gives impetus and affords
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a structural guide to analogous studies of DNA crossover binding by eukaryal cellular TopIB
enzymes.
Materials and Methods
DraTopIB was produced and purified as described previously (Patel et al., 2006).
Complementary 12-mer DNA oligonucleotides (5′-GAATAAGGGCGC-3′ and 3′-
CTTATTCCCGCG-5′) were purified by reverse phase HPLC (Aggarwal, 1990) and then
annealed. Initial crystallization trials with mixtures of DraTopIB and 12-bp duplex DNA were
performed by the sitting drop vapor diffusion method in 96 well plates set up with a Hydra-II
crystallization robot. Small crystals grew in polyethylene glycol at 10°C with a 1:1 molar ratio
of protein and DNA. Refined conditions using the hanging drop vapor diffusion method yielded
larger crystals in 30% PEG 400 (w/v), 0.1 M sodium acetate (pH 4.5), 0.2 M calcium chloride.
Prior to data collection, the crystals were cryoprotected with 25% glycerol by stepwise transfer
through cooled (4° C) solutions of the well buffer containing increasing concentrations of
glycerol (in 5% glycerol increments per step and soaking for 2 min/step). Crystals were
harvested with a rayon crystal-mounting loop, and flash cooled in liquid nitrogen.
A complete diffraction data set was collected initially using a laboratory x-ray source. Later,
a higher resolution data set (to 1.65 Å) was collected using synchrotron radiation at DND CAT
at the Advanced Photon Source (Argonne National Laboratory). All data were processed using
XDS (Kabsch, 1993) and scaled using SCALA (Collaborative Computational Project 4,
1994). The crystals belong to space group C2 with unit cell dimensions a=119.7 Å, b=53.4 Å,
and c=77.4 Å, β = 96.3° and had one DraTopIB protomer in the asymmetric unit. Data
collection statistics are listed in Table I.
The structure was solved by molecular replacement with the program PHASER (McCoy et al.,
2007) using the C-terminal domain of the DraTopIB apoenzyme (PDB 2F4Q) as a search
model. Initial electron density maps clearly showed Fo−Fc difference density for DNA, but
only weak density for the N-terminal domain. After refinement, the entire DNA and most of
the C-terminal domain were built, but some regions of the N-terminal domain were disordered.
Refinement was performed with REFMAC5 (Murshudov et al., 1997) and model rebuilding
was executed in COOT (Emsley and Cowtan, 2004). The structure has a final Rwork and
Rfree of 19.3% and 22.2% respectively with 99.7% of the residues in the favored regions of the
Ramachandran plot and no outliers, and good rotamer distributions (Davis et al., 2004).
Refinement statistics for the complex are compiled in Table I. The coordinates of the final
model and the structure factors have been deposited in the PDB with accession code 3M4A.
Figures were made with PyMOL (DeLano 2002).
Wild-type vaccinia virus TopIB and three mutants with clustered alanine substitutions at the
secondary DNA binding site (5xAla, 3xAla, and 8xAla) were produced in E. coli with C-
terminal His6 tags and purified from soluble bacterial lysates by Ni-agarose and
phosphocellulose chromatography (Yakovleva et al., 2008). Topoisomerase reaction mixtures
containing (per 20 μl) 50 mM Tris-HCl (pH 7.5), 100 mM NaCl, 2.5 mM EDTA, 250 ng
supercoiled pUC19 plasmid DNA, and 2.5 ng of TopIB were incubated at 37° C. Aliquots (20
μl) were withdrawn at 5, 10, 20, 30, 60, 120 and 240 s and then quenched immediately with
SDS. “Time 0” samples were taken prior to adding TopIB. The mixtures were analyzed by
electrophoresis through a 1% horizontal agarose gel in TBE buffer (90 mM Tris-borate, 2.5
mM EDTA). DNA was visualized by staining with ethidium bromide and UV
transillumination.
Biochemical analysis of the topological changes in relaxed circular DNA accompanying
intramolecular synapsis was performed as described previously (Shuman et al., 1997). To
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prepare the relaxed plasmid DNA, a reaction mixture (100 μl) containing 50 mM Tris-HCl (pH
7.5), 100 mM NaCl, 5 mM MgCl2, 25 μg supercoiled pUC19 DNA, and 0.5 μg wild-type
vaccinia TopIB was incubated for 30 min at 37° C. The reaction was quenched by adding SDS
to 0.2% and EDTA to 20 mM. The mixture was digested with 1 μg proteinase K for 2 h at 37°
C, then extracted twice with phenol: chloroform and once with chloroform. The relaxed DNA
was recovered by ethanol precipitation and resuspended in 10 mM Tris-HCl (pH 7.5), 1 mM
EDTA. Vaccinia TopIB-F274 and F274 variants of the 5xAla, 3xAla, and 8xAla cluster
mutants were produced in E. coli with C-terminal His6 tags and purified by Ni-agarose and
phosphocellulose chromatography. To assay synaptic plectoneme formation, reaction mixtures
(20 μl) containing 50 mM Tris-HCl (pH 7.5), 100 mM NaCl, 2.5 mM EDTA, 250 ng relaxed
pUC19 DNA, and 100 or 200 ng of vaccinia TopIB-F274 or its Ala-cluster variants were
incubated for 15 min at 37° C. The mixtures were then supplemented where with 5 ng wild-
type vaccinia TopIB where specified and incubated for another 15 min at 37° C. The reactions
were halted by adding SDS to 0.5%. The mixtures were digested with 10 μg proteinase K for
1 h at 37° C and then analyzed by electrophoresis through a 1% horizontal agarose gel in TBE
buffer, in parallel with samples of supercoiled and relaxed pUC19. The DNA was visualized
by staining with ethidium bromide.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
Support from the R.H. Lurie Comprehensive Cancer Center of Northwestern University to the Structural Biology
Facility is acknowledged. This research was supported by NIH grants GM51350 (to A.M.) and GM46330 (to S.S.).
S.S. is an American Cancer Society Research Professor. Portions of this work were performed at the DuPont-
Northwestern-Dow Collaborative Access Team (DND-CAT) Synchrotron Research Center at the Advanced Photon
Source (APS). DND-CAT is supported by Dupont, DOW and the NSF. Use of the APS is supported by the Department
of Energy (DOE). We thank Greg Van Duyne for providing the coordinates of the poxvirus TopIB–DNA vanadate
transition state mimetic in advance of publication.
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Figure 1. Models for TopIB-mediated DNA synapsis
Single molecule imaging by EM and AFM has shown that poxvirus TopIB binds cooperatively
to linear plasmid DNA and forms protein-DNA filaments in which distant segments in the
same DNA molecule are synapsed. The TopIB protomer consists of a small N-terminal domain
(depicted as a cyan sphere) and a larger C-terminal domain (magenta sphere) that contains the
active site. The primary DNA binding site resides within a protein clamp formed by the N and
C domains. Two possible mechanisms to account for synapsis are depicted as models A and
B. In model A, TopIB-TopIB interactions promote synapsis of two TopIB-DNA filaments. In
model B, the synapsed DNA duplex is captured at a distinct secondary DNA binding site on
the TopIB protomer.
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Figure 2. A secondary DNA binding site on DraTopIB
The C-terminal catalytic domains of poxvirus TopIB (from the structure of the covalent TopIB-
DNA intermediate; PDB 2H7F) and DraTopIB (from the present DraTopIB-DNA cocrystal
structure) were superimposed and offset horizontally. The tertiary structures, depicted as beige
ribbons traces, are homologous throughout. The C-domains consist of two globular lobes (1
and 2, denoted for poxvirus TopIB in panel A). The active sites are located on the superior
surface of the C-domain (denoted by the arrow for DraTopIB in panel B); the catalytic amino
acid side chains are shown as stick models. The specificity helix is shown in magenta. The
duplex DNA segments of the respective DNA ligands are shown as gray spacing-filling models.
The DNA duplex is covalently linked to poxvirus TopIB is the primary (catalytic) DNA binding
site. By contrast, DraTopIB binds its DNA ligand on the opposite face of the C-domain ~30
Å away from the primary DNA site. See also Figure S1 for a comparison of the DNA contacts
of the specificity helix in poxvirus TopIB versus DraTopIB.
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Figure 3. Close-up view of the interactions between DNA and DraTopIB
(A) A stereo image of the secondary DNA binding site is shown. Side chains and a main chain
amide that contact the DNA ligand are depicted as sticks. Hydrogen bonds are denoted by
dashed lines. (B) The 12mer duplex DNA ligand is depicted as a two-dimensional base-paired
ladder. Atomic contacts between the indicated amino acid side chains (or the Gly117 amide)
and the DNA phosphates or nucleobases are indicated by arrows.
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Figure 4. Conservation of the secondary DNA binding site
(A) A stereo image of the secondary DNA binding site in DraTopIB in shown, viewed from
the opposite side of the C-domain as the image in Fig. 3. The DNA is depicted as a cartoon
with the phosphate backbone chain shown in yellow and the nucleobases as blue sticks. (B)
The corresponding region of the C-domain of poxvirus TopIB is shown in the same orientation
as in panel A. The stereo image highlights the basic and polar side chains that are candidates
to comprise the secondary DNA site in poxvirus TopIB. (C) This panel shows the aligned
amino acids sequences of the two protein segments comprising the actual or imputed secondary
DNA binding sites in exemplary bacterial, eukaryal cellular, and poxvirus TopIB enzymes –
D. radiodurans (Dra) TopIB, Homo sapiens (Hsa) TopIB, and vaccinia (vac) TopIB,
respectively – for which crystal structures are available. The aligment is based on superposition
of the tertiary structures. The nine DraTopIB amino acids that contact the DNA in the secondary
site are denoted by ●. The amino acids clusters in vaccinia TopIB that, when simultaneously
mutated to alanine abolished synaptic plectoneme formation, are denoted by |. The four lysines
in HsaTopIB implicated in crossover recognition on the basis of the effects of clustered lysine-
to-glutamate changes (Yang et al., 2009) are underlined. Positions of side chain identity/
similarity at the actual or imputed DNA binding residues are highlighted in yellow shading.
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Figure 5. Mutations at the secondary binding site of poxvirus TopIB abolish formation of
plectonemic synaptic complexes but have little or no effect on supercoil relaxation
(A) Kinetics of relaxation of 250 ng pUC19 DNA by 2.5 ng of vaccinia TopIB, either wild-
type (WT) or the indicated mutants with clustered alanine substitutions at the putative
secondary DNA binding site. The DNA products were resolved by native agarose gel
electrophoresesis. The supercoiled (S) and relaxed (R) circular DNAs are indicated on the
left. (B) Plectonemic supercoiling. Relaxed circular pUC19 DNA was incubated with 100 or
200 ng of vaccinia TopIB-Phe274 or the indicated Ala-cluster mutants thereof, then treated
where indicated with 5 ng of wild-type TopIB to relax any supercoils introduced by binding
of the F274 protein(s). The products were resolved by native agarose gel electrophoresis.
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Figure 6. Model of TopIB with primary and secondary DNA sites occupied
The diagram shows a model of poxvirus TopIB with both DNA binding sites occupied. The
model was built by aligning the DraTopIB-DNA complex to the crystal structure of poxvirus
TopIB bound covalently to its DNA target site as a vanadate transition state mimetic (Perry et
al., 2010; PDB 3IGC) and then deleting the DraTopIB protein to leave just the 12-mer DNA
in the secondary site. Three different views of the model are shown in A, B and C. The primary
DNA ligand is enveloped within a circumferential protein clamp formed by the N- and C-
domains. The secondary DNA ligand is docked at the base of the C-domain with one strand of
the duplex fitting into the secondary binding site (B).
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Table I
Diffraction data and refinement statistics
Data collection
Space group
C2
Cell
119.7 Å, 53.4 Å, 77.4 Å, β=96.33
Detector type/source
MAR-CCD/APS
Wavelength (Å)
1.00
Resolution (Å)
1.65
Detector type/source
MAR345/home
Wavelength (Å)
1.5418
Resolution (Å)
1.92
Merged data sets
Measured reflections
265,853
Unique reflections
56,385
Completeness (%)a
96.2 (89.5)
Rsym (%)a,b
4.8 (27.6)
Rmeas (%)a,c
5.3 (32.5)
Redundancy
4.7 (3.4)
Mean(I/σ(I))
24.1 (4.1)
Refinement
Resolution (Å)
76 – 1.65 (1.65 – 1.693)
Number of reflections: working set/test set
53,565/2,820 (3,633/209)
Rworkf
19.3 (22.0)
Rfreed
22.2 (23.2)
Protein atoms
2,495
DNA atoms
487
Water molecules
278
Other
12
r.m.s.d. from target values
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Bond lengths (Å)
0.008
Bond angles (°)
1.29
Average B-factor (Å2):
Main chain
16.0
Side chain
17.7
DNA
15.2
Solvent
24.6
Ramachandran plot
Favored (%)f
99.7
Outliers (%)f
0.0
Rotamer outliers (%)f
0.4
aNumbers in parenthesis represent values in the highest resolution shell.
bRsym=Σ|I−<I>|/ΣI, where I=observed intensity, and <I>=average intensity obtained from multiple measurements.
cRmeas as defined by (Diederichs and Karplus, 1997).
dR-factor=Σ||Fo| − |Fc||/Σ|Fo|, where |Fo|=observed structure factor amplitude and |Fc|=calculated structure factor amplitude.
eRfree: R-factor based on 5% of the data excluded from refinement.
fCalculated with MolProbity (Davis et al., 2004).
Structure. Author manuscript; available in PMC 2011 June 9.
|
3M4D
|
Crystal structure of the M113N mutant of alpha-hemolysin
|
Molecular bases of cyclodextrin adapter interactions
with engineered protein nanopores
Arijit Banerjeea, Ellina Mikhailovaa, Stephen Cheleya, Li-Qun Gub,2, Michelle Montoyac,3, Yasuo Nagaokaa,4,
Eric Gouauxd, and Hagan Bayleya,1
aDepartment of Chemistry, University of Oxford, Oxford, OX1 3TA, United Kingdom; bDepartment of Medical Biochemistry & Genetics, Texas A&M
University System Health Science Center, College Station, TX 77843-1114; cDepartment of Biochemistry and Molecular Biophysics and Howard Hughes
Medical Institute, Columbia University, New York, NY 10032; and dVollum Institute and Howard Hughes Medical Institute, Oregon Health and Science
University, Portland, OR 97239
Edited by Gregory A. Petsko, Brandeis University, Waltham, MA, and approved March 3, 2010 (received for review December 15, 2009)
Engineered protein pores have several potential applications in
biotechnology: as sensor elements in stochastic detection and
ultrarapid DNA sequencing, as nanoreactors to observe single-
molecule chemistry, and in the construction of nano- and micro-
devices. One important class of pores contains molecular adapters,
which provide internal binding sites for small molecules. Mutants
of the α-hemolysin (αHL) pore that bind the adapter β-cyclodextrin
(βCD) ∼104 times more tightly than the wild type have been ob-
tained. We now use single-channel electrical recording, protein en-
gineering including unnatural amino acid mutagenesis, and high-
resolution x-ray crystallography to provide definitive structural in-
formation on these engineered protein nanopores in unparalleled
detail.
alpha-hemolysin ∣single molecule ∣stochastic sensing ∣structure ∣
unnatural amino acid
M
any research groups have used protein engineering to
obtain enzymes and antibodies with new properties suited
for specific tasks (1–6). Fewer groups have taken on the difficult
problem of engineering membrane proteins (7). We have engi-
neered the α-hemolysin protein pore, mindful of several potential
applications in biotechnology, including its ability to act as a de-
tector in stochastic sensing (8) and ultrarapid DNA sequencing
(9), to serve as a nanoreactor for the observation of single-
molecule chemistry (10) and to act as a component for the con-
struction of nano- and microdevices (11).
An important breakthrough in this area, which enabled the sto-
chastic sensing of organic molecules including the detection of
DNA bases in the form of nucleoside monophosphates (12, 13),
was the discovery of internal molecular adapters, a form of non-
covalent protein modification (14). Most useful have been cyclo-
dextrin (CD) adapters, which have until now been used in the
absence of detailed structural information about how they work.
The present paper is a definitive investigation, which provides
such information through the application of a wide variety of
technical approaches: single-channel electrical recording, protein
engineering including unnatural amino acid mutagenesis, and
x-ray crystallography. The studies employing mutagenesis show
that the striking interactions seen in the crystal structures also
occur in individual pores in lipid bilayers.
We reveal that the tight-binding αHL mutants (15) M113N7
and M113F7 bind βCD in different orientations within the hep-
tameric pore. In the case of M113N7, the top (primary hydroxyls)
of the CD ring faces the trans entrance of the pore. In the case of
M113F7, the bottom (secondary hydroxyls) of the CD ring faces
the trans entrance, while the top of the ring is bonded to the pore
through remarkable CH-π interactions. Another tight-binding
mutant, M113V7, can bind the CD in both orientations. These
results illustrate the exquisite level of engineering that can be
achieved with protein nanopores, which is, for example, far be-
yond what is possible with solid-state pores. The work also pro-
vides information valuable for the design of new binding sites
within the lumen of the αHL pore or within other β-barrel pro-
teins. Our results will be of interest to others exploring the inter-
actions of CDs with the αHL pore (16, 17), including groups
involved in computational studies (18, 19). In addition CDs bind
to a variety of other pores, including porins (20, 21) and connex-
ins (22), and are being tested in vivo as blockers of the anthrax
protective antigen pore (23, 24). The CD adapter concept has
also been incorporated into other formats, e.g., with glass nano-
pores (25), and artificial pores based on CDs have been made by
several groups (26–28). Our work is pertinent to these studies.
Results
Kinetics and Thermodynamics of the Interactions of βCD with αHL
Pores Containing Met, Phe and Asn at Position 113. We showed earlier
that position 113 in the αHL pore (Fig. 1A) is critical for the bind-
ing of βCD (14). Subsequently, residue 113, which is Met in the
WT protein, was changed to each of the remaining 19 naturally
occurring amino acids by site-directed mutagenesis (15). We
found that 11 of these mutants, expressed as homoheptamers,
bound βCD with a similar affinity and with similar kinetics to
the WT homoheptamer. Two mutants (P, W) bound βCD about
10 times more strongly than the WT homoheptamer, while six of
them (V, H, Y, D, N, F) bound with high affinity, i.e., with a Kd
value 103 to 104 times lower than the WT.
Remarkably, the side chains of the latter six amino acids bear
little resemblance to one another, and this issue is addressed in the
present paper. We first examined the two amino acids with the
most disparate side chains (Fand N) by making heteromeric pores
containing WT (Met-113), M113F, and M113N subunits. Three
series of heteroheptamers were produced: WT7−nM113Nn,
WT7−nM113Fn, and M113F7−nM113Nn. The heteroheptamers
were separated by SDS-polyacrylamide gel electrophoresis aided
by an oligoaspartate (D8) tail on the first of the two types of sub-
unit (Fig. 1B) (29). All 21 combinations of WT, M113F, and
M113N subunits formed αHL pores that interacted with βCD as
shown by single-channel current recordings, which revealed the
extent of block by βCD (Fig. S1), the association and dissociation
Author contributions: A.B., S.C., E.G., and H.B. designed research; A.B., E.M., S.C., L.-Q.G.,
M.M., and Y.N. performed research; A.B., E.G., and H.B. analyzed data; and A.B. and
H.B. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
1To whom correspondence should be addressed. E-mail: hagan.bayley@chem.ox.ac.uk.
2Present address: Department of Biological Engineering and Dalton Cardiovascular
Research Center, University of Missouri, Columbia, MO 65211.
3Present address: Nature Structural & Molecular Biology, 75 Varick Street, 9th Floor, New
York NY 10013-1917.
4Present address: Department of Biotechnology, Faculty of Engineering, Kansai University,
3-3-35 Yamate-cho, Suita, Osaka 564-8680, Japan.
This article contains supporting information online at www.pnas.org/cgi/content/full/
0914229107/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.0914229107
PNAS ∣May 4, 2010 ∣vol. 107 ∣no. 18 ∣8165–8170
BIOCHEMISTRY
rate constants for βCD (kon and koff), and (from the latter) the
equilibrium dissociation constant for βCD (Kd ¼ koff∕kon) (15).
The kon values for βCD for the 21 combinations of subunits
were all similar at ∼5 × 105 M−1 s−1 (Fig. 1C, Upper). By contrast,
the koff values differed widely, ranging from ∼5 × 10−2 s−1 to
∼103 s−1. For WT7−nM113Nn and WT7−nM113Fn, the koff values
decreased as M113N or M113F subunits were added. In the case
of M113N, there was a steep drop in the value of koff after the
fifth subunit had been incorporated. In the case of M113F,
the decrease in the value of koff occurred less precipitously as the
M113F subunits were added (Fig. 1C, Lower). Intriguingly, with
M113F7−nM113Nn, koff first increased as M113N subunits were
added to M113F7 until n ¼ 4 (M113F3M113N4) and then de-
creased for larger values of n (Fig. 1C, Lower). We recognize that
there is more than one permutation of heteromers containing two
to five mutant subunits (Fig. 1B), but we have ignored this fact
here because no significant differences in the properties of indi-
vidual heteromers were observed. For example, 42 recordings
were made of WT5M113N2, which has three permutations.
Because, kon showed little variation with subunit composition,
the variation in Kd was similar to the variation in koff (Fig. 1C).
While these studies were in progress, the crystal structures of
βCD complexed to M113N7 (Fig. 2B) and M113F7 (Fig. 2C) were
solved (Table S1) (30). High-resolution structures could be
obtained because the CD and the αHL pore have the same C7
symmetry. In the case of M113N7, βCD is bound with the second-
ary hydroxyl face “upward” (Fig. 2B). In each glucose unit of the
βCD, the 2-hydroxyl is hydrogen bonded to the side-chain amide
of an Asn-113 (the residue introduced by mutagenesis) and the
3-hydroxyl is hydrogen bonded to the ϵ-amino group of Lys-147.
In the case of M113F7, two βCDs are bound to the αHL pore
(Fig. 2C). It is the top βCD in the structure that concerns us, be-
cause it is in contact with the Phe-113 residues introduced by mu-
tagenesis. It is immediately apparent that the top βCD in M113F7
is in the opposite orientation to the βCD in M113N7 with each
6-hydroxyl group in a CH-π bonding interaction (31–35) with a
Phe-113 side chain. The opposite orientations of the βCDs in
M113N7 and M113F7 immediately explain why heteromers
formed from similar numbers of M113N and M113F subunits
(e.g., M113N4M113F3) bind βCD weakly (see also Discussion).
Unnatural Amino Acid Mutagenesis. To further explore the range of
noncovalent interactions that are available when βCD binds to
the αHL pore, five unnatural amino acids (Fig. 3A and Fig. S2)
were incorporated at position 113, by using the in vitro nonsense
codon suppression method (36). In particular, we had noted that
M113V7 containing the β-branched Val binds βCD tightly (15),
and therefore we compared cyclopropylglycine (Cpg) and cyclo-
propylalanine (Cpa). We also further examined the means by
which M113F7 binds βCD tightly, by comparing the properties of
4-fluorophenylalanine (f1F), pentafluorophenylalanine (f5F),
and cyclohexylalanine (Cha) at position 113.
The five homomeric pores all produced single-channel cur-
rents with unitary conductance values in the range expected
for properly assembled heptamers (Fig. S3). All five bound βCD
(Fig. 3B, Level 2), either tightly (f1F, Cpg) or weakly (f5F, Cha,
Cpa) as described in detail below. During the long βCD binding
events, additional current spikes were seen (Fig. 3B). Similar
Fig. 1.
Binding of βCD by heteromeric αHL pores formed by WT, M113F and M113N subunits. (A) Crystal structure of WT-αHL (61) showing residue 113 (Met,
yellow). Left panel, side view and right panel, top view. (B) Separation of 35S-labeled αHL heteroheptamers by SDS-polyacrylamide electrophoresis. The
separation of the M113F7-nM113Nn heteromers is shown as detected by autoradiography of a dried gel. The M113F subunits carried a D8 tail. Lane 1,
M113N7; lane 2, M113F7−nM113Nn (the heteromers formed from several preparations made with differing ratios of M113F and M113N subunits were mixed
to give roughly equal amounts of each subunit combination); lane 3, M113F7. A diagram of the eight different combinations of subunits and their permuta-
tions is shown to the right of the autoradiogram. The various permutations are not separated by electrophoresis. (C) Kinetics of the interaction of βCD with
single heteromeric αHL pores as determined by bilayer recording. Values of kon were calculated by using kon ¼ 1∕ðτon½βCDÞ, where τon is the mean interevent
interval. Values of koff were determined by using koff ¼ 1∕τoff, where τoff is the mean dwell time of βCD in the pore. Values of Kd were calculated by using
Kd ¼ koff∕kon. Each point represents the mean s:d: for three or more determinations. Where they cannot be seen, the s.d. values lie within the symbol. Black
squares, WT7−nM113Nn; gray squares, M113F7−nM113Nn; empty squares, M113F7−nWTn.
8166
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Banerjee et al.
events had been observed previously with certain Met-113 repla-
cement mutants and may represent movement of the βCD at its
binding site (e.g., rotation about axes perpendicular to the C7
axis) (15). The additional current spikes were more prevalent
for M113V7 and M113Cpg7, which may take part in more con-
formationally labile interactions with βCD, compared with say
M113F7 (Fig. S4).
Interactions of βCD with Homoheptamers Bearing Aromatic Residues
at Position 113. To further understand the nature of the binding of
βCD to aromatic side chains, we examined the kinetics of βCD
binding to the homoheptamers containing f1F or f5F at position
113, M113f1F7 and M113f5F7 (Fig. 3C). For both mutants, the
value of kon was very similar to that of WT7, but the values of koff
and therefore Kd for M113f1F7 differed dramatically from WT7
and were close to the values for the tight-binding mutant M113F7
(Table S2A). By contrast, koff and Kd for M113f5F7 were similar
to the values for WT7 (Table S2A).
To determine whether M113f1F7 binds βCD in the same orien-
tation as M113F7 (Fig. 2C), we made heteromers of the M113f1F
subunit with M113N or M113F and examined M113F4M113f1F3
and M113N4M113f1F3. M113F4M113f1F3 binds βCD as strongly
as either M113F7 or M113f1F7, but M113N4M113f1F3 binds
βCD weakly with a similar affinity to WT7 (Fig. 3D and
Table S3). Therefore, it is reasonable to infer that M113F7
and M113f1F7 bind βCD in the same orientation with the 6-
hydroxyl groups of the CD in proximity to the aromatic rings
on the protein.
Cyclohexylalanine (Cha) was used to replace the aromatic side
chains with a roughly isosteric hydrophobic group. Again the va-
lue of kon for βCD was little changed, but koff for M113Cha7 had
an intermediate value of 42 6 s−1. Therefore, M113Cha7 binds
βCD more weakly than M113F7 but distinctly more strongly than
the WT7 pore (Table S2A and Fig. 3C).
Interactions of βCD with Homoheptamers Bearing Hydrophobic Resi-
dues at Position 113. M113V7 binds βCD very strongly, and there-
fore we compared αHL pores with Cpg or Cpa at position 113.
Cpg is roughly isosteric with Val, and like Val has a β-branched
side chain. Gratifyingly, M113Cpg7 has a kon value similar to the
other αHL pores, and koff and Kd values close to those of
M113V7 (Table S2B and Fig. 3E). Cyclopropylalanine (Cpa), with
an additional methylene group compared to Cpg, is roughly
isosteric with Leu, a weak binder, and M113Cpa7 also binds
βCD weakly with kon, koff and Kd values similar to those of
WT7 (Table S2B and Fig. 3E). M113I7 and M113T7, which are
β-branched, are also weak binders, but Ile and Thr are less closely
related to Val than Cpg.
To determine whether M113V7 binds βCD in the same orien-
tation as M113F7 or M113N7 (Fig. 2), we made heteromers of
M113V and the M113N or M113F subunits. M113V3M113F4,
M113V4M113F3, M113V3M113N4, and M113V4M113N3 were
examined in detail. All four heteroheptamers bound βCD more
weakly than M113V7, M113F7 or M113N7 (Fig. 3F and Table S4),
suggesting that Val at position 113 interacts with βCD strongly but
in a different manner to either Phe or Asn. Each heteromer
exhibited a range of Kd values, perhaps reflecting the various pos-
sible permutations of the two different subunits around the cen-
tral axis of the heptamer, although this heterogeneity was not
seen for heteromers made from WT, M113F and M113N (Fig. 1).
Discussion
Soon after we discovered that βCD binds to the WT-αHL pore for
around a millisecond, we found a mutant pore, M113N7, that re-
leases βCD ∼104 times more slowly (14). This prompted us to
examine all 19 mutants in which residue 113 is replaced by a nat-
ural amino acid, with the surprising result that a collection of ami-
no acids with structurally unrelated side chains (V, H, Y, D, N, F)
are tight binders (15). Here, we have examined the nature of the
binding interactions more closely by single-channel electrical re-
cording, protein engineering including unnatural amino acid mu-
tagenesis, and high-resolution x-ray crystallography, and we
provide the first definitive structural information on an engi-
neered protein nanopore.
We find that βCD can bind tightly to the αHL pore in three
different ways depending on the residue at 113, as exemplified
by Asn, Phe, and Val. Because Asn and Phe have quite different
side chains, we first compared the ability of M113N and M113F
subunits to take part in binding the CD. The examination of het-
eromeric proteins containing WT (Met-113), M113N and M113F
subunits showed that the replacement of WT subunits in WT7
with M113N or M113F subunits led to increased affinity for
βCD. The more M113N or M113F subunits that were added, the
tighter binding became. By contrast, when subunits in M113N7
were replaced with M113F subunits, binding became weaker,
reaching a minimum at three to four M113F subunits, and then
increasing in strength with five M113F subunits or more (Fig. 1C).
Parallel structural studies (30) revealed the basis of the “oppos-
ing” effects of the M113N and M113F subunits. βCD binds to
M113N7 in the opposite orientation to that in which it binds
to M113F7. In M113N7, the secondary hydroxyls in the βCD ring
are hydrogen bonded to Lys-147 and Asn-113 (Fig. 2A). By con-
trast, βCD interacts with M113F7 through its primary hydroxyl
face (Fig. 2B).
It seemed likely that M113V7, bound βCD in yet another way,
and this was examined by forming heteromers between M113V
and M113N or M113F. The presence of three or four subunits
of either M113N or M113F greatly decreases the affinity of
the pore for βCD (Fig. 3F), with an average koff of 7.3 × 101 s−1,
indicating that a third binding mode is indeed operating
Fig. 2.
X-ray structures of M113N and
M113F homoheptamers with βCD bound.
(A) Side view of heptameric αHL. βCD binds
in the blue highlighted region. (B) βCD
bound to M113N7 (dotted lines indicate hy-
drogen bonding). The side chains of Lys-147
are in pale brown and the side chains of Asn-
113 in yellow. (C) βCD bound to M113F7
(dotted lines indicate CH-π bonding). The
side chains of Phe-113 are in yellow. The sec-
ond βCD in the M113F7 · ðβCDÞ2 structure is
hydrogen bonded to the top βCD in a head-
to-head arrangement and has no apparent
interactions with the protein. For both (B)
and (C), four β strands were omitted from
the barrel to give a better view.
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(Table S4). In summary, the three groups of tight-binding mutants
comprise αHL pores incorporating, at position 113: (i) the hydro-
gen-bonding amino acids N, D (the latter would have to be largely
in the protonated form), and possibly H; (ii) the aromatics F, Y,
f1F, and possibly H, and more weakly W; (iii) the β-branched ami-
no acids V, Cpg. There may be yet other means by which CDs can
bind to the αHL pore. For example, we earlier found that hepta-
6-sulfato-βCD can bind tightly to αHL pores containing the
N139Q mutation (37). Presumably, this CD is bound at a site low-
er down in the β barrel in a fashion that includes hydrogen bond-
ing to the Gln at position 139. While the various mutants
exhibited widely different koff values, the value of kon was almost
invariant and averaged ∼2.3 × 105 M−1 s−1 (Table S2) (15). Ap-
parently, transport to the binding site is rate limiting, through
a route unaffected by mutagenesis.
koff increased precipitously with the addition of WTsubunits to
M113N7 (Fig. 1C). Crystal structures of M113N7 show that resi-
dues
111,
113,
and
147
are
reorganized
by
compari-
son with WT7 and then undergo a more limited rearrangement
when βCD binds (Fig. S5). For example, the side chain of
Lys-147 shifts position to form a bifurcated hydrogen bond with
a 3-hydroxyl group of βCD and the side chain carbonyl of an Asn-
113 (Fig. S6). Therefore, the side chains of residues 111, 113, and
147 might be in a variety of conformations in WT7−nM113Nn het-
eromers and offer less well preorganized binding sites for βCD
than they do in M113N7. Further, the intramolecular hydrogen
bonds of the secondary hydroxyls in βCD (38) must be disrupted
upon binding as both hydroxyls on each glucose ring form hydro-
gen bonds to the mutant subunits (Fig. 2B). Because the hydrogen
bonds that are broken in βCD are arranged in a circle, the break-
age of bonds involving a single glucose (three bonds in all) will be
relatively more disruptive than those involving adjoining glucose
residues or the entire circle. The overall binding cooperativity in
M113N7 could be attributed to enthalpic cooperativity outweigh-
ing entropic penalties to binding (39). Positive cooperativity has
been observed previously in fairly rigid model systems (40).
By contrast with M113N7, there is little movement of side
chains in ðM113FÞ7 by comparison with WT7 and little move-
ment, including Phe-113, upon binding βCD (Fig. S7A). Further,
the crystal structure of M113F7 · βCD suggests that each Phe re-
sidue interacts independently with the βCD through what appear
to be CH-π interactions (Fig. S7B). These interactions are ex-
pected to be weak and not strongly directional and hence offer
less enthalpic cooperativity, as supported by the B-factors (crys-
tallographic temperatures factors) at the primary βCD binding
site, which are between ∼40 and 50. Positive cooperativity is ob-
served, but it is less pronounced than in the case of M113N7
(Table S5). In the case of M113N7, the B-factors of the residues
that bind βCD are in the 20s implying that the βCD is more rigidly
held than it is in M113F7.
The binding of sugars to aromatic residues in proteins can in-
clude CH-π bonding (41) or OH-π bonding or a finely balanced
complement of both (42, 43). However, we have dismissed the
possibility of an OH-π interaction between Phe-113 and the
6-hydroxyl groups of βCD as the distance between the center
of the phenyl rings to the nearest hydroxyl oxygen is higher
(5.2 0.65 Å, n ¼ 7) than that expected for a favorable OH-π
interaction (33). While we propose that βCD binds to Phe-113
through a C-6 CH-π interaction (Fig. S7B), the distances between
the center of the Phe-113 ring and the nearest C-6 of βCD ob-
served in the M113F7 · βCD structure (4.66 0.24 Å, n ¼ 7)
are in the upper range of the expected distance for a strong inter-
action, which is ∼4.5 Å (33). The angle between the normal to the
aromatic rings and the line connecting the C-6 atoms to the aro-
matic midpoint is 8.0 5.6°, which is well within the expected
range (44). The measurements with M113f5F7 argue against a
hydrophobic interaction between Phe residues at position 113
and the βCD ring. In f5F, the hydrophobicity of the phenyl ring
is significantly increased (45) yet M113f5F7 binds βCD weakly,
like WT7 (Fig. 3C and Table S2A).
By contrast with F, f1F, Y and N, homomeric αHL pores with
f5F and W at position 113 bound βCD relatively weakly (Fig. 3C
and Table S2A). In the case of f5F, the powerful electron with-
drawing action of the five fluorine atoms leaves a highly increased
positive charge at the center of the ring (46, 47), mitigating
against a hydrogen-bonding interaction. The electron-rich Trp
Fig. 3.
Properties of pores containing natural and unnatural amino acid sub-
stitutions at position 113. The data were recorded at þ40 mV in 1.0 M NaCl,
10 mM sodium phosphate, pH 7.5. (A) Unnatural amino acids used in this
study: 4-fluorophenylalanine, f1F; pentafluorophenylalanine, f5F; cyclohex-
ylalanine, Cha; cyclopropylglycine, Cpg; cyclopropylalanine, Cpa. (B) Repre-
sentative
current
traces
from
single
homoheptameric
αHL
pores,
containing unnatural amino acids at position 113, in the presence of βCD.
βCD (40 μM final) was added to the trans chamber. Level 1, open pore current;
level 2, pore occupied by βCD. The broken line indicates zero current. (C) In-
teraction of βCD with homomeric αHL pores containing aromatic amino acids
at position 113. Kd values for the interaction between βCD and the αHL pore
were calculated by using Kd ¼ koff∕kon. Each column represents the mean
s:d: for 10 or more determinations: dark gray, natural amino acids; light gray,
unnatural amino acids. Data adapted from Gu and colleagues (15) are
marked (*). (D) Representative current traces from single-channel recordings
of βCD binding to M113F4M113f1F3 and M113N4M113f1F3. βCD (40 μM final)
was added to the trans chamber. The broken line indicates zero current. (E)
Interaction of βCD with homomeric αHL pores containing hydrophobic amino
acids at position 113. Kd values for the interaction between βCD and the αHL
pore were calculated by using Kd ¼ koff∕kon. Each column represents the
mean s:d: for ten or more determinations: dark gray, natural amino acids;
light gray, unnatural amino acids. Data adapted from Gu and colleagues (15)
are marked (*). (F) koff values for βCD from heteroheptamers formed with
M113F and M113V subunits and with M113N and M113V subunits. βCD
(40 μM final) was added to the trans chamber. The kon values for βCD for
all these mutants are similar, at ∼3 × 105 M−1 s−1. Empty square: average
koff values for the mutant (bar is s:d). Filled square: M113V3M113F4; filled
circle: M113V4M113F3; filled upright triangle: M113V3M113N4; filled in-
verted triangle: M113V4M113N3.
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Banerjee et al.
ring (44, 46, 47) should favor hydrogen bonding, but here we can-
not make a direct comparison with the crystal structure of
M113F7 as the indole ring is far larger than benzene. It is possible
that it cannot become oriented in the same manner and that it is
misaligned for hydrogen bonding.
Our experiments suggest that M113V7 and M113Cpg7 bind
βCD in a third way. In heteromers with M113V, both M113F
and M113N reduce the affinity of the pore for βCD suggesting that
neither the CH-π interaction with Phe-113 nor the hydrogen-
bonding interactions with Asn-113 and Lys-147 are compatible
with binding to Val. Close interactions of Val with glucose rings
have been noted previously (48). Therefore, we propose that the
Val side-chain interacts with the side of the glucose ring. This in-
teraction might occur in one or both orientations of the CD
ring (Fig. 4).
Conclusion
We provide structural information on engineered protein nano-
pores and describe three distinct ways in which βCD can bind
within the lumen of mutant αHL pores in atomic detail. Our re-
sults will be useful in several areas of basic science and biotech-
nology. By using host molecules lodged within the αHL pore,
host-guest interactions can be investigated in fine detail at the
single-molecule level (17, 49). The present work will now permit
us to examine binding events at a known face of a CD. The work
also provides information for designing new binding sites within
the lumen of the αHL pore (37) or within other β barrel proteins
(21, 50) and for using molecular design to devise ways in which to
covalently attach CDs within pores (13, 51). These areas impact
practical applications of nanopore technology including stochas-
tic sensing (8), single-molecule DNA sequencing (9, 12, 13, 52),
the use of nanoreactors for the observation of single-molecule
chemistry (10), and the construction of nano- and microdevices
(11, 53), as well as the design of CDs as therapeutic agents
(23, 24).
Methods
Full details of the experimental procedures can be found in SI Appendix.
Materials
L-Amino acids were obtained as follows: 4-fluorophenylalanine (f1F) (Fluka);
pentafluorophenylalanine (f5F) (PepTech Corp.); cyclopropylglycine (Cpg) (Ty-
ger); cyclopropylalanine (Cpa) (Tyger). 4-N-benzoyl-5′-O-(4,4′-dimethoxytri-
tyl)-2′-deoxycytidine-3′-[(2-cyanoethyl)-(N,N-diisopropyl)]-phosphoramidite
and bis(2-cyanoethyl)-N,N-diisopropylphosphoramidite for the synthesis of
pdCpA were purchased from Glen Research and Toronto Research Chemicals,
respectively.
Preparation
of
NVOC-Protected
Aminoacyl-pdCpA.
NVOC-protected
aminoacyl-pdCpAs were prepared as reported previously by reacting the
dinucleotide pdCpA with N-protected, carboxylic acid-activated, amino
acids (54–56).
Preparation of NVOC-Protected Aminoacyl-tRNA. NVOC-protected aminoacyl-
pdCpAs were ligated enzymatically with a truncated tRNA, prepared by using
methods described elsewhere (57, 58).
Genetic Constructs and Mutagenesis. All new αHL constructs were verified by
DNA sequencing. Details of each construct can be found in SI Appendix.
Synthesis, Assembly, and Purification of Mutant αHL pores. αHL monomers (WT
and mutants) were prepared in vitro by coupled transcription and translation
(IVTT) and assembled into homoheptamers on rabbit red blood cell
membranes followed by purification by SDS–PAGE as described earlier
(59). Heteroheptamers were prepared by mixing the two required DNAs
(one encoding an αHL with a D8 tail) before IVTT and then oligomerizing
the mixed translation products on rabbit red blood cell membranes. Pores
with the desired combinations of subunits were purified by SDS–PAGE (59).
Synthesis, Assembly, and Purification of αHL Mutants Containing Unnatural Ami-
no Acids. αHL polypeptides containing unnatural amino acids were synthe-
sized by IVTT in the presence of rabbit red blood cell membranes. The
plasmid with a stop codon (TAG) at position 113 was used. Deprotected ami-
noacyl-tRNAs (SI Appendix) were added to the IVTT mixtures. For heterohep-
tamers with subunits containing unnatural amino acids in combination with
M113N or M113F, monomers were first made, which were then coassembled
on rabbit red blood cell membranes and subsequently purified by SDS–PAGE.
Single-Channel Current Recordings in Planar Lipid Bilayers. (15, 60) Recordings
were made with 1.0 M NaCl, 10 mM sodium phosphate, pH 7.5, in both cham-
bers, at an applied potential of þ40 mV. Data were recorded at 22 2°C. The
bilayer was formed from 1,2-diphytanoyl-sn-glycero-phosphocholine (Avanti
Polar Lipids). Proteins were added to the cis chamber, and βCD to the trans
chamber. Single-channel currents were recorded with an Axopatch 200B
patch-clamp amplifier (Axon Instruments) and filtered at 2 kHz with a
built-in 4-pole low-pass Bessel Filter. The data were acquired at a sampling
rate of 10 kHz. For mutants that bind βCD strongly, the data were acquired
for at least 30 min and for weak-binding mutants for at least 10 min.
Kinetic Data Analysis. Current amplitude and dwell-time histograms were
made by using ClampFit 9.0. The mean dwell times, τoff, were determined
by fitting the dwell-time histograms to single exponentials. Values of kon
and koff were obtained by using the mean dwell times and mean interevent
intervals, as described previously (15, 60). This analysis assumes a binary in-
teraction, which was supported in all cases examined by the finding of only
one major blockade level and a single exponential distribution of dwell
times (τoff).
Fig. 4.
Molecular model showing the three classes of interaction between
the αHL pore and βCD identified in this work. The model identifies the region
of βCD responsible for each interaction (H atoms interacting with Phe-113 or
Asn-113 and Lys-147: gray). The first class of interaction is with aromatic
residues and involves the seven -CH2OH groups of the βCD. The second class
is typified by the interactions with Asn at position 113, which involve hydro-
gen-bonds to the secondary 2-hydroxyls of the βCD. Structural studies show
that this interaction is supported by hydrogen bonding between Lys-147 and
the secondary 3-hydroxyls of the βCD. Structural studies and experiments
with heteromers suggest that the βCD in M113F7 is in the opposite orienta-
tion to the βCD in M113N7, in support of the model shown here. As the inter-
action with Val is hydrophobic, it is not directional and βCD may not bind at
the same position inside the β barrel as it does in M113F7 or M113N7.
Banerjee et al.
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Protein Crystallography. Details can be found in SI Appendix. Protein Data
Bank: The coordinates and structure factors of the described structures have
been deposited with accession codes 3M2L ðM113F7Þ, 3M3R ðM113F7 · βCDÞ,
3M4D ðM113N7Þ, 3M4E ðM113N7 · βCDÞ.
ACKNOWLEDGMENTS. We thank Dennis Dougherty for the plasmid pTHG73.
This work was funded by a Royal Society Wolfson Research Merit Award
(to H.B.), the Medical Research Council (H.B.), the National Institutes of
Health (H.B.), and the Howard Hughes Medical Institute (E.G.).
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|
3M4E
|
Crystal structure of the M113N mutant of alpha-hemolysin bound to beta-cyclodextrin
|
Molecular bases of cyclodextrin adapter interactions
with engineered protein nanopores
Arijit Banerjeea, Ellina Mikhailovaa, Stephen Cheleya, Li-Qun Gub,2, Michelle Montoyac,3, Yasuo Nagaokaa,4,
Eric Gouauxd, and Hagan Bayleya,1
aDepartment of Chemistry, University of Oxford, Oxford, OX1 3TA, United Kingdom; bDepartment of Medical Biochemistry & Genetics, Texas A&M
University System Health Science Center, College Station, TX 77843-1114; cDepartment of Biochemistry and Molecular Biophysics and Howard Hughes
Medical Institute, Columbia University, New York, NY 10032; and dVollum Institute and Howard Hughes Medical Institute, Oregon Health and Science
University, Portland, OR 97239
Edited by Gregory A. Petsko, Brandeis University, Waltham, MA, and approved March 3, 2010 (received for review December 15, 2009)
Engineered protein pores have several potential applications in
biotechnology: as sensor elements in stochastic detection and
ultrarapid DNA sequencing, as nanoreactors to observe single-
molecule chemistry, and in the construction of nano- and micro-
devices. One important class of pores contains molecular adapters,
which provide internal binding sites for small molecules. Mutants
of the α-hemolysin (αHL) pore that bind the adapter β-cyclodextrin
(βCD) ∼104 times more tightly than the wild type have been ob-
tained. We now use single-channel electrical recording, protein en-
gineering including unnatural amino acid mutagenesis, and high-
resolution x-ray crystallography to provide definitive structural in-
formation on these engineered protein nanopores in unparalleled
detail.
alpha-hemolysin ∣single molecule ∣stochastic sensing ∣structure ∣
unnatural amino acid
M
any research groups have used protein engineering to
obtain enzymes and antibodies with new properties suited
for specific tasks (1–6). Fewer groups have taken on the difficult
problem of engineering membrane proteins (7). We have engi-
neered the α-hemolysin protein pore, mindful of several potential
applications in biotechnology, including its ability to act as a de-
tector in stochastic sensing (8) and ultrarapid DNA sequencing
(9), to serve as a nanoreactor for the observation of single-
molecule chemistry (10) and to act as a component for the con-
struction of nano- and microdevices (11).
An important breakthrough in this area, which enabled the sto-
chastic sensing of organic molecules including the detection of
DNA bases in the form of nucleoside monophosphates (12, 13),
was the discovery of internal molecular adapters, a form of non-
covalent protein modification (14). Most useful have been cyclo-
dextrin (CD) adapters, which have until now been used in the
absence of detailed structural information about how they work.
The present paper is a definitive investigation, which provides
such information through the application of a wide variety of
technical approaches: single-channel electrical recording, protein
engineering including unnatural amino acid mutagenesis, and
x-ray crystallography. The studies employing mutagenesis show
that the striking interactions seen in the crystal structures also
occur in individual pores in lipid bilayers.
We reveal that the tight-binding αHL mutants (15) M113N7
and M113F7 bind βCD in different orientations within the hep-
tameric pore. In the case of M113N7, the top (primary hydroxyls)
of the CD ring faces the trans entrance of the pore. In the case of
M113F7, the bottom (secondary hydroxyls) of the CD ring faces
the trans entrance, while the top of the ring is bonded to the pore
through remarkable CH-π interactions. Another tight-binding
mutant, M113V7, can bind the CD in both orientations. These
results illustrate the exquisite level of engineering that can be
achieved with protein nanopores, which is, for example, far be-
yond what is possible with solid-state pores. The work also pro-
vides information valuable for the design of new binding sites
within the lumen of the αHL pore or within other β-barrel pro-
teins. Our results will be of interest to others exploring the inter-
actions of CDs with the αHL pore (16, 17), including groups
involved in computational studies (18, 19). In addition CDs bind
to a variety of other pores, including porins (20, 21) and connex-
ins (22), and are being tested in vivo as blockers of the anthrax
protective antigen pore (23, 24). The CD adapter concept has
also been incorporated into other formats, e.g., with glass nano-
pores (25), and artificial pores based on CDs have been made by
several groups (26–28). Our work is pertinent to these studies.
Results
Kinetics and Thermodynamics of the Interactions of βCD with αHL
Pores Containing Met, Phe and Asn at Position 113. We showed earlier
that position 113 in the αHL pore (Fig. 1A) is critical for the bind-
ing of βCD (14). Subsequently, residue 113, which is Met in the
WT protein, was changed to each of the remaining 19 naturally
occurring amino acids by site-directed mutagenesis (15). We
found that 11 of these mutants, expressed as homoheptamers,
bound βCD with a similar affinity and with similar kinetics to
the WT homoheptamer. Two mutants (P, W) bound βCD about
10 times more strongly than the WT homoheptamer, while six of
them (V, H, Y, D, N, F) bound with high affinity, i.e., with a Kd
value 103 to 104 times lower than the WT.
Remarkably, the side chains of the latter six amino acids bear
little resemblance to one another, and this issue is addressed in the
present paper. We first examined the two amino acids with the
most disparate side chains (Fand N) by making heteromeric pores
containing WT (Met-113), M113F, and M113N subunits. Three
series of heteroheptamers were produced: WT7−nM113Nn,
WT7−nM113Fn, and M113F7−nM113Nn. The heteroheptamers
were separated by SDS-polyacrylamide gel electrophoresis aided
by an oligoaspartate (D8) tail on the first of the two types of sub-
unit (Fig. 1B) (29). All 21 combinations of WT, M113F, and
M113N subunits formed αHL pores that interacted with βCD as
shown by single-channel current recordings, which revealed the
extent of block by βCD (Fig. S1), the association and dissociation
Author contributions: A.B., S.C., E.G., and H.B. designed research; A.B., E.M., S.C., L.-Q.G.,
M.M., and Y.N. performed research; A.B., E.G., and H.B. analyzed data; and A.B. and
H.B. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
1To whom correspondence should be addressed. E-mail: hagan.bayley@chem.ox.ac.uk.
2Present address: Department of Biological Engineering and Dalton Cardiovascular
Research Center, University of Missouri, Columbia, MO 65211.
3Present address: Nature Structural & Molecular Biology, 75 Varick Street, 9th Floor, New
York NY 10013-1917.
4Present address: Department of Biotechnology, Faculty of Engineering, Kansai University,
3-3-35 Yamate-cho, Suita, Osaka 564-8680, Japan.
This article contains supporting information online at www.pnas.org/cgi/content/full/
0914229107/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.0914229107
PNAS ∣May 4, 2010 ∣vol. 107 ∣no. 18 ∣8165–8170
BIOCHEMISTRY
rate constants for βCD (kon and koff), and (from the latter) the
equilibrium dissociation constant for βCD (Kd ¼ koff∕kon) (15).
The kon values for βCD for the 21 combinations of subunits
were all similar at ∼5 × 105 M−1 s−1 (Fig. 1C, Upper). By contrast,
the koff values differed widely, ranging from ∼5 × 10−2 s−1 to
∼103 s−1. For WT7−nM113Nn and WT7−nM113Fn, the koff values
decreased as M113N or M113F subunits were added. In the case
of M113N, there was a steep drop in the value of koff after the
fifth subunit had been incorporated. In the case of M113F,
the decrease in the value of koff occurred less precipitously as the
M113F subunits were added (Fig. 1C, Lower). Intriguingly, with
M113F7−nM113Nn, koff first increased as M113N subunits were
added to M113F7 until n ¼ 4 (M113F3M113N4) and then de-
creased for larger values of n (Fig. 1C, Lower). We recognize that
there is more than one permutation of heteromers containing two
to five mutant subunits (Fig. 1B), but we have ignored this fact
here because no significant differences in the properties of indi-
vidual heteromers were observed. For example, 42 recordings
were made of WT5M113N2, which has three permutations.
Because, kon showed little variation with subunit composition,
the variation in Kd was similar to the variation in koff (Fig. 1C).
While these studies were in progress, the crystal structures of
βCD complexed to M113N7 (Fig. 2B) and M113F7 (Fig. 2C) were
solved (Table S1) (30). High-resolution structures could be
obtained because the CD and the αHL pore have the same C7
symmetry. In the case of M113N7, βCD is bound with the second-
ary hydroxyl face “upward” (Fig. 2B). In each glucose unit of the
βCD, the 2-hydroxyl is hydrogen bonded to the side-chain amide
of an Asn-113 (the residue introduced by mutagenesis) and the
3-hydroxyl is hydrogen bonded to the ϵ-amino group of Lys-147.
In the case of M113F7, two βCDs are bound to the αHL pore
(Fig. 2C). It is the top βCD in the structure that concerns us, be-
cause it is in contact with the Phe-113 residues introduced by mu-
tagenesis. It is immediately apparent that the top βCD in M113F7
is in the opposite orientation to the βCD in M113N7 with each
6-hydroxyl group in a CH-π bonding interaction (31–35) with a
Phe-113 side chain. The opposite orientations of the βCDs in
M113N7 and M113F7 immediately explain why heteromers
formed from similar numbers of M113N and M113F subunits
(e.g., M113N4M113F3) bind βCD weakly (see also Discussion).
Unnatural Amino Acid Mutagenesis. To further explore the range of
noncovalent interactions that are available when βCD binds to
the αHL pore, five unnatural amino acids (Fig. 3A and Fig. S2)
were incorporated at position 113, by using the in vitro nonsense
codon suppression method (36). In particular, we had noted that
M113V7 containing the β-branched Val binds βCD tightly (15),
and therefore we compared cyclopropylglycine (Cpg) and cyclo-
propylalanine (Cpa). We also further examined the means by
which M113F7 binds βCD tightly, by comparing the properties of
4-fluorophenylalanine (f1F), pentafluorophenylalanine (f5F),
and cyclohexylalanine (Cha) at position 113.
The five homomeric pores all produced single-channel cur-
rents with unitary conductance values in the range expected
for properly assembled heptamers (Fig. S3). All five bound βCD
(Fig. 3B, Level 2), either tightly (f1F, Cpg) or weakly (f5F, Cha,
Cpa) as described in detail below. During the long βCD binding
events, additional current spikes were seen (Fig. 3B). Similar
Fig. 1.
Binding of βCD by heteromeric αHL pores formed by WT, M113F and M113N subunits. (A) Crystal structure of WT-αHL (61) showing residue 113 (Met,
yellow). Left panel, side view and right panel, top view. (B) Separation of 35S-labeled αHL heteroheptamers by SDS-polyacrylamide electrophoresis. The
separation of the M113F7-nM113Nn heteromers is shown as detected by autoradiography of a dried gel. The M113F subunits carried a D8 tail. Lane 1,
M113N7; lane 2, M113F7−nM113Nn (the heteromers formed from several preparations made with differing ratios of M113F and M113N subunits were mixed
to give roughly equal amounts of each subunit combination); lane 3, M113F7. A diagram of the eight different combinations of subunits and their permuta-
tions is shown to the right of the autoradiogram. The various permutations are not separated by electrophoresis. (C) Kinetics of the interaction of βCD with
single heteromeric αHL pores as determined by bilayer recording. Values of kon were calculated by using kon ¼ 1∕ðτon½βCDÞ, where τon is the mean interevent
interval. Values of koff were determined by using koff ¼ 1∕τoff, where τoff is the mean dwell time of βCD in the pore. Values of Kd were calculated by using
Kd ¼ koff∕kon. Each point represents the mean s:d: for three or more determinations. Where they cannot be seen, the s.d. values lie within the symbol. Black
squares, WT7−nM113Nn; gray squares, M113F7−nM113Nn; empty squares, M113F7−nWTn.
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Banerjee et al.
events had been observed previously with certain Met-113 repla-
cement mutants and may represent movement of the βCD at its
binding site (e.g., rotation about axes perpendicular to the C7
axis) (15). The additional current spikes were more prevalent
for M113V7 and M113Cpg7, which may take part in more con-
formationally labile interactions with βCD, compared with say
M113F7 (Fig. S4).
Interactions of βCD with Homoheptamers Bearing Aromatic Residues
at Position 113. To further understand the nature of the binding of
βCD to aromatic side chains, we examined the kinetics of βCD
binding to the homoheptamers containing f1F or f5F at position
113, M113f1F7 and M113f5F7 (Fig. 3C). For both mutants, the
value of kon was very similar to that of WT7, but the values of koff
and therefore Kd for M113f1F7 differed dramatically from WT7
and were close to the values for the tight-binding mutant M113F7
(Table S2A). By contrast, koff and Kd for M113f5F7 were similar
to the values for WT7 (Table S2A).
To determine whether M113f1F7 binds βCD in the same orien-
tation as M113F7 (Fig. 2C), we made heteromers of the M113f1F
subunit with M113N or M113F and examined M113F4M113f1F3
and M113N4M113f1F3. M113F4M113f1F3 binds βCD as strongly
as either M113F7 or M113f1F7, but M113N4M113f1F3 binds
βCD weakly with a similar affinity to WT7 (Fig. 3D and
Table S3). Therefore, it is reasonable to infer that M113F7
and M113f1F7 bind βCD in the same orientation with the 6-
hydroxyl groups of the CD in proximity to the aromatic rings
on the protein.
Cyclohexylalanine (Cha) was used to replace the aromatic side
chains with a roughly isosteric hydrophobic group. Again the va-
lue of kon for βCD was little changed, but koff for M113Cha7 had
an intermediate value of 42 6 s−1. Therefore, M113Cha7 binds
βCD more weakly than M113F7 but distinctly more strongly than
the WT7 pore (Table S2A and Fig. 3C).
Interactions of βCD with Homoheptamers Bearing Hydrophobic Resi-
dues at Position 113. M113V7 binds βCD very strongly, and there-
fore we compared αHL pores with Cpg or Cpa at position 113.
Cpg is roughly isosteric with Val, and like Val has a β-branched
side chain. Gratifyingly, M113Cpg7 has a kon value similar to the
other αHL pores, and koff and Kd values close to those of
M113V7 (Table S2B and Fig. 3E). Cyclopropylalanine (Cpa), with
an additional methylene group compared to Cpg, is roughly
isosteric with Leu, a weak binder, and M113Cpa7 also binds
βCD weakly with kon, koff and Kd values similar to those of
WT7 (Table S2B and Fig. 3E). M113I7 and M113T7, which are
β-branched, are also weak binders, but Ile and Thr are less closely
related to Val than Cpg.
To determine whether M113V7 binds βCD in the same orien-
tation as M113F7 or M113N7 (Fig. 2), we made heteromers of
M113V and the M113N or M113F subunits. M113V3M113F4,
M113V4M113F3, M113V3M113N4, and M113V4M113N3 were
examined in detail. All four heteroheptamers bound βCD more
weakly than M113V7, M113F7 or M113N7 (Fig. 3F and Table S4),
suggesting that Val at position 113 interacts with βCD strongly but
in a different manner to either Phe or Asn. Each heteromer
exhibited a range of Kd values, perhaps reflecting the various pos-
sible permutations of the two different subunits around the cen-
tral axis of the heptamer, although this heterogeneity was not
seen for heteromers made from WT, M113F and M113N (Fig. 1).
Discussion
Soon after we discovered that βCD binds to the WT-αHL pore for
around a millisecond, we found a mutant pore, M113N7, that re-
leases βCD ∼104 times more slowly (14). This prompted us to
examine all 19 mutants in which residue 113 is replaced by a nat-
ural amino acid, with the surprising result that a collection of ami-
no acids with structurally unrelated side chains (V, H, Y, D, N, F)
are tight binders (15). Here, we have examined the nature of the
binding interactions more closely by single-channel electrical re-
cording, protein engineering including unnatural amino acid mu-
tagenesis, and high-resolution x-ray crystallography, and we
provide the first definitive structural information on an engi-
neered protein nanopore.
We find that βCD can bind tightly to the αHL pore in three
different ways depending on the residue at 113, as exemplified
by Asn, Phe, and Val. Because Asn and Phe have quite different
side chains, we first compared the ability of M113N and M113F
subunits to take part in binding the CD. The examination of het-
eromeric proteins containing WT (Met-113), M113N and M113F
subunits showed that the replacement of WT subunits in WT7
with M113N or M113F subunits led to increased affinity for
βCD. The more M113N or M113F subunits that were added, the
tighter binding became. By contrast, when subunits in M113N7
were replaced with M113F subunits, binding became weaker,
reaching a minimum at three to four M113F subunits, and then
increasing in strength with five M113F subunits or more (Fig. 1C).
Parallel structural studies (30) revealed the basis of the “oppos-
ing” effects of the M113N and M113F subunits. βCD binds to
M113N7 in the opposite orientation to that in which it binds
to M113F7. In M113N7, the secondary hydroxyls in the βCD ring
are hydrogen bonded to Lys-147 and Asn-113 (Fig. 2A). By con-
trast, βCD interacts with M113F7 through its primary hydroxyl
face (Fig. 2B).
It seemed likely that M113V7, bound βCD in yet another way,
and this was examined by forming heteromers between M113V
and M113N or M113F. The presence of three or four subunits
of either M113N or M113F greatly decreases the affinity of
the pore for βCD (Fig. 3F), with an average koff of 7.3 × 101 s−1,
indicating that a third binding mode is indeed operating
Fig. 2.
X-ray structures of M113N and
M113F homoheptamers with βCD bound.
(A) Side view of heptameric αHL. βCD binds
in the blue highlighted region. (B) βCD
bound to M113N7 (dotted lines indicate hy-
drogen bonding). The side chains of Lys-147
are in pale brown and the side chains of Asn-
113 in yellow. (C) βCD bound to M113F7
(dotted lines indicate CH-π bonding). The
side chains of Phe-113 are in yellow. The sec-
ond βCD in the M113F7 · ðβCDÞ2 structure is
hydrogen bonded to the top βCD in a head-
to-head arrangement and has no apparent
interactions with the protein. For both (B)
and (C), four β strands were omitted from
the barrel to give a better view.
Banerjee et al.
PNAS
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vol. 107
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8167
BIOCHEMISTRY
(Table S4). In summary, the three groups of tight-binding mutants
comprise αHL pores incorporating, at position 113: (i) the hydro-
gen-bonding amino acids N, D (the latter would have to be largely
in the protonated form), and possibly H; (ii) the aromatics F, Y,
f1F, and possibly H, and more weakly W; (iii) the β-branched ami-
no acids V, Cpg. There may be yet other means by which CDs can
bind to the αHL pore. For example, we earlier found that hepta-
6-sulfato-βCD can bind tightly to αHL pores containing the
N139Q mutation (37). Presumably, this CD is bound at a site low-
er down in the β barrel in a fashion that includes hydrogen bond-
ing to the Gln at position 139. While the various mutants
exhibited widely different koff values, the value of kon was almost
invariant and averaged ∼2.3 × 105 M−1 s−1 (Table S2) (15). Ap-
parently, transport to the binding site is rate limiting, through
a route unaffected by mutagenesis.
koff increased precipitously with the addition of WTsubunits to
M113N7 (Fig. 1C). Crystal structures of M113N7 show that resi-
dues
111,
113,
and
147
are
reorganized
by
compari-
son with WT7 and then undergo a more limited rearrangement
when βCD binds (Fig. S5). For example, the side chain of
Lys-147 shifts position to form a bifurcated hydrogen bond with
a 3-hydroxyl group of βCD and the side chain carbonyl of an Asn-
113 (Fig. S6). Therefore, the side chains of residues 111, 113, and
147 might be in a variety of conformations in WT7−nM113Nn het-
eromers and offer less well preorganized binding sites for βCD
than they do in M113N7. Further, the intramolecular hydrogen
bonds of the secondary hydroxyls in βCD (38) must be disrupted
upon binding as both hydroxyls on each glucose ring form hydro-
gen bonds to the mutant subunits (Fig. 2B). Because the hydrogen
bonds that are broken in βCD are arranged in a circle, the break-
age of bonds involving a single glucose (three bonds in all) will be
relatively more disruptive than those involving adjoining glucose
residues or the entire circle. The overall binding cooperativity in
M113N7 could be attributed to enthalpic cooperativity outweigh-
ing entropic penalties to binding (39). Positive cooperativity has
been observed previously in fairly rigid model systems (40).
By contrast with M113N7, there is little movement of side
chains in ðM113FÞ7 by comparison with WT7 and little move-
ment, including Phe-113, upon binding βCD (Fig. S7A). Further,
the crystal structure of M113F7 · βCD suggests that each Phe re-
sidue interacts independently with the βCD through what appear
to be CH-π interactions (Fig. S7B). These interactions are ex-
pected to be weak and not strongly directional and hence offer
less enthalpic cooperativity, as supported by the B-factors (crys-
tallographic temperatures factors) at the primary βCD binding
site, which are between ∼40 and 50. Positive cooperativity is ob-
served, but it is less pronounced than in the case of M113N7
(Table S5). In the case of M113N7, the B-factors of the residues
that bind βCD are in the 20s implying that the βCD is more rigidly
held than it is in M113F7.
The binding of sugars to aromatic residues in proteins can in-
clude CH-π bonding (41) or OH-π bonding or a finely balanced
complement of both (42, 43). However, we have dismissed the
possibility of an OH-π interaction between Phe-113 and the
6-hydroxyl groups of βCD as the distance between the center
of the phenyl rings to the nearest hydroxyl oxygen is higher
(5.2 0.65 Å, n ¼ 7) than that expected for a favorable OH-π
interaction (33). While we propose that βCD binds to Phe-113
through a C-6 CH-π interaction (Fig. S7B), the distances between
the center of the Phe-113 ring and the nearest C-6 of βCD ob-
served in the M113F7 · βCD structure (4.66 0.24 Å, n ¼ 7)
are in the upper range of the expected distance for a strong inter-
action, which is ∼4.5 Å (33). The angle between the normal to the
aromatic rings and the line connecting the C-6 atoms to the aro-
matic midpoint is 8.0 5.6°, which is well within the expected
range (44). The measurements with M113f5F7 argue against a
hydrophobic interaction between Phe residues at position 113
and the βCD ring. In f5F, the hydrophobicity of the phenyl ring
is significantly increased (45) yet M113f5F7 binds βCD weakly,
like WT7 (Fig. 3C and Table S2A).
By contrast with F, f1F, Y and N, homomeric αHL pores with
f5F and W at position 113 bound βCD relatively weakly (Fig. 3C
and Table S2A). In the case of f5F, the powerful electron with-
drawing action of the five fluorine atoms leaves a highly increased
positive charge at the center of the ring (46, 47), mitigating
against a hydrogen-bonding interaction. The electron-rich Trp
Fig. 3.
Properties of pores containing natural and unnatural amino acid sub-
stitutions at position 113. The data were recorded at þ40 mV in 1.0 M NaCl,
10 mM sodium phosphate, pH 7.5. (A) Unnatural amino acids used in this
study: 4-fluorophenylalanine, f1F; pentafluorophenylalanine, f5F; cyclohex-
ylalanine, Cha; cyclopropylglycine, Cpg; cyclopropylalanine, Cpa. (B) Repre-
sentative
current
traces
from
single
homoheptameric
αHL
pores,
containing unnatural amino acids at position 113, in the presence of βCD.
βCD (40 μM final) was added to the trans chamber. Level 1, open pore current;
level 2, pore occupied by βCD. The broken line indicates zero current. (C) In-
teraction of βCD with homomeric αHL pores containing aromatic amino acids
at position 113. Kd values for the interaction between βCD and the αHL pore
were calculated by using Kd ¼ koff∕kon. Each column represents the mean
s:d: for 10 or more determinations: dark gray, natural amino acids; light gray,
unnatural amino acids. Data adapted from Gu and colleagues (15) are
marked (*). (D) Representative current traces from single-channel recordings
of βCD binding to M113F4M113f1F3 and M113N4M113f1F3. βCD (40 μM final)
was added to the trans chamber. The broken line indicates zero current. (E)
Interaction of βCD with homomeric αHL pores containing hydrophobic amino
acids at position 113. Kd values for the interaction between βCD and the αHL
pore were calculated by using Kd ¼ koff∕kon. Each column represents the
mean s:d: for ten or more determinations: dark gray, natural amino acids;
light gray, unnatural amino acids. Data adapted from Gu and colleagues (15)
are marked (*). (F) koff values for βCD from heteroheptamers formed with
M113F and M113V subunits and with M113N and M113V subunits. βCD
(40 μM final) was added to the trans chamber. The kon values for βCD for
all these mutants are similar, at ∼3 × 105 M−1 s−1. Empty square: average
koff values for the mutant (bar is s:d). Filled square: M113V3M113F4; filled
circle: M113V4M113F3; filled upright triangle: M113V3M113N4; filled in-
verted triangle: M113V4M113N3.
8168
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www.pnas.org/cgi/doi/10.1073/pnas.0914229107
Banerjee et al.
ring (44, 46, 47) should favor hydrogen bonding, but here we can-
not make a direct comparison with the crystal structure of
M113F7 as the indole ring is far larger than benzene. It is possible
that it cannot become oriented in the same manner and that it is
misaligned for hydrogen bonding.
Our experiments suggest that M113V7 and M113Cpg7 bind
βCD in a third way. In heteromers with M113V, both M113F
and M113N reduce the affinity of the pore for βCD suggesting that
neither the CH-π interaction with Phe-113 nor the hydrogen-
bonding interactions with Asn-113 and Lys-147 are compatible
with binding to Val. Close interactions of Val with glucose rings
have been noted previously (48). Therefore, we propose that the
Val side-chain interacts with the side of the glucose ring. This in-
teraction might occur in one or both orientations of the CD
ring (Fig. 4).
Conclusion
We provide structural information on engineered protein nano-
pores and describe three distinct ways in which βCD can bind
within the lumen of mutant αHL pores in atomic detail. Our re-
sults will be useful in several areas of basic science and biotech-
nology. By using host molecules lodged within the αHL pore,
host-guest interactions can be investigated in fine detail at the
single-molecule level (17, 49). The present work will now permit
us to examine binding events at a known face of a CD. The work
also provides information for designing new binding sites within
the lumen of the αHL pore (37) or within other β barrel proteins
(21, 50) and for using molecular design to devise ways in which to
covalently attach CDs within pores (13, 51). These areas impact
practical applications of nanopore technology including stochas-
tic sensing (8), single-molecule DNA sequencing (9, 12, 13, 52),
the use of nanoreactors for the observation of single-molecule
chemistry (10), and the construction of nano- and microdevices
(11, 53), as well as the design of CDs as therapeutic agents
(23, 24).
Methods
Full details of the experimental procedures can be found in SI Appendix.
Materials
L-Amino acids were obtained as follows: 4-fluorophenylalanine (f1F) (Fluka);
pentafluorophenylalanine (f5F) (PepTech Corp.); cyclopropylglycine (Cpg) (Ty-
ger); cyclopropylalanine (Cpa) (Tyger). 4-N-benzoyl-5′-O-(4,4′-dimethoxytri-
tyl)-2′-deoxycytidine-3′-[(2-cyanoethyl)-(N,N-diisopropyl)]-phosphoramidite
and bis(2-cyanoethyl)-N,N-diisopropylphosphoramidite for the synthesis of
pdCpA were purchased from Glen Research and Toronto Research Chemicals,
respectively.
Preparation
of
NVOC-Protected
Aminoacyl-pdCpA.
NVOC-protected
aminoacyl-pdCpAs were prepared as reported previously by reacting the
dinucleotide pdCpA with N-protected, carboxylic acid-activated, amino
acids (54–56).
Preparation of NVOC-Protected Aminoacyl-tRNA. NVOC-protected aminoacyl-
pdCpAs were ligated enzymatically with a truncated tRNA, prepared by using
methods described elsewhere (57, 58).
Genetic Constructs and Mutagenesis. All new αHL constructs were verified by
DNA sequencing. Details of each construct can be found in SI Appendix.
Synthesis, Assembly, and Purification of Mutant αHL pores. αHL monomers (WT
and mutants) were prepared in vitro by coupled transcription and translation
(IVTT) and assembled into homoheptamers on rabbit red blood cell
membranes followed by purification by SDS–PAGE as described earlier
(59). Heteroheptamers were prepared by mixing the two required DNAs
(one encoding an αHL with a D8 tail) before IVTT and then oligomerizing
the mixed translation products on rabbit red blood cell membranes. Pores
with the desired combinations of subunits were purified by SDS–PAGE (59).
Synthesis, Assembly, and Purification of αHL Mutants Containing Unnatural Ami-
no Acids. αHL polypeptides containing unnatural amino acids were synthe-
sized by IVTT in the presence of rabbit red blood cell membranes. The
plasmid with a stop codon (TAG) at position 113 was used. Deprotected ami-
noacyl-tRNAs (SI Appendix) were added to the IVTT mixtures. For heterohep-
tamers with subunits containing unnatural amino acids in combination with
M113N or M113F, monomers were first made, which were then coassembled
on rabbit red blood cell membranes and subsequently purified by SDS–PAGE.
Single-Channel Current Recordings in Planar Lipid Bilayers. (15, 60) Recordings
were made with 1.0 M NaCl, 10 mM sodium phosphate, pH 7.5, in both cham-
bers, at an applied potential of þ40 mV. Data were recorded at 22 2°C. The
bilayer was formed from 1,2-diphytanoyl-sn-glycero-phosphocholine (Avanti
Polar Lipids). Proteins were added to the cis chamber, and βCD to the trans
chamber. Single-channel currents were recorded with an Axopatch 200B
patch-clamp amplifier (Axon Instruments) and filtered at 2 kHz with a
built-in 4-pole low-pass Bessel Filter. The data were acquired at a sampling
rate of 10 kHz. For mutants that bind βCD strongly, the data were acquired
for at least 30 min and for weak-binding mutants for at least 10 min.
Kinetic Data Analysis. Current amplitude and dwell-time histograms were
made by using ClampFit 9.0. The mean dwell times, τoff, were determined
by fitting the dwell-time histograms to single exponentials. Values of kon
and koff were obtained by using the mean dwell times and mean interevent
intervals, as described previously (15, 60). This analysis assumes a binary in-
teraction, which was supported in all cases examined by the finding of only
one major blockade level and a single exponential distribution of dwell
times (τoff).
Fig. 4.
Molecular model showing the three classes of interaction between
the αHL pore and βCD identified in this work. The model identifies the region
of βCD responsible for each interaction (H atoms interacting with Phe-113 or
Asn-113 and Lys-147: gray). The first class of interaction is with aromatic
residues and involves the seven -CH2OH groups of the βCD. The second class
is typified by the interactions with Asn at position 113, which involve hydro-
gen-bonds to the secondary 2-hydroxyls of the βCD. Structural studies show
that this interaction is supported by hydrogen bonding between Lys-147 and
the secondary 3-hydroxyls of the βCD. Structural studies and experiments
with heteromers suggest that the βCD in M113F7 is in the opposite orienta-
tion to the βCD in M113N7, in support of the model shown here. As the inter-
action with Val is hydrophobic, it is not directional and βCD may not bind at
the same position inside the β barrel as it does in M113F7 or M113N7.
Banerjee et al.
PNAS
∣
May 4, 2010
∣
vol. 107
∣
no. 18
∣
8169
BIOCHEMISTRY
Protein Crystallography. Details can be found in SI Appendix. Protein Data
Bank: The coordinates and structure factors of the described structures have
been deposited with accession codes 3M2L ðM113F7Þ, 3M3R ðM113F7 · βCDÞ,
3M4D ðM113N7Þ, 3M4E ðM113N7 · βCDÞ.
ACKNOWLEDGMENTS. We thank Dennis Dougherty for the plasmid pTHG73.
This work was funded by a Royal Society Wolfson Research Merit Award
(to H.B.), the Medical Research Council (H.B.), the National Institutes of
Health (H.B.), and the Howard Hughes Medical Institute (E.G.).
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Banerjee et al.
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3M4G
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H57A HFQ from Pseudomonas Aeruginosa
|
structural communications
760
doi:10.1107/S1744309110017331
Acta Cryst. (2010). F66, 760–764
Acta Crystallographica Section F
Structural Biology
and Crystallization
Communications
ISSN 1744-3091
The structures of mutant forms of Hfq from
Pseudomonas aeruginosa reveal the importance of
the conserved His57 for the protein hexamer
organization
Olga Moskaleva, Bogdan Melnik,
Azat Gabdulkhakov, Maria
Garber, Stanislav Nikonov, Elena
Stolboushkina and Alexei
Nikulin*
Institute of Protein Research, RAS, Russia
Correspondence e-mail: nikulin@vega.protres.ru
Received 12 March 2010
Accepted 11 May 2010
PDB References: P. aeruginosa Hfq, H57A
mutant, 3inz; H57T mutant, 3m4g.
The bacterial Sm-like protein Hfq forms homohexamers both in solution and in
crystals. The monomers are organized as a continuous -sheet passing through
the whole hexamer ring with a common hydrophobic core. Analysis of the
Pseudomonas aeruginosa Hfq (PaeHfq) hexamer structure suggested that
solvent-inaccessible intermonomer hydrogen bonds created by conserved
amino-acid residues should also stabilize the quaternary structure of the
protein. In this work, one such conserved residue, His57, in PaeHfq was replaced
by alanine, threonine or asparagine. The crystal structures of His57Thr and
His57Ala Hfq were determined and the stabilities of all of the mutant forms and
of the wild-type protein were measured. The results obtained demonstrate the
great importance of solvent-inaccessible conserved hydrogen bonds between the
Hfq monomers in stabilization of the hexamer structure.
1. Introduction
In bacteria, Hfq protein acts as a global post-transcriptional regulator
which binds small regulatory RNAs and promotes their interaction
with mRNAs (Valentin-Hansen et al., 2004; Brennan & Link, 2007).
It controls the expression of many genes by its action on mRNA
translation, stability or polyadenylation (Zhang et al., 1998; Vytvytska
et al., 2000; Hajnsdorf & Re´gnier, 2000; Sledjeski et al., 2001). Hfq is a
small (70–110 amino-acid residues) thermostable protein which exists
in a homohexameric form in solution (Brennan & Link, 2007; Zhang
et al., 2002; Møller et al., 2002). A hexameric organization has also
been observed in the crystal structures of Hfq from Staphylococcus
aureus (Schumacher et al., 2002), the core part of Escherichia coli Hfq
(Sauter et al., 2003) and Hfq from Pseudomonas aeruginosa (Nikulin
et al., 2005). All of these proteins formed doughnut-shaped rings with
outer and inner diameters of about 65 and 10 A˚ and a thickness of
25–30 A˚ .
Hfq belongs to the Sm/Sm-like protein family. This family includes
eukaryotic Sm and Sm-like (Lsm) proteins and archaeal Lsm proteins
(Wilusz & Wilusz, 2005). Eukaryotic Sm/Lsm proteins are involved in
RNA processing in the cell (Kufel et al., 2004; Verdone et al., 2004). In
crystals they form heptamers (Achsel et al., 1999; Mayes et al., 1999;
Walke et al., 2001) or octamers (Naidoo et al., 2008), whereas in
cytoplasm the Sm proteins are found as heterodimers or trimers and
are only able to form heptamers in the presence of U-rich small
nuclear RNAs (UsnRNAs; Achsel et al., 1999; Will & Lu¨hrmann,
2001).
Archaeal genomes usually encode one or two distinct Lsm proteins
called Lsm1 and Lsm2 (Salgado-Garrido et al., 1999). Lsm3 proteins
have only been identified in a few archaeal species (Mura et al., 2003;
Kilic et al., 2006). Lsm1 is the most abundant of the archaeal species.
Archaeal Lsm1 and Lsm2 proteins form stable homoheptamers, with
the exception of Archaeoglobus fulgidus AF-Sm2, which can exist in
hexameric or heptameric forms depending on the pH or the presence
of RNA (To¨ro¨ et al., 2001; Achsel et al., 2001; Kilic et al., 2006).
# 2010 International Union of Crystallography
All rights reserved
The Sm-protein family is characterized by a conserved motif of
about 70 amino acids, which is called the Sm-domain. It is a -barrel-
type structure consisting of five -strands, which are capped by an
N-terminal -helix (Fig. 1a). The Sm-domain contains two conserved
sequence motifs (Sm1 and Sm2) linked by a loop that differs in length
and sequence depending on the species (Valentin-Hansen et al., 2004;
Se´raphin, 1995). Strands 1, 2 and 3 form the Sm1 motif and
strands 4 and 5 constitute the Sm2 motif of the domain. The
sequence of the Sm1 motif is conserved among all bacteria, archaea
and eukarya (Kambach et al., 1999; Hajnsdorf & Re´gnier, 2000;
Valentin-Hansen et al., 2004; Kilic et al., 2006). In contrast, the Sm2
motif has different consensus sequences in bacterial Hfq and
eukaryal/archaeal Sm/Lsm proteins (Sauter et al., 2003). Analysis of
the known crystal structures (To¨ro¨ et al., 2002; Nikulin et al., 2005;
Brennan & Link, 2007) has shown that the -strands of the Sm2
motifs organize the protein ring by means of hydrogen bonds formed
by the main-chain O and N atoms. The quaternary structure is
additionally stabilized by contacts between strongly conserved amino
acids. Previously, we have suggested (Nikulin et al., 2005) that the
conserved YKHI consensus sequence of the Sm2 motif in Hfq should
define its hexamer formation and that His57 could play a very
important role in the stabilization of the hexamer structure. To prove
this hypothesis, we mutated His57 in P. aeruginosa Hfq (PaeHfq) to
alanine, threonine and asparagine, measured the stability of the wild-
type hexamer and the obtained mutant forms and solved the crystal
structures of PaeHfq with His57Thr and His57Ala mutations.
2. Materials and methods
2.1. Site-directed mutagenesis, gene expression and recombinant
protein purification
To prepare mutant forms of PaeHfq, site-directed mutagenesis was
carried out by PCR using oligonucleotides which contained the
desired mutations. All of the mutants (hfqH57A, hfqH57N and
hfqH57T) were constructed in two steps. In step 1, fragments carrying
a mutation were amplified from pET22b(+)/Hfq DNA by PCR with
the reverse primer 50-CGGGATCCTCAAGCGTTGCCC-30 and
corresponding oligonucleotides for each fragment (H57A, 50-GTT-
TACAAGGCGGCGATCTCC-30;
H57N,
50-GTTTACAAGAAC-
GCGATCTCC-30; H57T, 50-GTTTACAAGACCGCGATCTCC-30).
The fragments were then completed by PCR using the forward
primer 50-GGGAATTCCATATGTCAAAAGGGCAT-30 and the
PCR products obtained in step 1. The final PCR products were
inserted into pET22b(+) plasmid DNA and verified by sequencing.
All of the mutant proteins were purified as described previously
(Nikulin et al., 2005).
2.2. Circular-dichroism (CD) measurements
CD measurements were performed on a Jasco J600 spectro-
polarimeter equipped with a Julabo F25 computer-controlled
thermostat. All spectra and melting experiments were measured
structural communications
Acta Cryst. (2010). F66, 760–764
Moskaleva et al.
Hfq
761
Figure 1
(a) Overall structure of the Hfq hexamer from P. aeruginosa. One monomer is coloured according to the conserved sequence motifs: the Sm1 motif (1, 2 and 3) is shown
in yellow, the Sm2 motif (4 and 5) in red and the N-terminal 1 helix in blue. The position of amino-acid residue 57 is shown by a black sphere. (b) Superposition of wild-
type PaeHfq (cyan), H57T PaeHfq (magenta) and H57A PaeHfq (green). The C-atom r.m.s. deviations of H57T PaeHfq and H57A PaeHfq from the wild-type protein are
0.43 and 0.49 A˚ , respectively. (c) The amino-acid sequence of PaeHfq with corresponding secondary-structure elements. Amino-acid residues that are conserved in Lsm
proteins from bacteria, archaea and eukarya are shown in green; those conserved in bacteria only are shown in cyan.
structural communications
762
Moskaleva et al.
Hfq
Acta Cryst. (2010). F66, 760–764
Figure 2
The interface of two adjacent monomers in the PaeHfq hexamer. The main chains of the monomers are shown in green and yellow. Side chains are shown for residue 57 only.
Hydrogen bonds are shown as dotted lines. (a) The wild-type PaeHfq crystal structure. (b) The H57A PaeHfq crystal structure. (c) The H57T PaeHfq crystal structure.
using a cell with a 0.1 mm path length. The melting experiments were
performed by monitoring the change in ellipticity at 220 nm.
2.3. Crystallization and data collection
Protein crystals were obtained using the hanging-drop vapour-
diffusion technique at 295 K. All drops were set up by mixing 2.0 ml
protein solution (8 mg ml1 protein, 100 mM NaCl, 50 mM Tris–HCl
pH 8.0) with 2.0 ml reservoir solution (200 mM NH4Cl, 15% PEG
MME 2000, 50 mM Tris–HCl pH 8.5, 20 mM CdCl2 or ZnCl2).
Crystals appeared after 1 d and reached maximum dimensions of
300 100 50 mm within one week. Before freezing, the crystals
were transferred to 15% PEG MME 2000, 15% PEG 400, 200 mM
ammonium chloride, 50 mM Tris–HCl pH 8.5. X-ray diffraction data
were collected from the crystals on EMBL beamline X12 (DESY,
Hamburg) or the BL14.1 beamline at BESSY (Berlin) and were
processed using XDS (Kabsch, 2010). Detailed data-collection
statistics are given in Table 1.
2.4. Structure determination and refinement
The protein structures were solved by the molecular-replacement
method using the PHENIX package (Adams et al., 2002) with a
hexamer of wild-type PaeHfq as the initial model (PDB code 1u1s;
Nikulin et al., 2005). The simulated-annealing protocol following
conventional residual refinement in combination with manual
inspection in Coot (Emsley & Cowtan, 2004) was used to refine the
model. Water molecules were introduced into the model using the
‘water pick’ function of Coot and the highest peaks in the Fo Fc
map were assigned to ions. At the final stage anisotropic ADP
refinement of H57T PaeHfq was implemented, improving the R and
Rfree factors from 0.199 and 0.244 to 0.149 and 0.218, respectively. The
structure coordinates of H57A PaeHfq and H57T PaeHfq have been
deposited in the Protein Data Bank (PDB codes 3inz and 3m4g,
respectively).
3. Results and discussion
3.1. Crystal structures of H57A PaeHfq and H57T PaeHfq
The crystal structures of H57A PaeHfq and H57T PaeHfq were
solved and refined to 2.05 and 1.7 A˚ resolution, respectively (Table 1).
The substitutions did not change the overall shape of the hexamer or
the conformations of the monomers (Fig. 1b).
In the wild-type protein the side chain of His57 formed two
hydrogen bonds to the main-chain O atoms of the adjacent monomer
(Fig. 2a). We supposed that the mutations would result in the
disappearance of one or both of these hydrogen bonds. Indeed, the
substitution of His57 by alanine led to a loss of the hydrogen bonds
(Fig. 2b). In contrast, the replacement of His57 by threonine gave rise
to the formation of new hydrogen bonds between adjacent monomers
that replaced those in the wild-type protein. Two water molecules
acted as bridges connecting the hydroxyl of the threonine of one
monomer to the main-chain carbonyl O atoms of Thr57 and Ile59 of
another molecule (Fig. 2c). Nevertheless, the compensation was not
completely equivalent. In the wild-type protein one of the hydrogen
bonds formed by His57 is inaccessible to solvent, whereas in H57T
PaeHfq the water-bridge hydrogen bonds are accessible. In this case
the protein atoms could easily form new hydrogen bonds to solvent.
At higher temperature the water molecules could even escape from
their sites. In this case, the hydroxyl group of Thr57 could be posi-
tioned at a short distance from the two carbonyl O atoms of the
neighbouring monomer, which is not desirable. To prove this
hypothesis, we measured the stability of the Hfq mutant proteins.
3.2. Stability of the Hfq mutant forms
To evaluate the influence of the His57 substitutions on PaeHfq
hexamer stability, CD spectra of the wild-type protein and its mutant
forms were measured. At room temperature all these proteins had
similar spectra corresponding to an / structure (Fig. 3a). It was
found that wild-type PaeHfq possesses extreme stability: its CD
spectrum did not change during heating to 366 K or on the addition of
urea up to 8 M. Difference scanning calorimetric experiments showed
that the denaturation peak of wild-type PaeHfq appeared near 393 K
(V. V. Filimonov, personal communication). Therefore, PaeHfq has
one of the highest denaturation temperatures of known proteins
(Tanaka et al., 2006). The secondary structure of wild-type PaeHfq, as
well as those of its H57A, H57T and H57N mutants, was completely
destroyed in the presence of 5 M GdnHCl (Fig. 3a). The GdnHCl-
induced unfolding of the proteins under equilibrium conditions
demonstrated that all of the substitutions changed the stability of the
protein considerably but in a similar way (Fig. 3b).
structural communications
Acta Cryst. (2010). F66, 760–764
Moskaleva et al.
Hfq
763
Figure 3
(a) CD spectrum of wild-type and mutant (H57A, H57T, H57N) PaeHfq proteins
under nondenaturing conditions (lower lines) and in the presence of 5 M GdnHCl
(upper lines). (b) Relative change of ellipticity at 220 nm during equilibrium
unfolding of the proteins by GdnHCl. (c) Relative change of ellipticity at 220 nm
during temperature unfolding of the mutant proteins in the presence of 1 M
GdnHCl.
Table 1
Data-collection and refinement statistics.
Values in parentheses are for the highest resolution shell.
H57T PaeHfq
H57A PaeHfq
Macromolecule details
PDB code
3inz
3m4g
No. of residues per monomer
82
82
Molecular assembly
Hexamer
Hexamer
Molecular weight of the hexamer (Da) 54411
54489
Data-collection statistics
Wavelength (A˚ )
1.00
0.91841
Resolution range (A˚ )
30.0–1.7 (1.74–1.7)
30.0–2.05 (2.16–2.05)
Space group
P21212
P1
Unit-cell parameters (A˚ , )
a = 61.3, b = 71.2,
c = 104.4,
= = = 90
a = 66.5, b = 66.6,
c = 68.7, = 91.8,
= 115.3, = 119.9
Total reflections
337107 (10581)
234040 (34408)
Unique reflections
50597 (2983)
53606 (7783)
Redundancy
6.7 (3.5)
4.4 (4.4)
Completeness (%)
94.1 (80.4)
97.4 (96.5)
Rmerge (%)
4.0 (36.2)
5.5 (49.0)
Average I/(I)
27.9 (3.9)
14.4 (3.0)
Wilson B factor (A˚ 2)
31.2
32.9
Refinement statistics
Resolution (A˚ )
30.0–1.70 (1.74–1.70) 30.0–2.05 (2.09–2.05)
Completeness (%)
94.1 (80.4)
97.4 (96.5)
Reflections
50571 (2983)
53561 (2704)
Test reflections
2528 (131)
2725 (127)
Rwork (%)
0.149 (0.178)
0.194 (0.296)
Rfree (%)
0.218 (0.247)
0.261 (0.393)
No. of waters
379
331
No. of ions
11
18
R.m.s. deviation from ideal geometry
Bonds (A˚ )
0.010
0.005
Angles ()
1.315
0.903
Chirality ()
0.103
0.059
Planarity ()
0.006
0.004
Average B value (A˚ 2)
Main chain
31.39
45.24
Side chain and water
38.74
50.82
MolProbity results
Ramachandran favoured (%)
95.20
95.76
Ramachandran allowed (%)
98.74
99.88
Ramachandran outliers (%)
1.26
0.12
To reveal the difference in stability of the PaeHfq mutants, the
relative changes in ellipticity at 220 nm were measured during
temperature unfolding (Fig. 3c). The presence of 1 M GdnHCl in the
buffer was important in order to melt the proteins within the oper-
ating range of the spectropolarimeter. The H57N, H57A and H57T
mutant forms of PaeHfq had melting temperatures of 346, 343 and
341 K, respectively, whereas wild-type PaeHfq retained its structure
up to 366 K. Compared with the other mutants, the H57T PaeHfq
had the lowest melting temperature, which was accompanied by a
deterioration of melting-process cooperativity. The reason for this
behaviour of H57T PaeHfq appears to be a consequence of the
incorporation of water molecules between the side chain of the
threonine and the main chain of the adjacent protein monomer as
discussed above. In the H57N PaeHfq protein stereochemical
analysis showed that the asparagine residue is able to organize a
direct but water-accessible hydrogen bond to the main-chain atoms of
the neighbouring monomer. Therefore, this substitution resulted in a
decreased melting temperature for the mutant protein forms but did
not lead to deterioration of the melting cooperativity.
The research was supported by the Russian Academy of Sciences,
the
Russian
Federal
Agency
for
Science
and
Innovation
(02.740.11.0295), the Russian Foundation for Basic Research (10-04-
00818) and the Program of the RAS on Molecular and Cellular
Biology.
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structural communications
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Acta Cryst. (2010). F66, 760–764
|
3M4I
|
Crystal structure of the second part of the Mycobacterium tuberculosis DNA gyrase reaction core: the TOPRIM domain at 1.95 A resolution
|
Structural Insights into the Quinolone Resistance
Mechanism of Mycobacterium tuberculosis DNA Gyrase
Je´re´mie Piton1,2,3, Ste´phanie Petrella4, Marc Delarue1,2, Gwe´nae¨lle Andre´-Leroux2,5, Vincent Jarlier4,
Alexandra Aubry4, Claudine Mayer1,2,6*
1 Unite´ de Dynamique Structurale des Macromole´cules, De´partement de Biologie Structurale et Chimie, Institut Pasteur, Paris, France, 2 URA 2185, CNRS, Paris, France,
3 UPMC Univ Paris 06, Paris, France, 4 UPMC Univ Paris 06, EA1541, Bacte´riologie-Hygie`ne, Paris, France, 5 Unite´ de Biochimie Structurale, De´partement de Biologie
Structurale et Chimie, Institut Pasteur, Paris, France, 6 Universite´ Paris Diderot Paris 7, Paris, France
Abstract
Mycobacterium tuberculosis DNA gyrase, an indispensable nanomachine involved in the regulation of DNA topology, is the
only type II topoisomerase present in this organism and is hence the sole target for quinolone action, a crucial drug active
against multidrug-resistant tuberculosis. To understand at an atomic level the quinolone resistance mechanism, which
emerges in extensively drug resistant tuberculosis, we performed combined functional, biophysical and structural studies of
the two individual domains constituting the catalytic DNA gyrase reaction core, namely the Toprim and the breakage-
reunion domains. This allowed us to produce a model of the catalytic reaction core in complex with DNA and a quinolone
molecule, identifying original mechanistic properties of quinolone binding and clarifying the relationships between amino
acid mutations and resistance phenotype of M. tuberculosis DNA gyrase. These results are compatible with our previous
studies on quinolone resistance. Interestingly, the structure of the entire breakage-reunion domain revealed a new
interaction, in which the Quinolone-Binding Pocket (QBP) is blocked by the N-terminal helix of a symmetry-related molecule.
This interaction provides useful starting points for designing peptide based inhibitors that target DNA gyrase to prevent its
binding to DNA.
Citation: Piton J, Petrella S, Delarue M, Andre´-Leroux G, Jarlier V, et al. (2010) Structural Insights into the Quinolone Resistance Mechanism of Mycobacterium
tuberculosis DNA Gyrase. PLoS ONE 5(8): e12245. doi:10.1371/journal.pone.0012245
Editor: Hendrik W. van Veen, University of Cambridge, United Kingdom
Received April 22, 2010; Accepted July 21, 2010; Published August 18, 2010
Copyright: 2010 Piton et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: J.P. is funded by the ‘‘Ministere de l’enseignement superieur et de la recherche.’’ The funders had no role in study design, data collection and analysis,
decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: mayer@pasteur.fr
Introduction
Type II topoisomerases are essential and ubiquitous nucleic acid-
dependent nanomachines involved in the regulation of DNA
topology and especially in the regulation of DNA supercoiling [1].
Type II topoisomerases act by an ATP-dependant double-stranded
DNA break [1]. Except archaeal topoisomerase VI [2,3], they all
belong to a single protein superfamily, the type IIA topoisomerases,
sharing homologous sequences and overall structures [4]. However,
they have acquired distinct functions during evolution [1]. Bacterial
genomes usually encode two type IIA enzymes, DNA gyrase and
topoisomerase IV. DNA gyrase facilitates DNA unwinding at
replication forks and topoisomerase IV has a specialized function in
mediating the decatenation of interlocked daughter chromosomes
[5]. Mycobacterium tuberculosis, the aetiologic agent of tuberculosis, is
unusual in possessing only one type II topoisomerase, DNA gyrase
[6]. Consequently, the M. tuberculosis DNA gyrase exhibits a different
activity spectrum as compared to other DNA gyrases, namely it
supercoils DNA with an efficiency comparable to that of other DNA
gyrases but shows enhanced relaxation, DNA cleavage, and
decatenation activities [7].
DNA gyrase and topoisomerase IV consist of two subunits
(GyrA and GyrB in DNA gyrase, ParC and ParE in topoisomerase
IV), which form the catalytically active heterotetrameric complex
(i.e. A2B2 and C2E2, respectively). Subunit A consists of two
domains, the N-terminal breakage-reunion domain and a carboxy-
terminal domain, termed CTD. Subunit B consists of the ATPase
domain followed by the Toprim domain. The GyrB Toprim and
GyrA breakage-reunion domains come from separate subunits and
cooperatively form the enzyme core (Figure 1A). The breakage-
reunion domain contains the catalytic tyrosine responsible for the
cleavage and religation of the DNA double helix. Although the
structure of a fully intact, active type IIA topoisomerase has yet to
be determined, structural and biochemical studies of the individual
fragments have led several authors to propose a model of its global
quaternary structure and a catalytic mechanism of the holoenzyme
[8]. The breakage-reunion domain binds a DNA segment termed
the ‘gate’ or G-segment at the DNA-gate. The N-terminal ATPase
domains dimerize upon ATP binding, capturing the DNA duplex
to be transported (T-segment). The T-segment is then passed
through a transient break in the G-segment opened by the
breakage-reunion domains, the DNA is resealed and the T-
segment released through a protein gate, the C-gate, prior to
resetting of the enzyme to the open clamp form.
Quinolones, which target the two bacterial type II topoisom-
erases, exert their powerful antibacterial activity by interfering
with the enzymatic reaction cycle. Specifically, they bind to the
enzyme-DNA binary complex, thereby stabilizing the covalent
enzyme tyrosyl-DNA phosphate ester. The resulting ternary
complexes block DNA replication and lead to cell death [9].
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August 2010 | Volume 5 | Issue 8 | e12245
Quinolones are one of the most effective second-line drugs in the
treatment of multidrug-resistant tuberculosis (MDR-TB; strains
resistant to the two main antituberculous drugs, rifampicin and
isoniazid) [10] and are currently under study for shortening
treatment duration of drug-susceptible tuberculosis [11]. Tuber-
culosis still remains the leading cause of death from a curable
infectious disease causing millions of deaths annually (http://www.
who.int). Unfortunately, due to the long and complex nature of
TB treatment, inappropriate use of first line antituberculous drugs
is common, leading to the emergence of drug-resistant bacilli,
especially MDR strains. Widespread dissemination of these bacilli
poses a serious threat to global TB control [12]. Compared with E.
coli, the ‘‘intrinsic resistance’’ of M. tuberculosis to quinolones is
relatively high, mainly due to the primary structure of DNA
gyrase. Namely, amino acids at positions 81 and 90 in GyrA and
482 in GyrB have been demonstrated to be involved in ‘‘intrinsic
quinolone resistance’’ [13]. Nonetheless, quinolones, and in
particular fluoroquinolones, are essential antibiotics for MDR-
Figure 1. Domain organization and structures of the individual domains from the M. tuberculosis DNA gyrase catalytic core. A.
Domain organization of the M. tuberculosis DNA gyrase. The catalytic core is composed by the Toprim domain and the breakage-reunion domain. B.
Three orthogonal views of the dimeric Toprim domain from M. tuberculosis colored by regions. The crystal structure of the complete Toprim domain
(TopBK) encompasses residues T448 to E654. The schematically represented primary sequence is colored as in the structure. The N-terminal residue
numbers of the regions (Toprim, tail and hinge) and the TopBK C-terminal residue number are indicated. The Toprim region, constituted by
discontinuous N- and C-terminal sequence segments and containing the magnesium-binding site (E459, D532 and D534) and the QRDR-B (Quinolone
Resistance Determining Region in GyrB) is colored in yellow, the Tail region in purple and the hinge between the two regions in blue. The second
monomer generated by a crystallographic two-fold axis is represented in grey. C. Three views of the dimeric breakage-reunion domain from M.
tuberculosis colored by regions. The crystal structure of the complete breakage-reunion domain (GA57BK) extends from D9 to A501. The N-terminal
helix is colored in red, the DNA-gate containing the catalytic residues R128 and Y129 and the QRDR-A in blue, the ‘tower’ in green, the helix-bundle in
orange and the C-gate in purple.
doi:10.1371/journal.pone.0012245.g001
M. tuberculosis DNA Gyrase
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August 2010 | Volume 5 | Issue 8 | e12245
TB [13,14]. However, M. tuberculosis develops ‘‘acquired resis-
tance’’ to quinolones following prolonged exposure, leading to the
emergence of extensively drug-resistant (XDR) strains (MDR-TB
strains resistant to any fluoroquinolone and to at least one of three
injectable second-line anti-TB drugs) [15,16,17]. This ‘‘acquired
resistance’’ is mainly a result of mutations in the DNA gyrase
sequence [18,19]. Mutations conferring bacterial resistance to
quinolones occur in two short discrete segments termed the
quinolone resistance-determining regions (QRDR) [20] located in
the breakage-reunion domain of GyrA subunit (QRDR-A) and less
frequently in the Toprim domain of GyrB (QRDR-B) [20,21,22].
Among the described mutations, we have unequivocally demon-
strated that the nature of the amino acids at positions 88, 90 and
94 in GyrA plays a crucial role in the ‘‘acquired resistance’’ to
quinolones (Table 1) [21,23].
The challenge of better understanding the complex mechanism
of quinolone resistance in M. tuberculosis requires high-resolution
structures of the antibiotic targets. Following our previous results,
the aim of this work was to obtain a 3-dimensional understanding
of the relationships between a given amino acid mutation and
quinolone resistance phenotype in M. tuberculosis. Simultaneously
to our results, two structures of M. tuberculosis DNA gyrase domains
were published last year, the low resolution GyrB’ structure (PDB
code 2ZJT, [24]) and the truncated MtGyrA59 domain (PDB code
3ILW, [25]). The first picture of the enzyme-quinolone interac-
tions was given by the low resolution structures of Streptococcus
pneumoniae ParC breakage-reunion and ParE Toprim domain in
complex with DNA and quinolones (PDB codes 3FOF and 3K9F,
[26]). Moreover, other efforts to develop new potent catalytic
inhibitors of bacterial DNA gyrase were illustrated by the crystal
structure of E. coli DNA gyrase in complex with the bifunctional
antibiotic simocyclinone D8 [27]. Its mode of action is unique in
that it directly interacts with DNA gyrase to prevent its binding to
DNA.
In this work, we combined X-ray crystallographic studies,
sedimentation velocity experiments and activity assays of the two
domains that form the enzyme core of M. tuberculosis DNA gyrase,
the GyrB Toprim and GyrA breakage-reunion domains. We
solved two high resolution structures of the Toprim domain
displaying two different conformations of the metal-binding site, to
2.1 and 1.95 A˚ resolution, respectively. The crystal structure of the
breakage-reunion domain we solved to 2.7 A˚ resolution, revealed
a promising interaction that will be further exploited for drug
design. This interaction involves the N-terminal helix, which is
anchored in the active site of a symmetry-related molecule.
Additionally, using the crystal structures of both domains, we
modeled the catalytic reaction core in complex with DNA and a
quinolone. This study brings the first structural explanation on
quinolone resistance mechanism of M. tuberculosis DNA gyrase.
Results
Crystal structures of the Toprim and breakage-reunion
domains are biologically relevant
The C-terminal GyrB domain (Toprim domain, residues 448–
654) and the entire N-terminal GyrA domain (breakage-reunion
domain, known as GyrA59 in E. coli, residues 1–502), hereafter
named TopBK and GA57BK, respectively, were overproduced
and purified. DNA cleavage activity assays show that TopBK is
able to catalyze DNA breaks when associated to the full-length A
subunit. Similarly, GA57BK is able to catalyze DNA breaks when
associated with the full-length B subunit (Figure 2A and B).
Interestingly, the GA57BK-TopBK complex has DNA cleavage
activity, showing that these domains possess all determinants for
DNA cleavage and confirming that these two domains form the
catalytic reaction core of the M. tuberculosis DNA gyrase (Figure 2A
and B). In addition to DNA cleavage, some nicking is also
observed when TopBK is associated either with the full length
GyrA, or with GA57BK (Figure 2B). This could be the result of a
decrease in the complex stability when TopBK is used in the
activity assays.
TopBK was crystallized in presence of magnesium (crystal I) and
calcium (crystal II) and the structures were solved at 2.1 A˚ and
1.95 A˚
resolution, respectively,
with one
monomer in the
asymmetric unit in both cases. Slight modifications of the
previously described crystallization conditions [24,28], e.g. mod-
ifying the pH value and adding divalent cations, led to a space
group change and a substantial increase in the resolution (2.8 to
1.95 A˚ ). A crystallographic two-fold axis generates a dimeric
structure, similar to the dimer observed in the asymmetric unit of
the GyrB’ structure (2ZJT). GA57BK corresponds to the entire N-
terminal domain with a molecular mass of 57 kDa. The crystals
belong to space group C2, with a dimer in the asymmetric unit.
Clear electron density was observed for the N-terminal fragment
that could be built either entirely (chain A) or partially (chain B)
because of different crystal contacts. Consequently, the final model
spans residues 9 to 499 for chain A and 29 to 501 for chain B.
Both TopBK and GA57BK display a dimeric structure in the
crystal (Figure 1B and C). The biological relevance of these
dimeric forms was investigated using analytical ultracentrifugation.
Sedimentation experiments reveal that TopBK and GA57BK
exhibit different behaviour in solution. In the case of TopBK, two
species are observed with a 50/50 distribution when using a
protein concentration corresponding to the crystallization condi-
tions (Figure 2C). The two species display a sedimentation
coefficient of 2.360.1 S and 3.660.2 S, corresponding to the
monomer and the dimer, respectively, according to the theoretical
sedimentation coefficient values calculated from the crystallo-
graphic structure (2.2 and 3.5 S, respectively). In contrast,
GA57BK is mainly dimeric in solution (Figure 2C). Sedimentation
experiments showed that one species was observed with a
sedimentation coefficient of 5.460.2 S, compatible with the value
calculated from the crystallographic dimer structure (5.6 S). The
good agreement between these experimental and theoretical
values indicates that the dimeric conformation of GA57BK is
stable in solution. These results suggest that the biological unit is a
dimer.
Table 1. Mutations described in M. tuberculosis strains
implicated in ‘‘acquired’’ resistance to quinolones.
Mutation
Effect on quinolone
susceptibility
Reference
GyrA
GyrB
G88A
resistance
13
A90V
resistance
21
D94A, G, N
resistance
21
N499D
resistance
21
T80A
no effect
21
T80A+A90G
hypersusceptibility
21
Summary of mutations described in M. tuberculosis strains (e.g. clinical strains or
strains cultured in vitro in presence of quinolone in order to select a resistant
strain), which have been unequivocally demonstrated as implicated in
‘‘acquired’’ resistance.
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The crystal structure of the isolated Toprim domain is a
dimer
The overall fold of the TopBK structure is very similar in both
crystal forms, and also similar to the previously published GyrB’
structure (2ZJT) [24] and to the Toprim domain of the known
eukaryotic counterpart, the yeast topoisomerase II [29]. The
structure displays a two-domain organization, a globular domain
constituted by discontinuous segments (residues 448–564 and 633–
654) and the Tail domain (residues 565–608) connected by a loop-
helix-loop hinge region (residues 609–632) (Figure 1B). The
globular domain, organized in a Rossmann-like fold, contains the
Toprim domain described by Aravind and collaborators [30] and
the QRDR-B (residues 461–499) (Figure 3A and S1). The Tail
domain comprises a three-stranded antiparallel b-sheet and an a-
helix. In the globular domain, the conserved acidic triad (E459,
D532, D534), which constitutes the signature of the Toprim
domain, binds the magnesium ion essential for the catalysis of
the cleavage-ligation reaction. In the structure of crystal I, the
magnesium ion is not visible, despite being present in the
crystallization condition. However, side chains of the catalytic
triad are in conformations which would allow ion coordination, as
observed in the yeast topoisomerase II in complex with DNA
(Figure 3B). Presumably, the ion is not bound due to the absence
of the DNA. When magnesium is substituted in the crystallization
conditions by calcium (TopBK crystal II), side chains of the triad
are observed in an inactive conformation similar to the one
observed for the low resolution M. tuberculosis Toprim domain
structure [24].
The Toprim domain forms a dimer with a symmetry related
molecule in both crystal structures (crystal I and II), burying
1017 A˚ 2 at the protein-protein interface, indicative of a biologi-
cally relevant interaction. The two species observed in sedimen-
tation experiments with a 50/50 distribution are identified as the
monomeric TopBK domain and the crystallographic dimer
suggesting that this crystallographic dimer exists in solution
outside the context of the full-length subunit.
Surprisingly, the high resolution structures of TopBK, revealed
two disordered regions, between b1 and b2 (residues 460–474) and
between b2 and a2 (residues 484–492) (Figure 3A). These regions
are structured in the context of the catalytic core or in presence of
DNA. The first disordered region corresponds to the a1-helix [30],
as observed in the three structures of the yeast topoisomerase II
[29,31,32] and in the structure of the S. pneumoniae reaction core
[26]. Interestingly, this region is located at the dimer interface and
placing an a-helix would generate steric hindrance between the
two helices of the crystallographic related monomers (Figure 3C).
The second disordered region, the loop between b2 and a2, is
exposed to the solvent explaining its high flexibility. In the
structures of type II topoisomerases in complex with DNA, this
loop (hereafter named DBL for DNA-Binding Loop) constitutes
the interface between the Toprim domain and DNA and is
stabilized through protein-DNA interactions.
The breakage-reunion domain is in a closed
conformation
GA57BK forms a biological dimer in the asymmetric unit,
generating a heart-like shaped structure with outer dimensions of
1006100690 A˚ (Figure 1C) and a central hole of 30 A˚ diameter
allows the passage of the T-segment from the DNA-gate to the C-
gate. GA57BK forms a biological dimer in a ‘closed’ conformation
in the asymmetric unit, as the C-gate, which constitutes the so-
called primary dimer interface, and the DNA-gate, the secondary
protein-protein interface, are both closed (Figure S2). This closed
conformation is observed in all isolated breakage-reunion domain
structures, the MtGyrA59 from M. tuberculosis [25], GyrA59 from
E. coli and of the two topoisomerase IV structures from S.
pneumoniae [33] and from S. aureus [34]. This shows that the closed
conformation is stable and energetically favorable. This stability is
Figure 2. Activity assays and oligomerization of the TopBK and
GA57BK domains. A. The quinolone-mediated DNA cleavage activity
test measured on supercoiled pBR322 DNA (0.4 mg) as a substrate in the
presence of moxifloxacin (50 mg/ml) and 2.5 mg of each subunit alone:
full length subunit A (ABK), full length subunit B (BBK), GA57BK and
TopBK. Lanes a and b are supercoiled pBR322 DNA and control of
cleavage activity with WT M. tuberculosis DNA gyrase (ABK and BBK),
respectively. B. The quinolone-mediated DNA cleavage activity test
measured on supercoiled pBR322 DNA (0.4 mg) as a substrate in the
presence of moxifloxacin (50 mg/ml) with various amounts (indicated by
values in mg) of GA57BK associated with the full length subunit B (BBK,
1 mg), various amounts of TopBK with the full length subunit A (ABK,
1 mg), and various amounts (indicated by values in mg) of the binary
complex constituted by GA57BK and TopBK. Lanes M, a and b are DNA
size markers, supercoiled pBR322 DNA and control of cleavage activity
with WT M. tuberculosis DNA gyrase (ABK and BBK), respectively. N, L and
S denote nicked, linear and supercoiled DNA, respectively. C. Sedimen-
tation experiments of GA57BK and TopBK. The single peak of GA57BK
corresponds to the dimer, with a sedimentation coefficient of 5.460.2 S.
The two peaks observed for TopBK correspond to the monomeric and
dimeric form, with sedimentation coefficients of 2.360.1 S and 3.460.2 S,
respectively. c(s) on the y-axis designates the distribution of the
sedimentation coefficients observed for the experiment.
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essential to generate the interface needed to trap the DNA G-
fragment in order to start the topoisomerase cycle. This is in
agreement with FRET experiments showing that the DNA-gate of
the Bacillus subtilis DNA gyrase is predominantly in the closed
conformation during the DNA relaxation and supercoiling
reactions [35]. When comparing all five protein-protein interfaces
(sum of DNA- and C-gate interfaces), the highest value is observed
for both structures of M. tuberculosis DNA gyrase (Table S1).
Whereas the C-gate displays similar values, ranging from 1029 to
1120 A˚ 2, differences in interface area are observed at the DNA-
gate with a value of more than 800 A˚ 2 for M. tuberculosis,
representing nearly one half of the total interface. Both structures
confirm that the M. tuberculosis breakage-reunion domain has a
compact closed conformation, especially at the level of the DNA-
gate, whatever the crystal environment.
Each monomer of GA57BK contains five distinct regions, the
N-terminal fragment (residues 9–41), deleted in MtGyrA59 and
disordered in the homologous structures, and the four typically
observed regions in breakage-reunion domains of all type II
topoisomerases (Figure 1C and S3). In this way, GA57BK
resembles the type II topoisomerase structures in complex with
Toprim, namely the yeast topoisomerase II or the structure of the
complex between ParC, ParE, DNA and a fluoroquinolone (see
below). The next four domains, the DNA-gate (residues 42–169),
the ‘tower’ (residues 170–355 and 491–501), the C-gate (residues
401–444) and the three-helix bundle (residues 356–400 and 445–
490) (Figure 1C), exhibit an overall structural fold similar to that
observed for other bacterial type II topoisomerases [33,34,36] and
the yeast topoisomerase II [29,31,32]. The DNA-binding helix-
turn-helix motif (a3 and a4 helices), the QRDR-A (residues 74–
113) and the catalytic residues involved in DNA cleavage, namely
R128 and Y129, are localised in the DNA-gate.
The active site is blocked through crystal contacts
established by the N-terminal helix
In contrast to other structures of the breakage-reunion domain
alone (i.e. E. coli DNA gyrase, S. aureus and S. pneumoniae topoisomerase
IV), the N-terminal segment of GA57BK (residues 9–41) is ordered
and is organized in two distinct secondary structures (Figure 4A).
Residues D9 to E16 form a loop whose B factors indicate high
flexibility, followed by a 24-residue long a-helix (Figure 4B). Until
now, this helix was only observed when the Toprim domain is also
present, whether DNA is complexed (in the structure of the S.
pneumoniae topoisomerase IV catalytic core in complex with DNA,
3FOF [26] and the yeast topoisomerase II catalytic core-DNA
complex, 2RGR [29]) or not (in the two structures of the yeast
topoisomerase II catalytic core, 1BJT [32] and 1BGW [31]).
A previously unobserved feature of our crystal structure of
GA57BK is the interaction between this N-terminal region with
neighbouring molecules in the crystal packing. As shown in
Figure 4, the N-terminal fragment residues of chain A in a given
asymmetric unit clearly establish direct contacts with the active site
residues of its nearest neighbour (chain A’) in the adjacent
asymmetric unit. As these two molecules are related by the
crystallographic two-fold axis, this interaction is reciprocal. The a-
helix is deeply anchored in the active site of its neighbouring
molecule. Several hydrogen-bonding interactions link E23 from
the a-helix to the a3–a4 region, namely D89, A90 and S91 from
the symmetry-related molecule (Figure 4C). R26 links the main
chain carbonyl-group of H87 via a water molecule. In addition,
D30 establishes hydrogen bonds with the hydroxyl group of the
catalytic tyrosine (Y129) and a salt bridge with the catalytic
arginine (R128). Finally, on the opposite face of the N-terminal
helix, S27 and Y31 form an H-bonding network with R39 and
R54 from the symmetry-related molecule (Figure 4C). This
arrangement buries a surface area of 1227 A˚ 2, indicating a stable
interaction. The resulting tetramer could explain the small peak
observed in sedimentation experiments (Figure 2B). Further
studies exploiting this interaction
for drug design will be
investigated. A peptide of 16 amino acids corresponding to
residues 15 to 30 of the M. tuberculosis breakage-reunion domain
will be used as an inhibitor for M. tuberculosis DNA gyrase in
activity and binding assays in order to develop structure-activity
relationships. Combined docking and molecular dynamics simu-
Figure 3. The TopBK magnesium-binding site. A. Overall view of the dimeric structure of the Toprim domain from M. tuberculosis. One
monomer constituting the asymmetric unit is represented in green, the second monomer generated by a crystallographic two-fold axis in grey. The
secondary structures are indicated by black labels. The locations of the two disordered regions, the DNA Binding Loop (DBL) and the a1-helix, are
indicated by the red labels ‘‘DBL’’ and ‘‘a1’’, respectively. B. The magnesium-binding site of both M. tuberculosis TopBK structures, TopBK crystal I
(3IFZ, in green) and TopBK crystal II (3M4I, in purple) with the conserved residues, E459, D532 and D534. The active site of the S. cerevisiae Toprim
domain (2RGR) is represented in blue and its bound magnesium ion in orange. C. Close view of the TopBK dimer interface. The two symmetry-related
a1 helices (shown in red and grey) generate steric clashes.
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lations will be used to design small molecules that mimic the
peptide-active site interactions [37].
The M. tuberculosis breakage-reunion domain possesses
two specific structural motifs
Unexpectedly, structural comparison of M. tuberculosis DNA
gyrase to other type II topoisomerases clearly reveals that there is no
significant
difference between a
DNA gyrase
from
species
containing only one type II topoisomerase and the other type II
topoisomerases, DNA gyrase and topoisomerase IV, generally
found in bacteria (Figure S4). However, we found that two regions
could be correlated to the wider substrate spectrum of M. tuberculosis
DNA gyrase function. First, a sequence motif (DPP) in the loop
between the a3–a4 DNA-binding motif and the catalytic tyrosine
residue resembles the sequence observed in topoisomerases IV and
is rarely observed in DNA gyrase sequences. Localised at the side of
Figure 4. The active site of M. tuberculosis DNA gyrase is blocked by the N-terminal helix of a symmetry-related molecule. A. Two
dimers of GA57BK, related by the crystallographic two-fold axis, interact through the N-terminal helix. B. Omit maps for the N-terminal helix. The
(2Fobs – Fcalc) map shown in blue is contoured at 1.5 s whilst the (Fobs – Fcalc) map shown in green is contoured at 3 s. C. Detailed interactions of the
N-terminal helix (chain A’, in hot pink) in the active site of the symmetry-related molecule (chain A, in light green). Y31 of the N-terminal helix and R54
of the symmetry-related molecule are located on the back-side of the helix and are not represented for better clarity. D. Based on the model
discussed in the text, the N-terminal helix (chain A’, in hot pink) occupies the quinolone-binding pocket (QBP) and clashes with the modeled DNA,
represented in orange, and the fluoroquinolone, in yellow, bound to the QBP.
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the DNA-gate and in direct interaction with DNA (Figure 5A and
B), this loop could contribute to the topoisomerase IV-like activity
(i.e. decatenation) of M. tuberculosis DNA gyrase. Second, a specific
insertion in the M. tuberculosis sequence consists in a negatively
charged motif DEEE (residues 211–214) (Figure S3). In the
structure, this motif is localised at the solvent-exposed surface of
the tower domain in the a10-loop-a10’ region (Figure 5C). SAXS
studies showed that this region interacts with the GyrA CTD [38].
Superimposition of the different breakage-reunion domains shows
that the structures display significant differences in this region and
can be clustered in three distinct groups according to the
conformation of the loop (Figure 5C). First, the eukaryotic
topoisomerase II group, represented by the three different structures
of the S. cerevisiae topoisomerase II, is characterized by the absence of
the helices a10’. The second group, which contains the bacterial
type IIA topoisomerases (topoisomerase IV or DNA gyrase) from
organisms containing two topoisomerases, possess a short a10’. The
interaction between this region and the CTD might therefore be
different in these two groups suggesting that this region may be
implicated in functional specificity of type II topoisomerases, as the
CTD plays a crucial role in DNA interaction. Finally, the two
structures of M. tuberculosis constitute the third group. The DEEE
motif creates an extension of the a10’ helix modifying the CTD
interface and could thus play an important role during the catalytic
cycle of the M. tuberculosis DNA gyrase. To confirm the relationships
between these two specific structural motifs and the function of M.
tuberculosis DNA gyrase, the role of the DPP and the DEEE motifs
will be studied through site-directed mutagenesis.
Structural modeling of the catalytic reaction core in
complex with DNA and quinolone
During the catalytic cycle of DNA gyrase, a ternary complex is
formed between the Toprim and the breakage-reunion domains
and DNA. Quinolones target this complex and inhibit the enzyme
through stabilization of the covalent DNA-protein complex
formed during catalysis. To explore the mechanistic implications
of the M. tuberculosis DNA gyrase and to understand how the N-
terminal helix would interfere in the context of the complex
structure, we performed structural modeling of the cleavage
complex based on the structure of a topoisomerase IV complex
[26]. This quaternary complex is composed of the catalytic
reaction
core
consisting
of
the
breakage-reunion
domain
(GA57BK), the Toprim domain (TopBK), a 34-bp DNA duplex
and one of the most promising fourth-generation fluoroquinolone,
moxifloxacin. In the structure of the complex, DNA is settled on
the DNA gate, is linked covalently to the two catalytic tyrosines
129, and is maintained on each side by the ‘tower’ of the breakage-
reunion domain and the Toprim domain (Figure 6). The two
Figure 5. Comparison of the M. tuberculosis breakage-reunion domain to other type II topoisomerase structures. A. Global view of the
breakage-reunion domain. The boxes indicate the three close-up views shown in B, C and D. B. The DPP loop of GA57BK represented in light green is
near the DNA phosphate backbone, in orange (see text for details of the model). C. Close view of the a10–a10’ loop. Both M. tuberculosis structures,
GA57BK (represented in light green) and MtGyrA59 (in yellow) possess a DEEX sequence insertion in this loop. The conformation of this loop is
different in other bacterial type II topoisomerases, namely the three topoisomerase IV structures represented in red and E. coli GyrA59 in green, and in
the three yeast topoisomerase II structures in blue. D. Close-up view of the a3–a4 loop. The conformations of GA57BK chain B (light green), and
MtGyrA59 (yellow) are different from the conformation of GA57BK chain A (light green) and E. coli GyrA59 (dark green).
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catalytic sites related by the heterotetramer two-fold axis are
separated by four base-pairs and each catalytic site contains one
quinolone molecule (Figure 6A and B). The quinolone carboxylate
group points towards the major groove, and the R7 group is
localised in the minor groove. The interaction energy between
each quinolone molecule and its devoted binding pocket is 2105
and 2112 kcal/mol, respectively. The slight discrepancy could
reflect some sequential binding. However, those values evidence a
very good binding affinity that is illustrated in Figures 6B and C.
Discussion
In the present work, we have structurally characterized the two
components of the catalytic reaction core. The structure of the
breakage-reunion domain (known as GyrA59 in E. coli) reveals a
new interaction promising for drug design, whilst the high
resolution structures of the Toprim domain highlights two
disordered regions that play a crucial role during the catalytic
reaction of DNA gyrase. The strong point of this study is that we
could identify original mechanistic properties of quinolone binding
that clarify relationships between amino acid mutations and
resistance phenotype. These structure-mechanism relationships
have been established from the modeling of the catalytic reaction
core based on the two crystal structures, DNA and quinolone,
using the crystal structure of the cleavage complex formed by the
S. pneumoniae breakage-reunion and Toprim domains of topoisom-
erase IV stabilized by a fluoroquinolone [26].
The Quinolone-Binding Pocket (QBP), a drug-binding
pocket composed of protein and DNA residues
Whereas the structures of the S. pneumoniae topoisomerase IV
and the M. tuberculosis DNA gyrase reaction core are very similar,
our model allowed us to establish clear relationships between
Figure 6. Model of the catalytic reaction core in complex with DNA and moxifloxacin. A. Overall structure of the complex. GA57BK is
represented in blue, TopBK in red, the DNA in orange and the moxifloxacin in green. B. Close-up view of the two quinolone-binding pockets (QBP).
The purple arrow highlights the rise of the intercalated base step that constitutes the DNA walls of the QBP. Protein residues that constitute the QBP
protein walls are indicated in red for TopBk and blue for GA57BK. The residues shown in sticks belong to the QRDR and are implicated in quinolone
resistance. C. Close-up view along the DNA axis of one of the two QBP. The same residues as in B are represented in sticks. D. Schematic
representation of the interactions between QBP residues and chemical groups of the quinolone.
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amino acid mutation and resistance phenotype in M. tuberculosis
DNA gyrase. We propose that the atypical quinolone-binding
mode in the Quinolone-Binding Pocket (QBP), whose walls are
constituted not only by regions of the Toprim and the breakage-
reunion domains but also by DNA (Figure 6B), explains the effect
of the amino acid nature at a given position on the observed
resistance. The drug is intercalated between the dinucleotide step
for which the DNA backbone of one strand is broken (the
phosphorus atom is covalently linked to oxygen atom of the
catalytic tyrosine). The intercalated dinucleotide step is strongly
perturbed, with a twist of nearly 10u and a rise of 7.3 A˚ (36u and
3.4 A˚ for a canonical B-helix, 33u and 2.7 A˚ for A-DNA), typical
of an intercalation mechanism (as observed, for example, in the
structure of a DNA-nogalamycin complex [39]). The two
intercalated base pairs form a saddle, where quinolone is stabilised
through p-p interactions (Figure 6B and 7). The quinolone
molecule is blocked in this DNA saddle mainly by Van der Waals
contacts with residues of both protein domains (Figure 6C, D and
7). On one side, the carboxylate and the R2 groups (R2 is a
hydrogen atom in the moxifloxacin) of the drug are maintained by
the a3–a4 loop and the beginning of the a4-helix of the breakage-
reunion domain (residues 86–91). On the other side, quinolone is
immobilized by three regions of the Toprim domain. The b1-a1
loop (residues 459–462) interacts with the R1 group, the b2-DBL
loop (residues 480–486) with the R7–R8 group and the beginning
of a2 (residues 498–502) with the R7 group (Figure 6C and D).
Consequently, both deformation (rise) of the intercalated dinucle-
otide step forming the DNA saddle, and the specific sequence of
the QRDR-A and B, are required to build up the QBP and
determine the geometrical characteristics of the binding pocket
(volume and shape). In addition, the conformation of the loop
connecting helices a3 and a4 (residues 84–88) also affects the
depth of the QBP (Figure 6C and D). Whereas this loop displays
two different conformations in the two monomers in the GA57BK
crystal structure (Figure 5D), our model clearly shows that the
presence of DNA tends to push this loop towards the conformation
observed in the E. coli structure, suggesting that only this
conformation is observed when the QBP is formed.
Structural insights into the mechanism of ‘‘intrinsic
resistance’’ to quinolone
The three residues, M81 and A90 in GyrA and R482 in GyrB
have been shown to be implicated in ‘‘intrinsic’’ quinolone
resistance of M. tuberculosis [13,14]. We have previously demon-
strated that A90S and R482K substitutions (S and K are the
corresponding residues in E. coli) have direct effect on resistance
level [13]. In our model, A90 and R482 are part of the QBP,
residing on the a4-helix and in the b2-DBL, respectively. As
shown in Figure 6C, A90 side chain is oriented toward the
carboxylate group of the quinolone. The substitution of this
alanine for serine could increase the stability of the drug through a
hydrogen bond between the serine side chain and the hydroxyl
group of the quinolone, as observed in the S. pneumoniae complex
(3K9F). Furthermore, the R482 side chain is located in the minor
groove and forms a gate which blocks the quinolone in the pocket
(Figure 7). It has been shown that removing a lysine from the
minor groove energetically costs more than removing an arginine
[40]. The gate will open more easily when the residue at this
position is an arginine, in contrast to lysine, contributing to the
destabilisation of the quinolone in the QBP. This open-close
mechanism of the gate could play a role in the ‘‘intrinsic
resistance’’ mechanism. Finally, in our previous study, we showed
that M81I substitution (I in E. coli) alone had not any effect, but
could raise the quinolone susceptibility when associated with the
A90S mutation [13]. This correlates well with the fact that M81 is
not directly located in the QBP. This residue is spatially too far
from the quinolone-binding site, but it could affect the QBP by
altering
the
conformation
of
the
a4-helix
through
direct
Figure 7. Two views of the Quinolone-Binding Pocket (QBP). The DNA-protein complex is represented in molecular surface and moxifloxacine
in sticks. GA57BK is colored in dark blue, TopBK in firebrick, DNA in orange and moxifloxacin in green. The residues of TopBK belonging to the QBP
are colored in yellow for the b1-a1 loop residues, in purple for the b2-DBL residues (including R482), and in pink for the DBL-a2 residues. The residues
of GA57BK belonging to the QBP are represented in light green and correspond to the a3–a4 region.
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interactions of the residue at position 81 and two residues of the
helix, namely D89 and I92. All these observations show that direct
interaction has direct effect on the resistance level, and gives a
synergetic effect role to the amino acid nature at position 81. The
role of amino acids at position 81 and 90 in ‘‘intrinsic resistance’’
will be further investigated through structural studies of the M.
tuberculosis DNA gyrase double mutant A90S-M81I in complex
with DNA and a quinolone.
Structural insights into the mechanism of ‘‘acquired
resistance’’ to quinolone
A number of mutations that lead to fluoroquinolone ‘‘acquired
resistance’’ have been described in the literature [15,16]. They are
all localised in the QRDR-A and -B (residues 74–113 of the
GA57BK structure and 461–499 of the TopBK structure,
respectively). Interestingly, our model shows that all the residues
in the QRDR implicated in the ‘‘acquired resistance’’ are localised
in the QBP (as defined above), highlighting the relationships
between the QRDR of both subunits and the structurally
identified QBP, as previously suggested [41]. The model showed
that the overall geometry of the QBP, rather than the network of
H-bonding, is crucial for the recognition and binding of quinolone
in the pocket. Consequently, amino acid changes in the QBP will
lead to modification of the pocket geometry, either (i) directly, for
residues whose side chains point into the QBP or, most
importantly, (ii) indirectly, through modification of the DNA
structure, for residues interacting with the DNA moiety of the
QBP. Mutations implicated in nearly ninety percent of the
resistant strains are located in the QRDR-A at positions 90 and
94. Interestingly, only A90, which also contributes to the intrinsic
resistance of M. tuberculosis, interacts through a CH-O bond
between its methyl group with the quinolone carboxylate group.
Substitution by a valine could generate steric hindrance and this
could explain why this mutation is known to increase quinolone
resistance [42]. Mutations at other positions on the a4-helix affect
the DNA backbone structure by changing the major groove
dimensions, as DNA stacks on the a4-helix (Figure 6B and C).
Consequently, the size of the saddle formed by the intercalated
base pairs will be modified (Figure 7). This size modification could
affect the binding and the stability of the drug in the QBP. To
illustrate this mechanism, the amino acid at position 94 has a
paradoxical effect on the resistance level. Indeed, substitution by
either smaller residues like glycine or alanine and bulky residues
like tyrosine both increase the resistance level [21]. These residues
will either expand or reduce the volume of the pocket, leading to
instability of the quinolone in the QBP. Mutations in the QRDR-
B, like N499, are much less frequent, but their effects on DNA
gyrase activity can also be explained by this shape recognition
mechanism. All these observations can be used to improve the
efficacy of already existing quinolones.
Conclusion
Taken together with our previous work concerning the role of
specific residues implicated in quinolone resistance [13,23], our
structural results concerning the M. tuberculosis breakage-reunion
and Toprim domains and the modeled complex of the catalytic
reaction core provide key insights into the relationship between the
amino acid sequence of the M. tuberculosis DNA gyrase and the
resistance mechanism to quinolones, a major class of antibiotics
against this pathogen. In addition, these results highlight two
directions for future work. First, M. tuberculosis DNA gyrase, the
single type II topoisomerase in this organism, possesses two specific
structural motifs, the DEEE loop and the DPP loop, which could
partially explain its different activity spectrum as compared to
topoisomerase IV or DNA gyrase. Hence, this atypical activity
spectrum could be explained by the unique nature of the amino
acids present in the DNA gate. Second, the N-terminal helix of the
GA57BK structure is structurally ordered and stabilised through
crystal contacts. Interestingly, this helix blocks the active site of a
symmetry-related molecule through interactions with residues of
the a3–a4 loop. In the asymmetric unit, the dimeric structure
displays two different conformations for this loop. In agreement
with what was proposed by Tretter et al. [25], this suggests that this
region is conformationally dynamic (Figure 5D). Furthermore, this
helix contacts active site residues important for the catalysis of the
breakage-ligation reaction. The presence of this N-terminal helix
would prevent DNA binding (Figure 4D). These observations will
be exploited for the design of a new inhibitor family using peptide-
based approaches that target DNA gyrase by competitive
inhibition of DNA binding. Thus, they open up new avenues for
the development of novel peptide-based DNA gyrase inhibitors,
providing valuable new strategies to combat this disease as strains
resistant to the current repertoire of drugs are emerging.
Materials and Methods
Cloning, expression, purification and crystallization of
GA57BK
The breakage-reunion domain of DNA gyrase subunit A from
M. tuberculosis (residues 1–502), hereafter named GA57BK because
of its molecular weight of 57 kDa, was cloned, expressed and
purified as reported previously [43]. Briefly, the PCR amplified
construct was ligated into the pET-29a vector (Novagen) between
the NdeI and XhoI sites. The C-terminal His-tagged protein was
overproduced after transforming the plasmid into Rosetta 2(DE3)
pLysS (novagen), and purified with a Ni-NTA column and a size
exclusion
chromatography
using
Superdex-75
10/300
(GE
Healthcare). The protein was concentrated to 10–15 mg/ml in
100 mM Tris-HCl pH 8.
Ga57BK crystals were prepared using the hanging drop vapor
diffusion method, mixing 2 volumes of protein sample against 1
volume of reservoir solution [100 mM Sodium HEPES pH 7.5,
4% PEG 4000, 30% MPD]. Crystals grew after several days at
21uC to a maximum size of 2006200650 mm3.
Data collection, structure determination and refinement
of GA57BK
Crystals were directly flash frozen in liquid nitrogen. Native
diffraction data were collected at the SOLEIL PROXIMA-1
beamline to 2.7 A˚ resolution. The XDS package [44] was used for
all data integration and scaling. The crystals belong to space group
C2
with
unit
cell
dimensions
a = 163.9 A˚ ,
b = 109.6 A˚ ,
c = 102.0 A˚ , b = 120.4u and contain one biological dimer in the
asymmetric unit corresponding to a Matthews coefficient value of
3.4 A˚ 3/Da [45]. Data collection statistics are shown in Table 2.
The
structure
of
GA57BK
was
determined
by
molecular
replacement with AMoRe [46] implemented in CCP4 [47] using
the breakage-reunion domain of the DNA gyrase from E. coli [36]
(pdb accession code 1AB4) as a search model. Two distinct
orientations and positions were found in the asymmetric unit.
Structure refinement was carried out with BUSTER-TNT [48]
using two-fold non-crystallographic symmetry restraints. Model
building was performed manually with the program Coot [49].
Model refinement statistics are summarized in Table 2. The
figures were prepared using PyMol [50], available at http://pymol.
sourceforge.net/. Interface areas were calculated with the PISA
server [51].
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Cloning, expression, purification and crystallization of
TopBK crystal I and II
The
Toprim
domain
of
DNA
gyrase
subunit
B
from
Mycobacterium tuberculosis (residues 448–675), hereafter named
TopBK, was cloned into the expression vector pRSF-2 Ek/LIC
(Novagen). The plasmid was transformed into Rosetta 2(DE3)
pLysS (Novagen). The transformed cells were grown in LB
medium in presence of chloramphenicol and kanamycin. Gene
expression was induced by addition of IPTG (Sigma) to a final
concentration of 1 mM at 22uC over night. Cells were harvested
by centrifugation and stored at 220uC one night. Cells were
resuspended in buffer B1 containing 20 mM Tris-HCl pH 8,
500 mM NaCl and 15 mM imidazole. The cells were lysed by
sonication. Following the centrifugation, the protein was run over
a Ni-NTA column (GE Healthcare) equilibrated with buffer B1 at
4uC. The TopBK protein was eluted using a linear gradient from
15 to 500 mM imidazole. Finally, the protein was loaded on a
Superdex-75 10/300 (GE Healthcare) equilibrated with a buffer
containing 20 mM Tris-HCl pH 8. The protein was then
concentrated to 5 mg/ml in the same buffer.
TopBK crystal I was obtained in 10% PEG 4K, 200 mM
ammonium sulfate, 15 mM magnesium chloride, 100 mM Tris-
HCl pH 8 by vapour diffusion with the hanging drop vapour
diffusion method mixing 2 volumes of protein sample with 1
volume of reservoir solution [10% PEG 4K, 200 mM ammonium
sulfate, 15 mM magnesium chloride, 100 mM Tris-HCl pH 8].
TopBK crystal II was obtained in similar conditions, except that
magnesium chloride was substituted by calcium chloride.
Data collection, structure determination and refinement
of TopBK
For TopBK crystal I, diffraction data were collected at ESRF on
beamline id23eh1 to 2.1 A˚ resolution. The XDS package was used
for data processing and scaling (Table 2). The crystals belong to
Table 2. Data collection and refinement statistics.
TopBK crystal I
TopBK crystal II
GA57BK
Data Collection
Beamline
ESRF ID23eh1
SOLEIL
PROXIMA 1
SOLEIL
PROXIMA 1
Space group
P43212
P43212
C2
Unit cell dimensions
a, b, c (A˚)
52.9, 52.9, 190.2
52.8, 52.8, 190.5
163.9, 109.6, 102.0
a, b, c (u)
90, 90, 90
90, 90, 90
90, 120.4, 90
Wavelength (A˚)
0.9762
0.9800
0.9800
Resolution (A˚)
14–2.1 (2.3–2.1)a
29–1.95 (2.06–1.95)
35–2.7 (2.8–2.7)
Rsym (%)b
13.0 (55.0)
7.5 (58.7)
8.6 (72.6)
Redundancya
8.6 (4.0)
7.6 (7.8)
3.5 (3.5)
Completeness (%)a
99.1 (86.5)
99.7 (99.2)
98.9 (99.0)
I/sig(I)a
13.1 (3.3)
16.9 (3.4)
12.44 (2.24)
Refinement
Resolution (A˚)
14.0–2.1
17.0–1.95
19.9–2.7
No. Reflections
16487
20626
42396
No. Atoms
Protein
1474
1482
7534
Water
107
147
238
Rwork/Rfree
c
0.214, 0.249
0.210, 0.230
0.192, 0.233
B-factors
Protein
38.3
37.8
52.5
Water
52.6
52.2
57.6
RMSD
Bond length (A˚)
0.004
0.007
0.004
Bond angles (u)
0.835
0.97
0.719
Ramachandran analysis
Most favored (%)
93.8
92.7
90.2
Additional allowed (%)
5.6
6.7
9.3
Generously allowed (%)
0.0
0.0
0.4
Disallowed (%)
0.6
0.6
0.1
aThe values in parentheses are statistics from the highest resolution shell.
bRsym~P P DIhkl{Ihkl jð ÞD=P Ihkl where Ihkl(j) is the jth observed intensity of Ihkl and Ihkl is the final average value of intensity.
cRwork~P DDFobsD{DFcalcDD=P DFobsD and Rfree~P DDFobsD{DFcalcDD=P DFobsD where the sum is restricted to reflections that belong to a test set of 5% randomly selected
data.
doi:10.1371/journal.pone.0012245.t002
M. tuberculosis DNA Gyrase
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11
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space group P43212 with unit cell dimensions a = b = 52.87 A˚ ,
c = 190.22 A˚ . The structure of TopBK was determined by
molecular replacement with Molrep [52] implemented in ccp4
using one monomer of the previously published structure (PDB
accession code 2ZJT, [24]) as the starting model. The asymmetric
unit contains one monomer corresponding to a Matthews
coefficient value of 2.4 A˚ 3/Da. Structure refinement was carried
out with BUSTER-TNT to 2.1 A˚ resolution. Model building was
performed manually with the program coot. Model refinement
statistics are summarized in Table 2. For TopBK crystal II,
diffraction data were collected at SOLEIL PROXIMA 1 to 1.95 A˚
resolution. TopBK crystal II is isomorphous to crystal I and
structure determination protocol was the same as for crystal I
(Table 2).
Analytical ultracentrifugation
Sedimentation velocity experiments were performed in a
Beckman XL-I analytical ultracentrifuge using a double sector
charcoal-Epon cell at 20uC and 42000 rpm. Absorbance scans
were taken at 276 nm every 6 min. The protein concentration was
1 mg/ml for GA57BK corresponding to 17.5 mM in 20 mM Tris
pH 8. For TopBK, experiments were performed at three protein
concentrations, 0.5, 1 and 4 mg/ml (corresponding to 18, 37 and
148 mM, respectively) in the same buffer. The program Sednterp
1.09
(available
at
http://www.rasmb.bbri.org)
was
used
to
calculate
solvent
density
(0.9988 g/cm3),
solvent
viscosity
(0.010069 Poise) and partial specific volume (0.7340 ml/g for
GA57BK and 0.7390 for TopBK) using the amino-acid compo-
sition. The sedimentation data were analyzed with the program
Sedfit [53] using the continuous c(s) and c(M) distributions.
Theoretical sedimentation coefficients were calculated from the
crystal structure PDB file using Hydropro 7c [54] with a hydrated
radius of 3.4 A˚ for the atomic elements. The same experiments
were performed for GA57BK and TopBK in 20 mM Tris pH 8
and 100 mM NaCl. Sedimentation data were analyzed with
appropriate values of solvent density and viscosity.
Activity assays
DNA supercoiling and cleavage assays were carried out as
previously described [7,13,23,55]. Briefly, DNA cleavage assays
were performed with various ratios of purified M. tuberculosis GyrA
and GyrB subunits or GA57BK and TopBK domains. The
reaction mixture (total volume 20 ml) contained DNA gyrase assay
buffer (40 mM Tris-HCl pH 7.5, 25 mM KCl, 6 mM magnesium
acetate, 2 mM spermidine, 4 mM DTT, 0.1 mg/ml E. coli tRNA,
BSA (0.36 mg/ml), 100 mM potassium glutamate), supercoiled
pBR322 DNA (0.4 mg) as the substrate and moxifloxacin (50 mg/
ml). Proteins were added and reaction mixtures were incubated at
25uC for 1 h. Three ml of 2% SDS and 3 ml of a 1 mg/ml solution
of proteinase K were added, and incubation was continued for
30 min at 37uC. Reactions were terminated by the addition of
50% glycerol containing 0.25% bromophenol blue, and the total
reaction mixture was subjected to electrophoresis in 1% agarose
gel in TBE 0.56 buffer (Tris-Borate-EDTA, pH 8.3). After
running for 3.5 hrs at 50 V, the gel was stained with ethidium
bromide (0.7 mg/ml), photographed and quantified with an Alpha
Innotech digital camera and associated software. All enzyme
assays were done at least twice, with reproducible results.
Molecular modeling
The catalytic core model (GA57BK2+TopBK2+DNA) was
generated by superposition onto the crystal structure of the
Streptococcus pneumoniae topoisomerase IV catalytic core [26] (pdb
accession code 3FOF). Chains A and B from 3IFZ (GA57BK)
were superposed to the corresponding chains from 3FOF,
respectively, using SSM implemented in coot. The two disordered
regions of the TopBK structure were modeled using the Toprim
domain of 3K9F as a template. The amino acid torsion angles in
these regions were validated using the Ramachandran plot. The
two monomers of TOPBK were superposed using the same
method to the chains C and D of the S. pneumoniae topoisomerase
IV catalytic core structure. The DNA coordinates (chain E, F, G,
H) without moxifloxacin were inserted in the complex and defined
as fixed atoms. The complex was then energy minimized. Energy
minimization was performed with the NAMD2 program [56]
using CHARMM27 force field. The system was minimized by
300 000 steps of conjugate gradient minimization. Non bonded
interaction parameters were set such that electrostatic interaction
is shifted to zero at 12 A˚ and the van der Waals interaction is
switched off from 10 A˚ to 12 A˚ .
For the docking, the two fluoroquinolone moieties were
extracted from 3FOF coordinates. They were positioned in the
minimized catalytic core with respect to their respective positions
in the 3FOF structure. The system was further minimized using
the Minimization module of Discovery Studio (Accelrys), the
CHARMM forcefield and a cascade of Steepest Descent, Gradient
Conjugate and Adopted Basis Newton Raphson minimizations,
during which the backbone of the protein complex plus the DNA
atoms were constrained while the side chains and ligand moieties
were allowed to relax (6,000 iterations with final RMS gradient
0.01). We computed energetic criteria as the potential energy of
the complex. The minimised model deviates from the crystal
structure of the Streptococcus pneumoniae topoisomerase IV catalytic
core with an rmsd of 2.4 A˚ over 1033 Ca atoms. Finally, we
computed the interaction energy (which corresponds to the sum of
VDW and electrostatics non-bonded interactions) between each
moxifloxacin and its devoted quinolone-binding pocket with the
Calculate Interaction Energy module of Discovery Studio
(Accelrys).
Accession numbers
Co-ordinates and structure factors of TopBK crystal I have been
deposited in the protein data bank with the code 3IG0, TopBK
crystal II with the code 3M4I and GA57BK with the code 3IFZ.
Supporting Information
Table S1
Values of the interfaces calculated by PISA for the five
structures of the breakage-reunion domain dimer in closed
conformation. The PDB codes for the five structures are given:
3IFZ (this work) and 3ILW (25) correspond to M. tuberculosis DNA
gyrase, 1AB4 (36) to E. coli DNA gyrase, 2INR (34) to S. aureus
topoisomerase IV, 2NOV (33) to S. pneumoniae topoisomerase IV.
Nat, Nres correspond to the number of atoms and residues,
respectively, in interaction between the two monomers.
Found
at:
doi:10.1371/journal.pone.0012245.s001
(1.36
MB
DOC)
Figure S1
Structure-based sequence alignment of the Toprim
domain from type II topoisomerases. The sequence names are as
follows: MtGyr (PDB code 3IFZ) (this work), M. tuberculosis DNA
gyrase; SpTopIV (PDB code 3FOF) (26), S. pneumoniae topoisom-
erase IV and ScTopII (PDB code 2RGR) (29), S. cerevisiae
topoisomerase
II.
alpha-helices
(cylinders)
and
beta-strands
(arrows) of M. tuberculosis GA57BK are shown with the sequences
and color-coded according to Figure 1 (Toprim region in yellow,
the hinge in blue and the Tail region in purple). Residues
emphasized by black shading are 100% conserved. The magne-
sium binding site residues are underlined by red stars (E and DxD).
M. tuberculosis DNA Gyrase
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The disordered regions are emphasized in pale grey and indicated
as alpha1 and DBL for DNA Binding Loop. The QRDR-B is
delimited by a blue frame.
Found
at:
doi:10.1371/journal.pone.0012245.s002
(0.04
MB
DOC)
Figure S2
The three different conformations of the breakage-
reunion domain.
A. The
breakage-reunion
domain of M.
tuberculosis (PDB id 3IFZ) (this work), representing the closed
conformation with the DNA-gate and the C-gate closed. This
closed conformation is also observed in the E. coli DNA gyrase (36),
S. pneumoniae and S. aureus topoisomerase IV breakage-reunion
domain structures (33,34). B. The breakage-reunion domain of S.
cerevisiae in complex with DNA (PDB id 2RGR) (29), representing
an open conformation with the DNA-gate open and the C-gate
closed. C. The breakage-reunion domain of S. cerevisiae (PDB id
1BGW) (31), representing an open conformation with the DNA-
gate closed and the C-gate open.
Found
at:
doi:10.1371/journal.pone.0012245.s003
(0.90
MB
DOC)
Figure S3
Structure-based sequence alignment of the breakage-
reunion domain from type II topoisomerases. The sequence names
are as follows: MtGyr (PDB code 3IFZ) (this work), M. tuberculosis
DNA gyrase; EcGyr (PDB code 1AB4) (36), E. coli DNA gyrase;
SaTopIV (PDB code 2INR) (34), S. aureus topoisomerase IV;
SpTopIV (PDB code 2NOV) (33), S. pneumoniae topoisomerase IV;
EcTopIV (PDB code 1ZVU), E. coli topoisomerase IV and
ScTopII (PDB code 2RGR) (29), S. cerevisiae topoisomerase II.
alpha-helices (cylinders) and beta-strands (arrows) of M. tuberculosis
GA57BK
are
shown
with
the
sequences
and
color-coded
according to Figure 1 (N-terminal helix in red, DNA-gate in blue,
Tower in green, helix bundle in orange and C-gate in purple).
Residues emphasized by black shading are 100% conserved. The
catalytic residues are underlined by red stars (R128 and Y129) and
GA57BK specific motifs by black stars (the DPP and DEEX
motifs). The QRDR-A is delimited by a blue frame.
Found
at:
doi:10.1371/journal.pone.0012245.s004
(0.06
MB
DOC)
Figure S4
Superimposition of the different monomer structures
of the breakage-reunion domain. M. tuberculosis DNA gyrase
GA57BK (3IFZ) (this work) in light green, M. tuberculosis DNA
gyrase MtGyrA59 (3ILW, 25) in pale green, E. coli DNA gyrase
(1AB4) (36) in dark green, S. pneumoniae topoisomerase IV (2NOV)
(33) in red, S. aureus topoisomerase IV (2INR) (34) in pale red, S.
pneumoniae complexed with DNA (3FOF) (26) in dark red and E. coli
topoisomerase IV (1ZVU) in firebrick. The rmsd (in Ang.) after
superimposition and the number of common Ca (in parenthesis)
are indicated in the table. The color code is conserved.
Found
at:
doi:10.1371/journal.pone.0012245.s005
(0.43
MB
DOC)
Acknowledgments
We greatly acknowledge the help of Ahmed Haouz and the PF6 facility
(Plate-Forme de cristalloge´ne`se et diffraction des Rayons X), Bertrand
Raynal and the PFBMI facility (Plate-Forme de Biophysique des
Macromole´cules et de leurs Interactions) from the Pasteur Institute. We
thank Fre´de´ric Poitevin for help with modeling and Nathalie Barilone for
generous donation of M. tuberculosis H37Rv genomic DNA. We are
especially grateful to Olivier Poch for helpful discussions. We thank Joseph
Cockburn for careful reading of the manuscript. We want to dedicate this
manuscript to the memory of Warren DeLano, the developer of PyMol,
who passed away in November 2009.
Author Contributions
Conceived and designed the experiments: JP AA CM. Performed the
experiments: JP GAL AA. Analyzed the data: JP SP MD CM. Contributed
reagents/materials/analysis tools: VJ. Wrote the paper: JP AA CM.
Contributed to the writing of the paper: SP MD GAL.
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M. tuberculosis DNA Gyrase
PLoS ONE | www.plosone.org
14
August 2010 | Volume 5 | Issue 8 | e12245
|
3M4J
|
Crystal structure of N-acetyl-L-ornithine transcarbamylase complexed with PALAO
|
Reversible Post-Translational Carboxylation Modulates The
Enzymatic Activity Of N-Acetyl-L-Ornithine Transcarbamylase†
Yongdong Li1,2, Xiaolin Yu1, Jeremy Ho1, David Fushman3, Norma M. Allewell3, Mendel
Tuchman1, and Dashuang Shi1,‡
1Research Center for Genetic Medicine and Department of Integrative Systems Biology,
Children’s National Medical Center, The George Washington University, Washington, DC 20010,
USA.
2Key Laboratory of Organo-Pharmaceutical Chemistry, Jiangxi Province, Gannan Normal
University, Ganzhou 341000, China.
3Department of Chemistry and Biochemistry, College of Chemical and Life Sciences, University of
Maryland, College Park, MD 20742, USA.
Abstract
N-acetyl-L-ornithine transcarbamylase (AOTCase), rather than ornithine transcarbamylase
(OTCase), is the essential carbamylase enzyme in the arginine biosynthesis of several plant and
human pathogens. The specificity of this unique enzyme provides a potential target for controlling
the spread of these pathogens. Recently, several crystal structures of AOTCase from Xanthomonas
campestris (xc) have been determined. In these structures, an unexplained electron density at the
tip of Lys302 side-chain was observed. Using 13C NMR spectroscopy, we show herein that
Lys302 is post-translationally carboxylated. The structure of wild-type AOTCase complexed with
the bisubstrate analogue, Nδ-(phosphonoacetyl)-Nα-acetyl-L-ornithine (PALAO), indicates that the
carboxyl group on Lys302 forms a strong hydrogen bonding network with surrounding active site
residues, Lys252, Ser253, His293, and Glu92 from the adjacent subunit either directly or via a
water molecule. Furthermore, the carboxyl group is involved in binding N-acetyl-L-ornithine via a
water molecule. Activity assays with the wild-type enzyme and several mutants demonstrate that
the post translational modification of lysine 302 has an important role in catalysis.
Post-translational modification of the ε-amino group of lysine residues in proteins is a
common mechanism used by organisms to regulate protein functions including DNA-protein
interactions, subcellular localization, transcriptional activity, and protein stability and
activity (1). Lysine residues can be modified by the addition of functional groups to become
acetylated, methylated, carbamylated or carboxylated. The role of histone lysine acetylation
and methylation in affecting chromatin structure and gene expression has been well
established for more than a decade (2). However, the biological roles for lysine
carbamylation and carboxylation have rarely been investigated.
†This work was supported by Public Health Service grants DK-47870 (MT) and DK-067935 (DS) from the National Institute of
Diabetes, Digestive and Kidney Diseases. JH was supported by a Scholarship from the Doug and Lynn Parsons Family Foundation.
The Cornell High Energy Synchrotron Source (CHESS) is supported by the National Science Foundation under award DMR 0225180
and the Macromolecular Diffraction Facility at CHESS (MacCHESS) is supported by award RR-01646 from the National Institutes of
Health, through its National Center for Research Resources.
‡Corresponding author. dshi@cnmcresearch.org. Phone: 202-476-5817. Fax: 202-476-6014.
SUPPORTING INFORMATION AVAILABLE
Figure S1. Structure and hydrogen bonding network around residue 302 for previously determined AOTCase structures. This material
is available free of charge via the Internet at http://pubs.acs.org.
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 August 17.
Published in final edited form as:
Biochemistry. 2010 August 17; 49(32): 6887–6895. doi:10.1021/bi1007386.
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In vivo, lysine acetylation and methylation are usually carried out by acetyltransferase and
methyltransferase enzymes, respectively (3). In addition, some proteins such as hemoglobin
and human serum albumin can be acetylated non-enzymatically by chemicals such as
aspirin, methyl acetyl phosphate, and other acetylating agents such as acetyl-CoA (4–8).
Lysine can also be methylated by small chemicals in vitro, and this has routinely been used
as a rescue method for protein crystallization (9). Lysine carbamylation and lysine
carboxylation have only been achieved by using chemicals and no enzyme has yet been
found to catalyze these modifications. Lysine carbamylation was one of the earliest post-
translational modification of proteins to be elucidated when it was identified as a product of
reversible denaturation-renaturation studies of proteins with urea (10,11). This
carbamylation, which produces homocitrulline, has also been detected in uremic patients
(12) and in patients with elevated plasma and/or urinary lysine levels (13). In contrast, lysine
carboxylation is not as commonly reported, but has been identified in a number of proteins
via crystal structure determinations. In most of these proteins, the carboxyl groups of
modified lysines are involved in bridging two metal ions that play a structural role in the
active site. In several other proteins, however, a direct role for a carboxylated lysine in the
catalytic mechanism has been reported (14–16).
N-acetyl-L-ornithine transcarbamylase (AOTCase, EC 2.1.3.9) was recently discovered to
be part of a novel arginine biosynthesis pathway in plant pathogens of the
Xanthomonadaceae family such as Xylella and Xanthomonas (17–19). These pathogens
attack a variety of economically important crops including citrus fruits, cotton, tomatoes,
and rice (20,21). Genome sequence analyses showed that an AOTCase-like gene is also
present in some human pathogens such as Stenotrophomonas maltophilia and members of
the genus Bacteroides (22). In the case of Bacteroides fragilis, this gene was later confirmed
to encode another novel transcarbamylase, N-succinyl-L-ornithine transcarbamylase
(SOTCase, EC 2.1.3.11) (23). Crystal structures of both AOTCase and SOTCase bound with
substrate or substrate analogues have recently been determined (17,18,23). An extended
density at the side-chain tip of Lys302 in AOTCase was observed suggesting a post-
translational modification. Since Lys302 is located within the active site of AOTCase and is
not present in SOTCase, it was proposed as one of three key signature residues to
distinguish the two carbamylases (22). Here, we demonstrate that Lys302 is post-
translationally modified by carboxylation and that this change affects the catalytic function
of the enzyme.
MATERIALS AND METHODS
Materials
All chemicals were purchased from Sigma Chemical Company unless otherwise specified.
ANOR was purchased from Indofine Chemical Co., Inc. N-acetyl-L-citrulline was custom
synthesized and purified by Chiral Quest Company. PALAO (>95% purity) was synthesized
by IMI TAMI Institute of Research and Development Ltd. (19). xcAOTCase was prepared
and purified as previously described (18). Mutants K302A (primer: 5’-
CTGCGTCGCAACGTCGCGGCTACTGATGCGGTG-3’), K302E (primer:5’-
CTGCGTCGCAACGTCGAGGCTACTGATGCGGTG-3’) and K302R (primer: 5’-
CTGCGTCGCAACGTCAGGGCTACTGATGCGGTG-3’) were generated by site-directed
mutagenesis using the “Quik Change” mutagenesis kit (Stratagene) according to the
manufacturer’s protocol. The correct mutants were confirmed by DNA sequencing.
Recombinant mutant proteins were expressed and purified in the same manner as the wild-
type enzyme.
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Activity assay
The modified colorimetric assay method, which detects the formation of the ureido group
during the transcarbamylation reaction (24), was used to measure enzyme activity. CP and
AORN concentration were kept constant as 4.0 mM. After an incubation of 5 minutes, the
reaction was stopped by the addition of 1 ml of color reagent, as described previously (19).
A set of tubes containing known amounts of N-acetyl-L-citrulline was included with each
rack of enzyme assays to produce a standard curve for calculation of the enzyme specific
activity.
Mass spectrometric analysis
In order to identify the post-translational modification, mass spectrometric analysis was
carried out on a 4700 ABI TOF/TOF mass spectrometer (Applier Biosystems) based on the
method described previously (25). In brief, 10 µg of native protein were digested overnight
at 310 K using trypsin in 50 mM ammonium bicarbonate pH 7.4. After desalting using a
C18 ZipTip micropipette tip, the resulting peptides were eluted in 10 µl of acetonitrile/0.1%
TFA [70:30(v:v)]. The sample was mixed with matrix solution and spotted on a MALDI
plate to be submitted to the mass spectrometric analysis.
Chemical rescue experiments
The assay in the presence of various selected chemical was conducted as described above.
The stock solutions of small chemicals were titrated to the pH of the assay with KOH or
HCl.
13C NMR experiments
The wild-type and K302A mutant protein of AOTCase (~10 mg) was precipitated by
degassed buffer (pH 4.5) containing 25 mM sodium acetate. After centrifugation, the
precipitate was re-dissolved by adding a buffer containing 20 mM NaH13CO3, 100 mM Tris
HCl (pH 8.0) and 50 mM NaCl. Before NMR experiments, 40 µl D2O was added to 500 µl
protein sample. The 13C NMR spectra were collected on a Bruker Avance 600 spectrometer
(operating at 14.1 T) equipped with a direct 13C-detection probe at 298 K. The experimental
settings and processing parameters for the wild-type protein and K302A variant were
identical. 512 transients were collected with 4K time domain points and a spectral width of
3019 Hz centered at 160 ppm. The spectra were processed using exponential multiplication
with the line broadening factor set to 3Hz. The similarity of protein concentration in both
samples was verified by 1H NMR (not shown).
Crystallization, data collection and processing
PALAO-bound wild-type and mutant AOTCase crystals were grown using the hanging-drop
vapor diffusion method, with conditions similar to those used to produce native and ligand-
complexed AOTCase crystals (18,23). 2.0 µl of ~10 mg/ml solution of AOTCase were
mixed with 1.6 µl of reservoir solution and 0.4 µl PALAO solution (~0.01 M). The reservoir
solution contained 20% (w/v) PEG 3350, 0.2 M lithium sulfate and 0.1 M bis-Tris, pH 6.0.
Diffraction data for the PALAO-bound crystal were collected at 100 K at the F1 beam line
of the Cornell High Energy Synchrotron Source. Data sets for the PALAO-bound mutant
AOTCase crystals were obtained using a Rigaku anode x-ray generator in the Molecular
Structure Section of the National Institute of Health. All data were processed using
HKL2000 package (26) and reduced using the program TRUNCATE in the CCP4 suite (27).
Data collection parameters are listed in Table 1.
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Structure solution and refinement
Molecular replacement was used for phase determination of the PALAO-bound wild-type
and mutant AOTCase structures. The coordinates of AOTCase (PDB code: 3KZO) after
removal of ligands or water molecules were used for phase determination. Upon rigid-body
refinement, electron density corresponding to the ligands could be clearly visualized. The
ligands were built into the map using the graphics program O (28). Refinements were
carried out using molecular annealing, energy minimization and restrained B factor
refinement with the program CNS1.1 (29). During refinements, 5% of the reflections at
various resolutions were randomly selected to set aside to calculate Rfree to monitor the
progress of refinement (30). After every cycle of refinement, the model was manually
adjusted using the program O (28). Water molecules were automatically assigned using the
program WATERPICK of CNS. Model quality was checked using the program
PROCHECK (31) to ensure good stereochemistry for all three models. The final refinement
statistics are listed in Table 1.
Figures 1 was drawn using the programs Pymol (http://www.pymol.org). Figure 3 was
drawn using ChemDraw 8.0. The coordinates have been deposited with the RCSB PDB as
entries 3M4J, 3M4N, 3M5C and 3M5D.
RESULTS
Lys302 in AOTCase is carboxylated
To investigate the nature of the modification of Lys302 and how it affects catalytic activity,
we revisited all AOTCase structures. In the PALAO-bound AOTCase structure, the electron
density map clearly indicates that Lys302 is post-translationally modified (Figure 1A). The
type of modification can include methylation, acetylation, carbamylation, and carboxylation.
The shape of the electron density can been used to distinguish methyl groups from larger
functional groups, but it is difficult to distinguish between acetyl, carbamyl, and carboxyl
groups, all of which have three non-hydrogen atoms in a plane. Given the hydrogen bonding
network with surrounding residues (Lys252, Ser253 and His293, Table 2), a carboxylated
modification is the most likely choice for the modification of Lys302 in AOTCase. To
exclude that the modification’s identity represents chemically stable moieties (methyl,
acetyl, carbamyl), we analyzed trypsin digested fragments of purified AOTCase by TOF-
TOF mass spectroscopy. As expected, only a peptide fragment with an unmodified Lys302
was observed, consistent with the lability of the carboxylic group in acidic solutions. At low
pH, the carboxyl group is spontaneously released as carbon dioxide (14, 32), in contrast to
other modified groups that are stably bound and can be observed by mass spectrometry
analysis after proteolysis (33).
The putative carboxyl group on the modified Lys302 forms direct hydrogen bonds with
main-chain or side-chain nitrogen atoms of Lys252, Ser253 and His293 (Figure 1A and
Table 2). Among these, Lys252 is involved in direct hydrogen bonding to the carboxyl
group of the AORN moiety of PALAO, and His293 forms a strong hydrogen bond with the
main-chain nitrogen atom of Leu295 in the conserved His293-Cys294-Leu295-Pro206
(HCLP) motif. The hydrogen bonding network between the carboxyl group of modified
Lys302, His293 and the main-chain nitrogen atom of Leu295 is reminiscent of the similar
hydrogen bonding network, Glu310-His302-Leu304 and Glu299-Leu272-Leu274, found in
human and E. coli OTCase, respectively (34, 35). These three residues are conserved in all
OTCase sequences, and the interactions between them are important for maintaining the
HCLP motif in a specific conformation to orientate their main-chain oxygen atoms towards
the active site. In all known transcarbamylase structures, a leucine residue corresponding to
Leu295 is in an energetically unfavorable conformation and the peptide bond between this
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leucine and Pro296 is in the cis conformation. In addition to the direct hydrogen bonding
interaction above, the carboxyl group on the modified Lys302 interacts with the α-amino
nitrogen atom of the AORN moiety of PALAO and Glu92 from the adjacent subunit via
water molecules.
When we revisited all previously determined AOTCase structures (see supplementary
Figure S1) we found: (1) Lys302 was carboxylated in the absence of substrate binding, but
substrate binding immobilizes the side-chain of Lys302 further by hydrogen bonding
interaction via water molecules. (2) Water-mediated hydrogen bonding promotes
interactions of carboxylated Lys302 with Glu92 from the adjacent subunit and the α-amino
nitrogen atom of AORN. (3) Similarly to AOTCase, in the structure of SOTCase E92Z (Z =
Ala, Ser, Pro, Val), mutant with N-succinyl-L-norvaline bound (22) the carboxylated Lys302
hydrogen bonds to the α-amino nitrogen atom and the succinyl carboxyl group of N-
succinyl-L-norvaline via water molecules (Figure S1).
To obtain direct, independent evidence for the carboxylation of Lys302, 13C NMR
experiments were carried out with both wild-type protein and the K302A mutant. As
observed for other proteins with carboxylated lysine (36,37), the strong 13C NMR signal at
164 ppm characteristic of a carboxyl group was clearly detectable in AOTCase wild-type
protein labeled by 13C-bicarbonate, in contrast to the K302A mutant where the signal was
weak (Figure 2). Since there are 17 other lysine residues in the protein, the weak signal seen
for the K302A mutant might be caused by the adventitious carboxylation of another lysine
with reduced pKa, as has been observed for the K392A mutant of the sensor domain of the
BlaR protein (38).
Functional and structural studies of Lys302 mutants
To investigate the effect of lysine carboxylation on enzyme activity, Lys302 was mutated to
alanine, glutamate or arginine. Each of these variants was expressed in E. coli and gave
similar yields. Enzymatic assays demonstrated a significant decrease in enzymatic activity in
all three mutants, reflecting the functional importance of Lys302 (Table 3). The level of
enzymatic activity for the wild-type (WT) and three mutants was WT > K302A > K302E ≫
K302R. To determine the structural basis of these results, the WT and mutant enzymes
bound with PALAO were crystallized and their structures were determined at 1.8–2.2 Å
resolution. Only the K302R mutation had and appreciable effect on the structure of the
protein. Since K302 is located near the AORN binding site, the mutations would weaken
AORN binding to the active site.
In the structure of the K302A mutant, three additional water molecules (labeled as w3, w4
and w5 in Figure 1B) replace the carboxylated lysine. The two water molecules (labeled w1
and w2 in Figure 1A–1D) that mediate the hydrogen bonding interaction of carboxylated
Lys302 with PALAO and Glu92 from the adjacent subunit are also found in the K302A
mutant structure. Furthermore, these water molecules maintain a similar hydrogen-bonding
network to the wild-type enzyme. These results might explain why the K302A mutant
retains significant catalytic activity (Table 3). To investigate whether adding short-chain
carboxylic acids to the K302A mutant increases its activity as other enzymes (14, 15, 39,
40), the activity of the K302A mutant was measured in the presence of high formate and
acetate concentration (0.5 M). Surprisingly, the activity of the K302A mutant was not
significantly improved. The crystal structure of the K302A mutant soaking with the
crystallization buffer in the presence of 0.5 M acetate was also determined (not shown) and
it was observed that the same five water molecules were present in the cavity that replaced
the side chain of the carboxylated lysine. This, the acetate’s inability to replace the water
molecules in the crystal structure, is consistent with the unchanged activity assay results.
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The side-chain of Glu302 in the K302E mutant structure is well defined and anchored by
hydrogen bonding interaction with the main-chain nitrogen atom of Arg298 and weakly
hydrogen bonded to the main-chain nitrogen atom of Ser253 (Figure 1C). Two of three
additional water molecules (w4 and w5) observed in the K302A mutant structure occupied
the same position as the carboxyl oxygen atoms of Glu302 and form a similar hydrogen-
bonding network. Relative to the PALAO-bound wild-type structure, there is only one more
water molecule (w3) at the position of the carboxyl group of the carboxylated Lys302. This
water molecule mediates a hydrogen bonding interaction between Glu302 and Lys252. Two
common water molecules (w1 and w2) that interact with PALAO and Glu92 from the
adjacent subunit, respectively, were also identified in the K302E structure.
Our observation that the K302E mutant had lower enzymatic activity than that of the K302A
mutant (Table 3) was surprising since the carboxyl group of the glutamate could conceivably
function similarly to a carboxylated lysine. The explanation may be that, in the K302A
mutant, the hydrogen bonding network is well maintained by water molecules in the cavity
that replaces the carboxylated lysine. In particular, w3 is optimally located for strong
hydrogen bonding to w1 (2.7 Å), which in turn binds AORN. The distances between w1 and
the carboxyl oxygen of carboxylated Lys302 in all wild-type crystal structures are within
2.4–2.7 Å, but the distance between w1 and w2 in the K302E structure is significantly
greater (3.2 Å). The weaker hydrogen bonding interaction may be a reason for lower
enzymatic activity of the K302E mutant.
In contrast to the K302A and K302E structures, the K302R structure shows a much larger
reduction in enzyme activity relative to the wild-type enzyme. The electron density for the
side-chain of Arg302 is weak and the temperature factor of its side-chain is 54.4 Å2,
significantly higher than those of carboxylated Lys302 (44.7 Å2) and Glu302 (33.4 Å2),
implying greater flexibility. Furthermore, the side-chain of Arg302 is oriented differently
from the carboxyl group of carboxylated Lys302 and pushes the nearby residues His180,
Pro181 and Lys182 outwards about 1.0 Å (Figure 1D). However, the water molecules
involved in hydrogen bonding to the α-amino nitrogen atom of PALAO (w1) and the side-
chains of Lys252 (w3) and Glu92 (w2) from the adjacent subunit are conserved. Consistent
with the K302E structure, the distance between w1 and w2 is even greater (3.4 Å) than in
the WT structure and the hydrogen bonding interaction between w2 and w3 is no longer
observed. Thus, the almost undetectable enzymatic activity of the K302R mutant probably
results from the changes at its active site, including the weakened hydrogen bonding
network involved in substrate binding.
DISCUSSION
Several lines of evidence clearly indicate that Lys302 in AOTCase is carboxylated. First, the
extra electron density indicates that the side-chain of Lys302 is modified. Second, the
hydrogen bonding environment of Lys302 for hydrogen bonding interactions is compatible
with a carboxyl group, but not for a positively charged lysine side-chain. Third, the
modification is labile at low pH, since mass spectroscopy of samples prepared at low pH
indicated that Lys302 was no longer modified. Fourth, the clear presence of the
indicative 13C NMR signal at 164 ppm for wild-type protein and its absence in the K302A
mutant confirms carboxylation of Lys302.
It is well known that lysine carboxylation is non-enzymatic and reversible, while other post-
translational modifications such as methylation, acetylation, and carbamylation are
irreversible and detectable by mass spectroscopy. Furthermore, lysine methylation and
acetylation usually require an enzyme-catalyzed reaction in vivo (41). Therefore, it is
unlikely that such lysine modifications will be observed in recombinant proteins
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overexpressed in a foreign host (e.g. E. coli). Lysine methylation can be achieved by using
special chemicals in vitro, but these chemicals are not present in vivo. Lysine carbamylation
and carboxylation use completely different mechanisms to form functionally different
groups (Figure 3). Carbamylation can be achieved by cyanate produced from
myeloperoxidase-catalyzed oxidation of thiocyanate, an anion abundant in plasma and
increased in smokers, or from urea in the plasma. Lysine carboxylation, on the other hand,
occurs readily in aqueous solution in the presence of carbon dioxide at a basic pH (32,42).
Even though carbamylation and carboxylation use very different mechanisms, the two are
confused in the literatures. Lysine carbamylation (or carbamoylation) is referred to in
several publications (15,32,42–44), when the actual reaction is in fact carboxylation.
The activity of the K302A mutant is almost half of that of the wild-type enzyme raising the
question of why AOTCase retains a lysine in this position. Perhaps this lysine was
maintained through evolution to distinguish AOTCase from SOTCase which uses N-
succinyl-L-ornithine (SORN) rather than AORN (22), and OTCase which uses L-ornithine.
An alternative explanation may be found in the very low activity of the K302R mutant. The
side-chain of arginine has a positive charge while carboxylated lysine has a negative charge.
The side chain of unmodified lysine is usually located in a similar position as that of
arginine, as observed in the structure of UV damage endonuclease (14). It would be
expected that the activity of AOTCase with an uncarboxylated lysine would be as low as the
K302R mutant’s. It could further be surmised that, the respective organisms need to use
carboxylation as a switch to turn “on” or “off” the arginine biosynthetic pathway. It has been
well known that rubulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) in plant cells
uses the carboxylation on Lys201 as a switch to turn the enzyme “on” during the day and
“off” at night by removing the carboxyl group (45,46). Carbon dioxide and bicarbonate have
been found to play an important biological role in modulating several biological processes
including photosynthetic carbon fixation (47), pH homeostasis (48), carbon metabolism
(49), activation of virulence in pathogenic organisms (50), sperm maturation (51),
stimulation of mammalian G-protein-responsive adenylyl cyclase (52), and as an alarmone
in Drosophila (53,54). Whether or not carboxylation of a key lysine in their related proteins
is used as an underlying regulatory mechanism should be investigated further.
There are 197 structures with carboxylated lysine residue (modified residue indicated as
Kcx) in the Protein Data Bank (PDB). If structures with 90% identity are counted only once,
there are still 52 unique structures remaining in this pool (Table 4). These proteins include
hydantoinase (40,55), folylpolyglutamate synthase (43), UV damage endonuclease (14),
OXA10, OXA-1 class D β-lactamase (38,56,57), urease (42), phosphotriesterase (58),
dihydroorotase (59), dihydropyrimidinase (60), organophosphate hydrolase (61) and MurE
and MurD ligases (44,62). In most of these proteins, the carboxylated lysine bridges two
metal ions, similar to the role of glutamate or aspartate in proteins with two metal-binding
sites (26 structures among 52). However, the urease apoenzyme can be activated in vitro
only in the presence of carbon dioxide prior to nickel binding (63), suggesting that the
carboxylated lysine may have other structural roles beyond binding metals. In some proteins
such as β-lactamase, UV damage endonuclease, Rubisco, MurD and MurE ligase and BlaR
signal transducer protein, a carboxylated lysine plays an essential catalytic role. More
interestingly, in three structures (PDB ID: 1HL9, 1PU6 and 2UYN for fucosiadase, 3-
methyladenine DNA glycosylase and TdcF protein of unknown function, respectively), the
carboxylated lysines are located near the surface of proteins, presumably playing primarily a
structure stabilizing role (64–66). Since the carboxyl group is labile at acidic pH, but easily
formed in the presence of carbon dioxide at basic pH, the number of proteins with lysine
carboxylation must be underestimated. Furthermore, the carboxylated lysine must be fixed
in place by metal ions (either one or two) or hydrogen bonding with other protein residues
(at least one). Therefore, any detection method involving denaturing the proteins will result
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in release of the carboxyl group. With current technology, 13C NMR (38) and
crystallography are the only methods that can detect this modification. However, these
methods are not amenable to high-throughput investigations. The majority (49 out of 52
structures in the PDB) of known lysine carboxylation modifications were found to be
located at or near the active site, probably because these sites receive the most attention.
Revisiting the structures in PDB with more attention to surface lysines might reveal more
structures with carboxylated lysines.
In conclusion, we have shown that Lys302 in AOTCase is post-translationally modified by
carboxylation and that this modification may be functionally important for enzymatic
activity. Lysine carboxylation is likely to be a more common event than currently
appreciated and may play a critical role in enzymatic activity and protein stability.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Abbreviations
ACIT
N-acteyl-L-citrulline
ANOR
N-acetyl-L-norvaline
AORN
N-acetyl-L-Ornithine
AOTCase
N-acetyl-L-ornithine transcarbamylase
ATCase
aspartate transcarbamlyase
OTCase
ornithine transcarbamylase
CP
carbamyl phosphate
ORN
L-ornithine
PALAO
Nδ-(phosphonacetyl)-Nα-acetyl-L-ornithine
SORN
N-succinyl-L-ornithine
WT
wild-type
xc
Xanthomonas campestris
Acknowledgments
We thank Dr. David Davies for facilitating our use of the diffraction equipment in the Molecular Structure Section
of the National Institute of Health and Dr. Fred Dyda for help in data collection and processing, and Dr. Yui-Fai
Lam in the University of Maryland for help in setting up NMR measurements.
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Figure 1.
Stereo view of the structure and hydrogen bonding network surrounding residue 302. A,
PALAO bound wild-type AOTCase, B, PALAO bound K302A AOTCase C, PALAO bound
K302E AOTCase, D, PALAO bound K302R AOTCase. Contours of the electron density
maps (2Fo-Fc) around PALAO, residue 302 and water molecules are shown as a brown cage
at 1.0σ. The final refined positions of the ligands and surrounding protein residues are
represented as colored sticks. The predicted hydrogen bonding interactions are in pink
dashed lines. The water molecules are represented as pink balls. The carbon of PALAO,
residue 302 and other protein residues are shown in pink, light blue and green sticks,
respectively.
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Figure 2.
13C NMR spectra of wild-type (upper panel) and K302A mutant (lower panel) AOTCase (1
mM). Experiments were performed in 100 mM Tris HCl, 50 mM NaCl, 7% D2O, pH 8.0,
supplemented with 20 mM NaH13CO3. The position of the resonance attributed to
carboxylated lysine in the enzyme is around 164 ppm.
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Figure 3.
Chemical structure of carbamylated vs. carboxylated lysine.
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Table 1
Data collection and refinement statistics
Dataset
PALAO
K302A
K302E
K302R
Space group
I213
I213
I213
I213
Resolution (Å)
2.2
1.9
1.85
2.2
Unit-cell parameters (Å)
a = b = c =128.88
a = b = c =128.92
a = b = c =129.29
a = b = c =127.39
Measurements
219,475
305,757
390,128
246,817
Unique reflections
18,269 (1,832) a
28,236 (1,365)
30,622 (1,456)
17,635 (879)
Redundancy
12.0 (11.8)
10.8(5.4)
12.8 (5.4)
14.0 (13.1)
Completeness (%)
99.8 (100.0)
100.0 (100.0)
99.7 (95.1)
100.0 (100.0)
<I/σ (I)>
15.0 (4.9)
16.4 (2.3)
19.8 (2.8)
8.7 (3.7)
Rmerg b
7.4 (48.4)
6.5(64.9)
5.2 (55.3)
9.8 (79.1)
Wilson B (Å2)
30.4
27.6
28.6
21.9
Refinement
Resolution range (Å)
50.0-2.2
50-1.9
50-1.85
50-2.2
No. of protein atoms
2620
2613
2617
2619
No. of water atoms
90
219
193
146
No. of hetero atoms
24
24
24
24
Rmsd of bond lengths (Å)
0.006
0.005
0.005
0.005
Rmsd of bond angle (°)
1.1
1.2
1.2
1.2
Rwork (%)c
20.0
19.8
20.0
18.9
Rfree (%)d
24.3
23.2
23.2
22.2
Average B factor (Å2)
41.7
32.2
32.3
35.3
aFigures in brackets apply to the highest-resolution shell.
bRmerg = ΣhΣi|I(h,i)-<I(h)>|/∑hΣiI(h,i), where I(h,i) is the intensity of the ith observation of reflection h, and < I(h)> is the average intensity of
redundant measurements of reflection h.
cRwork= Σh‖Fobs| – |Fcalc‖/Σh|Fobs|.
dRfree = Σh‖Fobs| – |Fcalc‖/Σh|Fobs| for 5% of the reserved reflections.
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Table 2
Interactions between carboxylated lysine and other residues at the active site of AOTCase
Kcx302
Other residues
Bound ligands
PALAO
CPa
AORNb
CP +ANORc
SO4+ACITd
OQ1
K252 NZ
2.6
2.6
2.7
2.6
2.6
OQ1
W1e
2.6
2.6
2.6
2.7
OQ2
S253 N
3.0
3.1
2.8
2.9
2.9
OQ2
H293 NE2
3.0
3.2
3.0
2.9
2.9
NZ
W2f
3.1
2.9
3.0
3.0
aThe values were calculated based on PDB ID 3KZM.
bThe values were calculated based on PDB ID 3KZN.
cThe values were calculated based on PDB ID 3KZO.
dThe values were calculated based on PDB ID 3KZK.
eThis water molecule hydrogen bonds to N1 atom of PALAO, AORN or ANOR, and backbone O atom of Pro296 as well.
fThis water molecule hydrogen bonds to OE1 atom of Glu92 from adjacent subunit as well.
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Table 3
Specific activity of wild-type and mutant AOTCase in the presence of acids (0.5M).
Compounds added
Specific activity(µmol/min/mg)
Wild-type
K302A
K302E
K302R
None
43.4 ± 0.4a
23.0 ± 0.5
7.1 ± 0.1
0.059±0.01
Formate
44.1 ± 1.2
26.4 ± 0.6
6.7 ± 0.2
0.093±0.01
Acetate
48.5 ± 1.1
21.2 ± 0.8
6.6 ± 0.5
0.104±0.03
aThe Mean ± S.D. are shown (n = 3).
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Table 4
Protein structures with lysine carboxylation modification
PDB ID
Enzyme name
Residue
Organism source
Funciton
2OGJ
Dihydroorotase
175
A.tumefaciens
Bridging two Zn(II)
2Z26
Dihydroorotase
102
E.coli
Bridging two Zn(II)
3JZE
Dihydroorotase
103
S.enterica
Bridging two Zn(II)
2GWN
Dihydroorotase
149
P. gingivalis
Bridging two Zn(II)
3F4C
Organophosphorus hydrolase
243
G. stearothermophilus
Bridging two Co(II)
3ICJ
Metal-dependent hydrolase
294
P. furiosus
Bridging two Zn(II)
3GTX
Organophosphorus hydrolase
243
D. radiodurans
Bridging two Co(II)
2QPX
Metal-dependent hydrolase
166
L. casei
Bridging two Zn(II)
2FTW
Dihydropyrimidinase
158
D. discoideum
Bridging two Zn(II)
2FVK
Dihydropyrimidinase
167
S. kluyveri
Bridging two Zn(II)
3DC8
Dihydropyrimidinase
147
S. meliloti
Bridging two Zn(II)
3GNH
L-Lys/Arg carboxypeptidase
211
C. crescentus cb15
Bridging two Zn(II)
3DUG
Arginine carboxypeptidase
182
Unidentified
Bridging two Zn(II)
2VC7
Phosphotriesterase
137
S. solfataricus
Bridging two Co(II)
2R1N
Metallophosphotriesterases
169
A. tumefaciens
Bridging two Co(II)
2OB3
Phosphotriesterase
169
B. diminuta
Bridging two Zn(II)
3E74
Allantoinase
146
E. coli
Bridging two Fe(III)
1EJX
Urease
217
K. aerogenes
Bridging two Ni(II)
1E9Z
Urease
219
H. pylori
Bridging two Ni(II)
4UBP
Urease
220
B. pasteurii
Bridging two Ni(II)
1ONW
Isoaspartyl dipeptidase
162
E. coli
Bridging two Zn(II)
1K1D
D-hydanroinase
150
G. stearothermophilus
Bridging two Zn(II)
1GKR
L-hydanroinase
147
A. aurescens
Bridging two Zn(II)
1GKP
D-hydanroinase
150
Thermus sp.
Bridging two Zn(II)
1NFG
D-hydantoinase
148
R. pickettii
Bridging two Zn(II)
2ICS
Adenine deaminase
154
E. faecalis
Bridging two Zn(II)
1RQB
Transcarboxylase
184
P. freudenreichii
Binding one Co(II)
2QF7
Pyruvate carboxylase
718
R. etli
Binding one Zn(II)
3BG3
Pyruvate carboxylase
741
H. sapiens
Binding one Mn(II)
2OEM
Rubisco-like protein
173
G. kaustophilus
Binding one Mg(II)
1WDD
Rubisco
201
O. sativa
Binding one Mg(II)
1GK8
Rubisco
201
C. reinhardtii
Binding one Mg(II)
1BWV
Rubisco
201
G. partita
Binding one Mg(II)
2WTZ
ATP-dependent MurE ligase
262
M. tuberculosis
Binding one Mg(II)
2JFG
MurD ligase
198
E. coli
Catalytic role?
1E8C
MurE ligase
224
E. coli
Catalytic role?
1JBW
Folypolyglutamate synthetase
185
L. casei
Catalytic role?
1W78
FolC bifunctional protein
188
E. coli
Binding one Mg(II)
3HBR
OXA-48 β-lactamase
73
K. pneumoniae
Catalytic role
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PDB ID
Enzyme name
Residue
Organism source
Funciton
3ISG
Class D β-lactamase
70
E. coli
Catalytic role
2P9V
AmpC beta-lactamase
315
E. coli
Catalytic role
1K55
OXA-10 β-lactamase
70
P. aeruginosa
Catalytic role
1K38
β-lactamase OXA-2
70
S. typhimurium
Catalytic role
1XQL
Alanine racemase
129
G. stearothermophilus
Binding substrate?
1VFS
Alanine racemase
129
S. lavendulae
Binding substrate?
1RCQ
Alanine racemase
122
P. aeruginosa
Binding substrate?
2J6V
UV damage endonuclease
229
T. thermophilus
Catalytic role
1H01
Cell division protein kinase 2
33
H. sapiens
Catalytic role?
2UYN
Protein TdcF
A58
E. coli
Structural role?
1HL9
Fucosidase
338
T. maritime
Structural role?
1PU6
3-methyladenine DNA glycosylase
205
H. pylori
Structural role?
Biochemistry. Author manuscript; available in PMC 2011 August 17.
|
3M4N
|
Crystal structure of N-acetyl-L-ornithine transcarbamylase K302A mutant complexed with PALAO
|
Reversible Post-Translational Carboxylation Modulates The
Enzymatic Activity Of N-Acetyl-L-Ornithine Transcarbamylase†
Yongdong Li1,2, Xiaolin Yu1, Jeremy Ho1, David Fushman3, Norma M. Allewell3, Mendel
Tuchman1, and Dashuang Shi1,‡
1Research Center for Genetic Medicine and Department of Integrative Systems Biology,
Children’s National Medical Center, The George Washington University, Washington, DC 20010,
USA.
2Key Laboratory of Organo-Pharmaceutical Chemistry, Jiangxi Province, Gannan Normal
University, Ganzhou 341000, China.
3Department of Chemistry and Biochemistry, College of Chemical and Life Sciences, University of
Maryland, College Park, MD 20742, USA.
Abstract
N-acetyl-L-ornithine transcarbamylase (AOTCase), rather than ornithine transcarbamylase
(OTCase), is the essential carbamylase enzyme in the arginine biosynthesis of several plant and
human pathogens. The specificity of this unique enzyme provides a potential target for controlling
the spread of these pathogens. Recently, several crystal structures of AOTCase from Xanthomonas
campestris (xc) have been determined. In these structures, an unexplained electron density at the
tip of Lys302 side-chain was observed. Using 13C NMR spectroscopy, we show herein that
Lys302 is post-translationally carboxylated. The structure of wild-type AOTCase complexed with
the bisubstrate analogue, Nδ-(phosphonoacetyl)-Nα-acetyl-L-ornithine (PALAO), indicates that the
carboxyl group on Lys302 forms a strong hydrogen bonding network with surrounding active site
residues, Lys252, Ser253, His293, and Glu92 from the adjacent subunit either directly or via a
water molecule. Furthermore, the carboxyl group is involved in binding N-acetyl-L-ornithine via a
water molecule. Activity assays with the wild-type enzyme and several mutants demonstrate that
the post translational modification of lysine 302 has an important role in catalysis.
Post-translational modification of the ε-amino group of lysine residues in proteins is a
common mechanism used by organisms to regulate protein functions including DNA-protein
interactions, subcellular localization, transcriptional activity, and protein stability and
activity (1). Lysine residues can be modified by the addition of functional groups to become
acetylated, methylated, carbamylated or carboxylated. The role of histone lysine acetylation
and methylation in affecting chromatin structure and gene expression has been well
established for more than a decade (2). However, the biological roles for lysine
carbamylation and carboxylation have rarely been investigated.
†This work was supported by Public Health Service grants DK-47870 (MT) and DK-067935 (DS) from the National Institute of
Diabetes, Digestive and Kidney Diseases. JH was supported by a Scholarship from the Doug and Lynn Parsons Family Foundation.
The Cornell High Energy Synchrotron Source (CHESS) is supported by the National Science Foundation under award DMR 0225180
and the Macromolecular Diffraction Facility at CHESS (MacCHESS) is supported by award RR-01646 from the National Institutes of
Health, through its National Center for Research Resources.
‡Corresponding author. dshi@cnmcresearch.org. Phone: 202-476-5817. Fax: 202-476-6014.
SUPPORTING INFORMATION AVAILABLE
Figure S1. Structure and hydrogen bonding network around residue 302 for previously determined AOTCase structures. This material
is available free of charge via the Internet at http://pubs.acs.org.
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 August 17.
Published in final edited form as:
Biochemistry. 2010 August 17; 49(32): 6887–6895. doi:10.1021/bi1007386.
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In vivo, lysine acetylation and methylation are usually carried out by acetyltransferase and
methyltransferase enzymes, respectively (3). In addition, some proteins such as hemoglobin
and human serum albumin can be acetylated non-enzymatically by chemicals such as
aspirin, methyl acetyl phosphate, and other acetylating agents such as acetyl-CoA (4–8).
Lysine can also be methylated by small chemicals in vitro, and this has routinely been used
as a rescue method for protein crystallization (9). Lysine carbamylation and lysine
carboxylation have only been achieved by using chemicals and no enzyme has yet been
found to catalyze these modifications. Lysine carbamylation was one of the earliest post-
translational modification of proteins to be elucidated when it was identified as a product of
reversible denaturation-renaturation studies of proteins with urea (10,11). This
carbamylation, which produces homocitrulline, has also been detected in uremic patients
(12) and in patients with elevated plasma and/or urinary lysine levels (13). In contrast, lysine
carboxylation is not as commonly reported, but has been identified in a number of proteins
via crystal structure determinations. In most of these proteins, the carboxyl groups of
modified lysines are involved in bridging two metal ions that play a structural role in the
active site. In several other proteins, however, a direct role for a carboxylated lysine in the
catalytic mechanism has been reported (14–16).
N-acetyl-L-ornithine transcarbamylase (AOTCase, EC 2.1.3.9) was recently discovered to
be part of a novel arginine biosynthesis pathway in plant pathogens of the
Xanthomonadaceae family such as Xylella and Xanthomonas (17–19). These pathogens
attack a variety of economically important crops including citrus fruits, cotton, tomatoes,
and rice (20,21). Genome sequence analyses showed that an AOTCase-like gene is also
present in some human pathogens such as Stenotrophomonas maltophilia and members of
the genus Bacteroides (22). In the case of Bacteroides fragilis, this gene was later confirmed
to encode another novel transcarbamylase, N-succinyl-L-ornithine transcarbamylase
(SOTCase, EC 2.1.3.11) (23). Crystal structures of both AOTCase and SOTCase bound with
substrate or substrate analogues have recently been determined (17,18,23). An extended
density at the side-chain tip of Lys302 in AOTCase was observed suggesting a post-
translational modification. Since Lys302 is located within the active site of AOTCase and is
not present in SOTCase, it was proposed as one of three key signature residues to
distinguish the two carbamylases (22). Here, we demonstrate that Lys302 is post-
translationally modified by carboxylation and that this change affects the catalytic function
of the enzyme.
MATERIALS AND METHODS
Materials
All chemicals were purchased from Sigma Chemical Company unless otherwise specified.
ANOR was purchased from Indofine Chemical Co., Inc. N-acetyl-L-citrulline was custom
synthesized and purified by Chiral Quest Company. PALAO (>95% purity) was synthesized
by IMI TAMI Institute of Research and Development Ltd. (19). xcAOTCase was prepared
and purified as previously described (18). Mutants K302A (primer: 5’-
CTGCGTCGCAACGTCGCGGCTACTGATGCGGTG-3’), K302E (primer:5’-
CTGCGTCGCAACGTCGAGGCTACTGATGCGGTG-3’) and K302R (primer: 5’-
CTGCGTCGCAACGTCAGGGCTACTGATGCGGTG-3’) were generated by site-directed
mutagenesis using the “Quik Change” mutagenesis kit (Stratagene) according to the
manufacturer’s protocol. The correct mutants were confirmed by DNA sequencing.
Recombinant mutant proteins were expressed and purified in the same manner as the wild-
type enzyme.
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Activity assay
The modified colorimetric assay method, which detects the formation of the ureido group
during the transcarbamylation reaction (24), was used to measure enzyme activity. CP and
AORN concentration were kept constant as 4.0 mM. After an incubation of 5 minutes, the
reaction was stopped by the addition of 1 ml of color reagent, as described previously (19).
A set of tubes containing known amounts of N-acetyl-L-citrulline was included with each
rack of enzyme assays to produce a standard curve for calculation of the enzyme specific
activity.
Mass spectrometric analysis
In order to identify the post-translational modification, mass spectrometric analysis was
carried out on a 4700 ABI TOF/TOF mass spectrometer (Applier Biosystems) based on the
method described previously (25). In brief, 10 µg of native protein were digested overnight
at 310 K using trypsin in 50 mM ammonium bicarbonate pH 7.4. After desalting using a
C18 ZipTip micropipette tip, the resulting peptides were eluted in 10 µl of acetonitrile/0.1%
TFA [70:30(v:v)]. The sample was mixed with matrix solution and spotted on a MALDI
plate to be submitted to the mass spectrometric analysis.
Chemical rescue experiments
The assay in the presence of various selected chemical was conducted as described above.
The stock solutions of small chemicals were titrated to the pH of the assay with KOH or
HCl.
13C NMR experiments
The wild-type and K302A mutant protein of AOTCase (~10 mg) was precipitated by
degassed buffer (pH 4.5) containing 25 mM sodium acetate. After centrifugation, the
precipitate was re-dissolved by adding a buffer containing 20 mM NaH13CO3, 100 mM Tris
HCl (pH 8.0) and 50 mM NaCl. Before NMR experiments, 40 µl D2O was added to 500 µl
protein sample. The 13C NMR spectra were collected on a Bruker Avance 600 spectrometer
(operating at 14.1 T) equipped with a direct 13C-detection probe at 298 K. The experimental
settings and processing parameters for the wild-type protein and K302A variant were
identical. 512 transients were collected with 4K time domain points and a spectral width of
3019 Hz centered at 160 ppm. The spectra were processed using exponential multiplication
with the line broadening factor set to 3Hz. The similarity of protein concentration in both
samples was verified by 1H NMR (not shown).
Crystallization, data collection and processing
PALAO-bound wild-type and mutant AOTCase crystals were grown using the hanging-drop
vapor diffusion method, with conditions similar to those used to produce native and ligand-
complexed AOTCase crystals (18,23). 2.0 µl of ~10 mg/ml solution of AOTCase were
mixed with 1.6 µl of reservoir solution and 0.4 µl PALAO solution (~0.01 M). The reservoir
solution contained 20% (w/v) PEG 3350, 0.2 M lithium sulfate and 0.1 M bis-Tris, pH 6.0.
Diffraction data for the PALAO-bound crystal were collected at 100 K at the F1 beam line
of the Cornell High Energy Synchrotron Source. Data sets for the PALAO-bound mutant
AOTCase crystals were obtained using a Rigaku anode x-ray generator in the Molecular
Structure Section of the National Institute of Health. All data were processed using
HKL2000 package (26) and reduced using the program TRUNCATE in the CCP4 suite (27).
Data collection parameters are listed in Table 1.
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Structure solution and refinement
Molecular replacement was used for phase determination of the PALAO-bound wild-type
and mutant AOTCase structures. The coordinates of AOTCase (PDB code: 3KZO) after
removal of ligands or water molecules were used for phase determination. Upon rigid-body
refinement, electron density corresponding to the ligands could be clearly visualized. The
ligands were built into the map using the graphics program O (28). Refinements were
carried out using molecular annealing, energy minimization and restrained B factor
refinement with the program CNS1.1 (29). During refinements, 5% of the reflections at
various resolutions were randomly selected to set aside to calculate Rfree to monitor the
progress of refinement (30). After every cycle of refinement, the model was manually
adjusted using the program O (28). Water molecules were automatically assigned using the
program WATERPICK of CNS. Model quality was checked using the program
PROCHECK (31) to ensure good stereochemistry for all three models. The final refinement
statistics are listed in Table 1.
Figures 1 was drawn using the programs Pymol (http://www.pymol.org). Figure 3 was
drawn using ChemDraw 8.0. The coordinates have been deposited with the RCSB PDB as
entries 3M4J, 3M4N, 3M5C and 3M5D.
RESULTS
Lys302 in AOTCase is carboxylated
To investigate the nature of the modification of Lys302 and how it affects catalytic activity,
we revisited all AOTCase structures. In the PALAO-bound AOTCase structure, the electron
density map clearly indicates that Lys302 is post-translationally modified (Figure 1A). The
type of modification can include methylation, acetylation, carbamylation, and carboxylation.
The shape of the electron density can been used to distinguish methyl groups from larger
functional groups, but it is difficult to distinguish between acetyl, carbamyl, and carboxyl
groups, all of which have three non-hydrogen atoms in a plane. Given the hydrogen bonding
network with surrounding residues (Lys252, Ser253 and His293, Table 2), a carboxylated
modification is the most likely choice for the modification of Lys302 in AOTCase. To
exclude that the modification’s identity represents chemically stable moieties (methyl,
acetyl, carbamyl), we analyzed trypsin digested fragments of purified AOTCase by TOF-
TOF mass spectroscopy. As expected, only a peptide fragment with an unmodified Lys302
was observed, consistent with the lability of the carboxylic group in acidic solutions. At low
pH, the carboxyl group is spontaneously released as carbon dioxide (14, 32), in contrast to
other modified groups that are stably bound and can be observed by mass spectrometry
analysis after proteolysis (33).
The putative carboxyl group on the modified Lys302 forms direct hydrogen bonds with
main-chain or side-chain nitrogen atoms of Lys252, Ser253 and His293 (Figure 1A and
Table 2). Among these, Lys252 is involved in direct hydrogen bonding to the carboxyl
group of the AORN moiety of PALAO, and His293 forms a strong hydrogen bond with the
main-chain nitrogen atom of Leu295 in the conserved His293-Cys294-Leu295-Pro206
(HCLP) motif. The hydrogen bonding network between the carboxyl group of modified
Lys302, His293 and the main-chain nitrogen atom of Leu295 is reminiscent of the similar
hydrogen bonding network, Glu310-His302-Leu304 and Glu299-Leu272-Leu274, found in
human and E. coli OTCase, respectively (34, 35). These three residues are conserved in all
OTCase sequences, and the interactions between them are important for maintaining the
HCLP motif in a specific conformation to orientate their main-chain oxygen atoms towards
the active site. In all known transcarbamylase structures, a leucine residue corresponding to
Leu295 is in an energetically unfavorable conformation and the peptide bond between this
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leucine and Pro296 is in the cis conformation. In addition to the direct hydrogen bonding
interaction above, the carboxyl group on the modified Lys302 interacts with the α-amino
nitrogen atom of the AORN moiety of PALAO and Glu92 from the adjacent subunit via
water molecules.
When we revisited all previously determined AOTCase structures (see supplementary
Figure S1) we found: (1) Lys302 was carboxylated in the absence of substrate binding, but
substrate binding immobilizes the side-chain of Lys302 further by hydrogen bonding
interaction via water molecules. (2) Water-mediated hydrogen bonding promotes
interactions of carboxylated Lys302 with Glu92 from the adjacent subunit and the α-amino
nitrogen atom of AORN. (3) Similarly to AOTCase, in the structure of SOTCase E92Z (Z =
Ala, Ser, Pro, Val), mutant with N-succinyl-L-norvaline bound (22) the carboxylated Lys302
hydrogen bonds to the α-amino nitrogen atom and the succinyl carboxyl group of N-
succinyl-L-norvaline via water molecules (Figure S1).
To obtain direct, independent evidence for the carboxylation of Lys302, 13C NMR
experiments were carried out with both wild-type protein and the K302A mutant. As
observed for other proteins with carboxylated lysine (36,37), the strong 13C NMR signal at
164 ppm characteristic of a carboxyl group was clearly detectable in AOTCase wild-type
protein labeled by 13C-bicarbonate, in contrast to the K302A mutant where the signal was
weak (Figure 2). Since there are 17 other lysine residues in the protein, the weak signal seen
for the K302A mutant might be caused by the adventitious carboxylation of another lysine
with reduced pKa, as has been observed for the K392A mutant of the sensor domain of the
BlaR protein (38).
Functional and structural studies of Lys302 mutants
To investigate the effect of lysine carboxylation on enzyme activity, Lys302 was mutated to
alanine, glutamate or arginine. Each of these variants was expressed in E. coli and gave
similar yields. Enzymatic assays demonstrated a significant decrease in enzymatic activity in
all three mutants, reflecting the functional importance of Lys302 (Table 3). The level of
enzymatic activity for the wild-type (WT) and three mutants was WT > K302A > K302E ≫
K302R. To determine the structural basis of these results, the WT and mutant enzymes
bound with PALAO were crystallized and their structures were determined at 1.8–2.2 Å
resolution. Only the K302R mutation had and appreciable effect on the structure of the
protein. Since K302 is located near the AORN binding site, the mutations would weaken
AORN binding to the active site.
In the structure of the K302A mutant, three additional water molecules (labeled as w3, w4
and w5 in Figure 1B) replace the carboxylated lysine. The two water molecules (labeled w1
and w2 in Figure 1A–1D) that mediate the hydrogen bonding interaction of carboxylated
Lys302 with PALAO and Glu92 from the adjacent subunit are also found in the K302A
mutant structure. Furthermore, these water molecules maintain a similar hydrogen-bonding
network to the wild-type enzyme. These results might explain why the K302A mutant
retains significant catalytic activity (Table 3). To investigate whether adding short-chain
carboxylic acids to the K302A mutant increases its activity as other enzymes (14, 15, 39,
40), the activity of the K302A mutant was measured in the presence of high formate and
acetate concentration (0.5 M). Surprisingly, the activity of the K302A mutant was not
significantly improved. The crystal structure of the K302A mutant soaking with the
crystallization buffer in the presence of 0.5 M acetate was also determined (not shown) and
it was observed that the same five water molecules were present in the cavity that replaced
the side chain of the carboxylated lysine. This, the acetate’s inability to replace the water
molecules in the crystal structure, is consistent with the unchanged activity assay results.
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The side-chain of Glu302 in the K302E mutant structure is well defined and anchored by
hydrogen bonding interaction with the main-chain nitrogen atom of Arg298 and weakly
hydrogen bonded to the main-chain nitrogen atom of Ser253 (Figure 1C). Two of three
additional water molecules (w4 and w5) observed in the K302A mutant structure occupied
the same position as the carboxyl oxygen atoms of Glu302 and form a similar hydrogen-
bonding network. Relative to the PALAO-bound wild-type structure, there is only one more
water molecule (w3) at the position of the carboxyl group of the carboxylated Lys302. This
water molecule mediates a hydrogen bonding interaction between Glu302 and Lys252. Two
common water molecules (w1 and w2) that interact with PALAO and Glu92 from the
adjacent subunit, respectively, were also identified in the K302E structure.
Our observation that the K302E mutant had lower enzymatic activity than that of the K302A
mutant (Table 3) was surprising since the carboxyl group of the glutamate could conceivably
function similarly to a carboxylated lysine. The explanation may be that, in the K302A
mutant, the hydrogen bonding network is well maintained by water molecules in the cavity
that replaces the carboxylated lysine. In particular, w3 is optimally located for strong
hydrogen bonding to w1 (2.7 Å), which in turn binds AORN. The distances between w1 and
the carboxyl oxygen of carboxylated Lys302 in all wild-type crystal structures are within
2.4–2.7 Å, but the distance between w1 and w2 in the K302E structure is significantly
greater (3.2 Å). The weaker hydrogen bonding interaction may be a reason for lower
enzymatic activity of the K302E mutant.
In contrast to the K302A and K302E structures, the K302R structure shows a much larger
reduction in enzyme activity relative to the wild-type enzyme. The electron density for the
side-chain of Arg302 is weak and the temperature factor of its side-chain is 54.4 Å2,
significantly higher than those of carboxylated Lys302 (44.7 Å2) and Glu302 (33.4 Å2),
implying greater flexibility. Furthermore, the side-chain of Arg302 is oriented differently
from the carboxyl group of carboxylated Lys302 and pushes the nearby residues His180,
Pro181 and Lys182 outwards about 1.0 Å (Figure 1D). However, the water molecules
involved in hydrogen bonding to the α-amino nitrogen atom of PALAO (w1) and the side-
chains of Lys252 (w3) and Glu92 (w2) from the adjacent subunit are conserved. Consistent
with the K302E structure, the distance between w1 and w2 is even greater (3.4 Å) than in
the WT structure and the hydrogen bonding interaction between w2 and w3 is no longer
observed. Thus, the almost undetectable enzymatic activity of the K302R mutant probably
results from the changes at its active site, including the weakened hydrogen bonding
network involved in substrate binding.
DISCUSSION
Several lines of evidence clearly indicate that Lys302 in AOTCase is carboxylated. First, the
extra electron density indicates that the side-chain of Lys302 is modified. Second, the
hydrogen bonding environment of Lys302 for hydrogen bonding interactions is compatible
with a carboxyl group, but not for a positively charged lysine side-chain. Third, the
modification is labile at low pH, since mass spectroscopy of samples prepared at low pH
indicated that Lys302 was no longer modified. Fourth, the clear presence of the
indicative 13C NMR signal at 164 ppm for wild-type protein and its absence in the K302A
mutant confirms carboxylation of Lys302.
It is well known that lysine carboxylation is non-enzymatic and reversible, while other post-
translational modifications such as methylation, acetylation, and carbamylation are
irreversible and detectable by mass spectroscopy. Furthermore, lysine methylation and
acetylation usually require an enzyme-catalyzed reaction in vivo (41). Therefore, it is
unlikely that such lysine modifications will be observed in recombinant proteins
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overexpressed in a foreign host (e.g. E. coli). Lysine methylation can be achieved by using
special chemicals in vitro, but these chemicals are not present in vivo. Lysine carbamylation
and carboxylation use completely different mechanisms to form functionally different
groups (Figure 3). Carbamylation can be achieved by cyanate produced from
myeloperoxidase-catalyzed oxidation of thiocyanate, an anion abundant in plasma and
increased in smokers, or from urea in the plasma. Lysine carboxylation, on the other hand,
occurs readily in aqueous solution in the presence of carbon dioxide at a basic pH (32,42).
Even though carbamylation and carboxylation use very different mechanisms, the two are
confused in the literatures. Lysine carbamylation (or carbamoylation) is referred to in
several publications (15,32,42–44), when the actual reaction is in fact carboxylation.
The activity of the K302A mutant is almost half of that of the wild-type enzyme raising the
question of why AOTCase retains a lysine in this position. Perhaps this lysine was
maintained through evolution to distinguish AOTCase from SOTCase which uses N-
succinyl-L-ornithine (SORN) rather than AORN (22), and OTCase which uses L-ornithine.
An alternative explanation may be found in the very low activity of the K302R mutant. The
side-chain of arginine has a positive charge while carboxylated lysine has a negative charge.
The side chain of unmodified lysine is usually located in a similar position as that of
arginine, as observed in the structure of UV damage endonuclease (14). It would be
expected that the activity of AOTCase with an uncarboxylated lysine would be as low as the
K302R mutant’s. It could further be surmised that, the respective organisms need to use
carboxylation as a switch to turn “on” or “off” the arginine biosynthetic pathway. It has been
well known that rubulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) in plant cells
uses the carboxylation on Lys201 as a switch to turn the enzyme “on” during the day and
“off” at night by removing the carboxyl group (45,46). Carbon dioxide and bicarbonate have
been found to play an important biological role in modulating several biological processes
including photosynthetic carbon fixation (47), pH homeostasis (48), carbon metabolism
(49), activation of virulence in pathogenic organisms (50), sperm maturation (51),
stimulation of mammalian G-protein-responsive adenylyl cyclase (52), and as an alarmone
in Drosophila (53,54). Whether or not carboxylation of a key lysine in their related proteins
is used as an underlying regulatory mechanism should be investigated further.
There are 197 structures with carboxylated lysine residue (modified residue indicated as
Kcx) in the Protein Data Bank (PDB). If structures with 90% identity are counted only once,
there are still 52 unique structures remaining in this pool (Table 4). These proteins include
hydantoinase (40,55), folylpolyglutamate synthase (43), UV damage endonuclease (14),
OXA10, OXA-1 class D β-lactamase (38,56,57), urease (42), phosphotriesterase (58),
dihydroorotase (59), dihydropyrimidinase (60), organophosphate hydrolase (61) and MurE
and MurD ligases (44,62). In most of these proteins, the carboxylated lysine bridges two
metal ions, similar to the role of glutamate or aspartate in proteins with two metal-binding
sites (26 structures among 52). However, the urease apoenzyme can be activated in vitro
only in the presence of carbon dioxide prior to nickel binding (63), suggesting that the
carboxylated lysine may have other structural roles beyond binding metals. In some proteins
such as β-lactamase, UV damage endonuclease, Rubisco, MurD and MurE ligase and BlaR
signal transducer protein, a carboxylated lysine plays an essential catalytic role. More
interestingly, in three structures (PDB ID: 1HL9, 1PU6 and 2UYN for fucosiadase, 3-
methyladenine DNA glycosylase and TdcF protein of unknown function, respectively), the
carboxylated lysines are located near the surface of proteins, presumably playing primarily a
structure stabilizing role (64–66). Since the carboxyl group is labile at acidic pH, but easily
formed in the presence of carbon dioxide at basic pH, the number of proteins with lysine
carboxylation must be underestimated. Furthermore, the carboxylated lysine must be fixed
in place by metal ions (either one or two) or hydrogen bonding with other protein residues
(at least one). Therefore, any detection method involving denaturing the proteins will result
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in release of the carboxyl group. With current technology, 13C NMR (38) and
crystallography are the only methods that can detect this modification. However, these
methods are not amenable to high-throughput investigations. The majority (49 out of 52
structures in the PDB) of known lysine carboxylation modifications were found to be
located at or near the active site, probably because these sites receive the most attention.
Revisiting the structures in PDB with more attention to surface lysines might reveal more
structures with carboxylated lysines.
In conclusion, we have shown that Lys302 in AOTCase is post-translationally modified by
carboxylation and that this modification may be functionally important for enzymatic
activity. Lysine carboxylation is likely to be a more common event than currently
appreciated and may play a critical role in enzymatic activity and protein stability.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Abbreviations
ACIT
N-acteyl-L-citrulline
ANOR
N-acetyl-L-norvaline
AORN
N-acetyl-L-Ornithine
AOTCase
N-acetyl-L-ornithine transcarbamylase
ATCase
aspartate transcarbamlyase
OTCase
ornithine transcarbamylase
CP
carbamyl phosphate
ORN
L-ornithine
PALAO
Nδ-(phosphonacetyl)-Nα-acetyl-L-ornithine
SORN
N-succinyl-L-ornithine
WT
wild-type
xc
Xanthomonas campestris
Acknowledgments
We thank Dr. David Davies for facilitating our use of the diffraction equipment in the Molecular Structure Section
of the National Institute of Health and Dr. Fred Dyda for help in data collection and processing, and Dr. Yui-Fai
Lam in the University of Maryland for help in setting up NMR measurements.
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Figure 1.
Stereo view of the structure and hydrogen bonding network surrounding residue 302. A,
PALAO bound wild-type AOTCase, B, PALAO bound K302A AOTCase C, PALAO bound
K302E AOTCase, D, PALAO bound K302R AOTCase. Contours of the electron density
maps (2Fo-Fc) around PALAO, residue 302 and water molecules are shown as a brown cage
at 1.0σ. The final refined positions of the ligands and surrounding protein residues are
represented as colored sticks. The predicted hydrogen bonding interactions are in pink
dashed lines. The water molecules are represented as pink balls. The carbon of PALAO,
residue 302 and other protein residues are shown in pink, light blue and green sticks,
respectively.
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Figure 2.
13C NMR spectra of wild-type (upper panel) and K302A mutant (lower panel) AOTCase (1
mM). Experiments were performed in 100 mM Tris HCl, 50 mM NaCl, 7% D2O, pH 8.0,
supplemented with 20 mM NaH13CO3. The position of the resonance attributed to
carboxylated lysine in the enzyme is around 164 ppm.
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Figure 3.
Chemical structure of carbamylated vs. carboxylated lysine.
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Table 1
Data collection and refinement statistics
Dataset
PALAO
K302A
K302E
K302R
Space group
I213
I213
I213
I213
Resolution (Å)
2.2
1.9
1.85
2.2
Unit-cell parameters (Å)
a = b = c =128.88
a = b = c =128.92
a = b = c =129.29
a = b = c =127.39
Measurements
219,475
305,757
390,128
246,817
Unique reflections
18,269 (1,832) a
28,236 (1,365)
30,622 (1,456)
17,635 (879)
Redundancy
12.0 (11.8)
10.8(5.4)
12.8 (5.4)
14.0 (13.1)
Completeness (%)
99.8 (100.0)
100.0 (100.0)
99.7 (95.1)
100.0 (100.0)
<I/σ (I)>
15.0 (4.9)
16.4 (2.3)
19.8 (2.8)
8.7 (3.7)
Rmerg b
7.4 (48.4)
6.5(64.9)
5.2 (55.3)
9.8 (79.1)
Wilson B (Å2)
30.4
27.6
28.6
21.9
Refinement
Resolution range (Å)
50.0-2.2
50-1.9
50-1.85
50-2.2
No. of protein atoms
2620
2613
2617
2619
No. of water atoms
90
219
193
146
No. of hetero atoms
24
24
24
24
Rmsd of bond lengths (Å)
0.006
0.005
0.005
0.005
Rmsd of bond angle (°)
1.1
1.2
1.2
1.2
Rwork (%)c
20.0
19.8
20.0
18.9
Rfree (%)d
24.3
23.2
23.2
22.2
Average B factor (Å2)
41.7
32.2
32.3
35.3
aFigures in brackets apply to the highest-resolution shell.
bRmerg = ΣhΣi|I(h,i)-<I(h)>|/∑hΣiI(h,i), where I(h,i) is the intensity of the ith observation of reflection h, and < I(h)> is the average intensity of
redundant measurements of reflection h.
cRwork= Σh‖Fobs| – |Fcalc‖/Σh|Fobs|.
dRfree = Σh‖Fobs| – |Fcalc‖/Σh|Fobs| for 5% of the reserved reflections.
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Table 2
Interactions between carboxylated lysine and other residues at the active site of AOTCase
Kcx302
Other residues
Bound ligands
PALAO
CPa
AORNb
CP +ANORc
SO4+ACITd
OQ1
K252 NZ
2.6
2.6
2.7
2.6
2.6
OQ1
W1e
2.6
2.6
2.6
2.7
OQ2
S253 N
3.0
3.1
2.8
2.9
2.9
OQ2
H293 NE2
3.0
3.2
3.0
2.9
2.9
NZ
W2f
3.1
2.9
3.0
3.0
aThe values were calculated based on PDB ID 3KZM.
bThe values were calculated based on PDB ID 3KZN.
cThe values were calculated based on PDB ID 3KZO.
dThe values were calculated based on PDB ID 3KZK.
eThis water molecule hydrogen bonds to N1 atom of PALAO, AORN or ANOR, and backbone O atom of Pro296 as well.
fThis water molecule hydrogen bonds to OE1 atom of Glu92 from adjacent subunit as well.
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Table 3
Specific activity of wild-type and mutant AOTCase in the presence of acids (0.5M).
Compounds added
Specific activity(µmol/min/mg)
Wild-type
K302A
K302E
K302R
None
43.4 ± 0.4a
23.0 ± 0.5
7.1 ± 0.1
0.059±0.01
Formate
44.1 ± 1.2
26.4 ± 0.6
6.7 ± 0.2
0.093±0.01
Acetate
48.5 ± 1.1
21.2 ± 0.8
6.6 ± 0.5
0.104±0.03
aThe Mean ± S.D. are shown (n = 3).
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Table 4
Protein structures with lysine carboxylation modification
PDB ID
Enzyme name
Residue
Organism source
Funciton
2OGJ
Dihydroorotase
175
A.tumefaciens
Bridging two Zn(II)
2Z26
Dihydroorotase
102
E.coli
Bridging two Zn(II)
3JZE
Dihydroorotase
103
S.enterica
Bridging two Zn(II)
2GWN
Dihydroorotase
149
P. gingivalis
Bridging two Zn(II)
3F4C
Organophosphorus hydrolase
243
G. stearothermophilus
Bridging two Co(II)
3ICJ
Metal-dependent hydrolase
294
P. furiosus
Bridging two Zn(II)
3GTX
Organophosphorus hydrolase
243
D. radiodurans
Bridging two Co(II)
2QPX
Metal-dependent hydrolase
166
L. casei
Bridging two Zn(II)
2FTW
Dihydropyrimidinase
158
D. discoideum
Bridging two Zn(II)
2FVK
Dihydropyrimidinase
167
S. kluyveri
Bridging two Zn(II)
3DC8
Dihydropyrimidinase
147
S. meliloti
Bridging two Zn(II)
3GNH
L-Lys/Arg carboxypeptidase
211
C. crescentus cb15
Bridging two Zn(II)
3DUG
Arginine carboxypeptidase
182
Unidentified
Bridging two Zn(II)
2VC7
Phosphotriesterase
137
S. solfataricus
Bridging two Co(II)
2R1N
Metallophosphotriesterases
169
A. tumefaciens
Bridging two Co(II)
2OB3
Phosphotriesterase
169
B. diminuta
Bridging two Zn(II)
3E74
Allantoinase
146
E. coli
Bridging two Fe(III)
1EJX
Urease
217
K. aerogenes
Bridging two Ni(II)
1E9Z
Urease
219
H. pylori
Bridging two Ni(II)
4UBP
Urease
220
B. pasteurii
Bridging two Ni(II)
1ONW
Isoaspartyl dipeptidase
162
E. coli
Bridging two Zn(II)
1K1D
D-hydanroinase
150
G. stearothermophilus
Bridging two Zn(II)
1GKR
L-hydanroinase
147
A. aurescens
Bridging two Zn(II)
1GKP
D-hydanroinase
150
Thermus sp.
Bridging two Zn(II)
1NFG
D-hydantoinase
148
R. pickettii
Bridging two Zn(II)
2ICS
Adenine deaminase
154
E. faecalis
Bridging two Zn(II)
1RQB
Transcarboxylase
184
P. freudenreichii
Binding one Co(II)
2QF7
Pyruvate carboxylase
718
R. etli
Binding one Zn(II)
3BG3
Pyruvate carboxylase
741
H. sapiens
Binding one Mn(II)
2OEM
Rubisco-like protein
173
G. kaustophilus
Binding one Mg(II)
1WDD
Rubisco
201
O. sativa
Binding one Mg(II)
1GK8
Rubisco
201
C. reinhardtii
Binding one Mg(II)
1BWV
Rubisco
201
G. partita
Binding one Mg(II)
2WTZ
ATP-dependent MurE ligase
262
M. tuberculosis
Binding one Mg(II)
2JFG
MurD ligase
198
E. coli
Catalytic role?
1E8C
MurE ligase
224
E. coli
Catalytic role?
1JBW
Folypolyglutamate synthetase
185
L. casei
Catalytic role?
1W78
FolC bifunctional protein
188
E. coli
Binding one Mg(II)
3HBR
OXA-48 β-lactamase
73
K. pneumoniae
Catalytic role
Biochemistry. Author manuscript; available in PMC 2011 August 17.
NIH-PA Author Manuscript
NIH-PA Author Manuscript
NIH-PA Author Manuscript
Li et al.
Page 21
PDB ID
Enzyme name
Residue
Organism source
Funciton
3ISG
Class D β-lactamase
70
E. coli
Catalytic role
2P9V
AmpC beta-lactamase
315
E. coli
Catalytic role
1K55
OXA-10 β-lactamase
70
P. aeruginosa
Catalytic role
1K38
β-lactamase OXA-2
70
S. typhimurium
Catalytic role
1XQL
Alanine racemase
129
G. stearothermophilus
Binding substrate?
1VFS
Alanine racemase
129
S. lavendulae
Binding substrate?
1RCQ
Alanine racemase
122
P. aeruginosa
Binding substrate?
2J6V
UV damage endonuclease
229
T. thermophilus
Catalytic role
1H01
Cell division protein kinase 2
33
H. sapiens
Catalytic role?
2UYN
Protein TdcF
A58
E. coli
Structural role?
1HL9
Fucosidase
338
T. maritime
Structural role?
1PU6
3-methyladenine DNA glycosylase
205
H. pylori
Structural role?
Biochemistry. Author manuscript; available in PMC 2011 August 17.
|
3M4O
|
RNA polymerase II elongation complex B
|
X-ray structure and mechanism of RNA polymerase II
stalled at an antineoplastic monofunctional
platinum-DNA adduct
Dong Wanga,b,1, Guangyu Zhuc, Xuhui Huangd, and Stephen J. Lippardc,1
aDepartment of Structural Biology, Stanford University School of Medicine, Stanford, CA 94305; bSkaggs School of Pharmacy and Pharmaceutical Sciences,
University of California, San Diego, La Jolla, CA 92093; cDepartment of Chemistry, Massachusetts Institute of Technology, Cambridge, MA 02139; and
dDepartment of Chemistry, Hong Kong University of Science and Technology, Clear Water Bay, Kowloon, Hong Kong, P.R. China
Contributed by Stephen J. Lippard, March 3, 2010 (sent for review February 9, 2010)
DNA is a major target of anticancer drugs. The resulting adducts
interfere with key cellular processes, such as transcription, to
trigger downstream events responsible for drug activity. cis-
Diammine(pyridine)chloroplatinum(II), cDPCP or pyriplatin, is a
monofunctional platinum(II) analogue of the widely used antican-
cer drug cisplatin having significant anticancer properties with a
different spectrum of activity. Its novel structure-activity properties
hold promise for overcoming drug resistance and improving the
spectrum of treatable cancers over those responsive to cisplatin.
However, the detailed molecular mechanism by which cells process
DNA modified by pyriplatin and related monofunctional complexes
is not at all understood. Here we report the structure of a transcri-
bing RNA polymerase II (pol II) complex stalled at a site-specific
monofunctional pyriplatin-DNA adduct in the active site. The re-
sults reveal a molecular mechanism of pol II transcription inhibition
and drug action that is dramatically different from transcription in-
hibition by cisplatin and UV-induced 1,2-intrastrand cross-links. Our
findings provide insight into structure-activity relationships that
may apply to the entire family of monofunctional DNA-damaging
agents and pave the way for rational improvement of monofunc-
tional platinum anticancer drugs.
anticancer ∣chemotherapy ∣DNA damage ∣pyriplatin ∣transcription
T
he DNA template for transcription is not only the site of in-
born errors of metabolism and of continuous attack by harm-
ful environmental agents, but it also represents a major target
for cancer therapy. Platinum-based anticancer drugs such as
cisplatin, cis-diamminedichloroplatinum(II), are widely used
and among the most effective antineoplastic treatments (1, 2).
Platinum-based drugs typically form bifunctional intra- or inter-
strand DNA cross-links by covalent bonding to the N7 positions of
two guanosine residues, triggering a variety of cellular processes,
including
transcription
inhibition
with
attendant
apoptosis
(1, 2). However, resistance and side effects can require with-
drawal of these drugs before they can effect a cure in certain types
of cancer (3).
In the effort to find new compounds that circumvent resis-
tance to conventional bifunctional platinum-based drugs, a class
of monofunctional platinum compounds were synthesized and
screened for anticancer activity (4–6). In contrast to other
inactive
monofunctional
platinum(II)
compounds
such
as
½PtðdienÞClþ and ½PtðNH3Þ3Clþ, cis-diammine(pyridine)chloro-
platinum(II) [cDPCP or “pyriplatin” (Fig. 1)] and related com-
plexes display significant anticancer properties and a different
spectrum of activity compared to conventional platinum-based
drugs. These features render them attractive candidates for treat-
ing cisplatin-refractory patients if the potency could be raised to
or beyond the level of that of cisplatin (4, 5, 7). Pyriplatin exhibits
unique chemical and biological properties, forming monofunc-
tional DNA adducts (Fig. 1 and Fig. S1) that can inhibit transcrip-
tion and better elude DNA repair (7). The x-ray crystal structure
of pyriplatin bound to a DNA duplex reveals substantially
different features than those of DNA adducts formed by conven-
tional, bifunctional platinum-based drugs. The overall DNA
duplex is much less distorted, with the pyridine ligand of the
cis-fPtðNH3Þ2ðpyÞg2þ moiety directed toward the 50-end of the
platinated strand. A hydrogen bond forms between the NH3
ligand trans to pyridine and O6 of the platinated guanosine
residue (7).
The detailed molecular mechanism by which cells process
DNA modified by monofunctional complexes such as pyriplatin
is not understood. Several important questions remain unan-
swered. By what process do monofunctional adducts block pol
II transcription? Does the mechanism differ from that of tran-
scription inhibition by 1,2- and 1,3-intrastrand cross-links that
comprise the major adducts of cisplatin? Why do pyriplatin
and its homologues, which violate the classical structure-activity
relationships (SARs) for active, bifunctional platinum drugs (8),
show such promise by comparison to related monofunctional
complexes like ½PtðNH3Þ3Clþ? Would knowledge of the struc-
ture of pyriplatin-modified DNA at its site(s) of biological action
inform the design of more potent analogues?
In the present work we take a combined biochemical and x-ray
structural approach to investigate the molecular mechanism of
pol II transcription inhibition by a site-specific monofunctional
platinum(II)-DNA adduct of pyriplatin. An unprecedented mo-
lecular mechanism for pol II transcription inhibition is revealed,
providing insight into structure-activity relationships that may ap-
ply to the entire family of monofunctional DNA-damaging
agents, whether or not they contain platinum.
Results
A Different Configuration of a Pyriplatin-DNA Adduct Accommodated
in the Pol II Active Site. To understand how a monofunctional
pyriplatin-DNA adduct is accommodated in the active site of
the transcribing pol II elongation complex, we designed and pre-
pared a DNA template containing a site-specific DNA lesion of
this complex, as described previously (7). A transcribing pol II
complex was then assembled in which the pyriplatin-DNA lesion
occupies the active (þ1) site (Complex B, Table 1). The crystal
structure of this complex reveals that the platinated nucleotide
is captured as a pol II complex in the post-translocation state,
in which the addition site is empty and ready for NTP loading
(Dashed Ring, Fig. 2A and Fig. S2). Fig. 2A reveals that the
Author contributions: D.W. and S.J.L. designed research; D.W., G.Z., and X.H. performed
research; D.W., G.Z., X.H., and S.J.L. analyzed data; and D.W., X.H., and S.J.L. wrote
the paper.
The authors declare no conflict of interest.
Data deposition: The atomic coordinates have been deposited in the Protein Data Bank,
www.pdb.org (PDB ID codes 3M4O and 3M3Y).
1To whom correspondence may be addressed. E-mail: dongwang@ucsd.edu or lippard@
mit.edu.
This article contains supporting information online at www.pnas.org/cgi/content/full/
1002565107/DCSupplemental.
9584–9589 ∣PNAS ∣May 25, 2010 ∣vol. 107 ∣no. 21
www.pnas.org/cgi/doi/10.1073/pnas.1002565107
positioning of the pyriplatin-damaged guanosine residue is lo-
cated above the bridge helix. This structure requires rotation
of the cis-fPtðNH3Þ2ðpyÞg2þ moiety and its bound guanosine re-
sidue into a different configuration compared to that adopted in
the pyriplatin-duplex DNA structure, in order to avoid a steric
clash with bridge helix (7). Fig. 2B depicts this comparison.
The rotation is energetically facilitated by the formation of hydro-
gen bonds between the ammine ligands on platinum with the
phosphodiester moiety of the backbone between positions þ1
and þ2, with concomitant loss of a hydrogen bond between O6
of the platinated guanosine residue and an ammine ligand. An
additional feature is that the pyridine group of the cis-
fPtðNH3Þ2ðpyÞg2þ fragment, which points downstream toward
the 50-direction of the template DNA, forms van der Waals inter-
actions with bridge helix residues Val 829 and Ala 832. The purine
base of the guanosine residue at position þ1 is displaced toward
the major groove of the RNA–DNA duplex by comparison with
structures having an undamaged base at this site in the post-trans-
location state (9–11).
Transcription Elongation Inhibited by a Pyriplatin–DNA Adduct. Be-
cause transcription inhibition is an important component in
the mechanism of action of platinum anticancer drugs (12–20),
we investigated the effect of a site-specific pyriplatin–DNA ad-
duct on the kinetics of pol II transcription elongation. We per-
formed an extension assay using platinated (Complex A,
Table 1) and unplatinated (Complex A0, control, Table 1) pol
II transcribing complexes having a 9mer RNA as primer. These
complexes were then incubated with a mixture of ATP, CTP, and
GTP. The RNA transcripts in A could be elongated from the 9mer
to the 11mer, stopping at a position corresponding to the Pt–
DNA lesion site observed in the pol II complex of the damaged
template DNA, whereas RNA transcripts in A0 were extended
much farther downstream on the undamaged template control
DNA (Fig. 3A). In order to avoid the possibility of misincorpora-
tion-induced transcription inhibition in this assay, we carried out
a similar extension assay using an RNA containing a 30-end CMP
matched against the damaged base (pol II complex C, 11mer)
(Table 1). A single matching GTP was incubated with this pol
II complex to test whether the enzyme could bypass the Pt–
DNA lesion. Consistent with the results of the previous assay,
RNA transcripts could not be extended beyond an 11mer in
the pol II complex with the damaged DNA template, whereas
RNA transcripts were efficiently extended farther downstream
along the undamaged DNA template (Fig. 3B). Similar extension
assay results were obtained using a chain-terminated GTP analo-
gue 30-dGTP or an RNA primer of different length (complex B,
10mer) (Table 1) (Fig. 3 C and D). Finally, to investigate whether
the presence of the damaged base affects the rate of NTP incor-
poration in a single round, we used complex B (10mer) and com-
plex C (11mer), incubating with CTP and 30-dGTP, respectively.
For CTP incorporation, RNA transcripts could be efficiently ex-
tended from the 10mer to the 11mer using both damaged and
nondamaged templates at a comparable rate (Fig. 3E), whereas
no obvious extension of RNA transcripts from the 11mer to a
12mer was observed on the damaged DNA template (Fig. 3C).
UTP failed to incorporate at the damaged template under the
same conditions (Fig. S3A). No obvious intrinsic cleavage was
observed for a pol II complex containing the 11mer RNA and
Pt-damaged DNA template in the presence of 20 mM Mg2þ
ion, suggesting that most of complex C (11mer) is not in the back-
tracked state (Fig. S3B) (21–23).
X-ray Structure of Pol II stalled at a Pyriplatin–DNA Adduct. To under-
stand the nature of the pol II complex stalled at the pyriplatin-
induced Pt–DNA adduct, we solved the x-ray crystal structure
Fig. 1.
Scheme depicting the formation of a monofunctional platinum-DNA
adduct by pyriplatin on double-stranded duplex DNA. The structure of the
pyriplatin-damaged DNA duplex used coordinates from the PDB (code
3CO3). The damaged and nondamaged DNA strands are shown in cyan
and green, respectively. The pyridine ligand and two ammine groups of
the cis-fPtðNH3Þ2ðpyÞg2þ moiety are depicted in magenta and blue, respec-
tively. The platinum atom and nitrogen atoms of the cis-fPtðNH3Þ2ðpyÞg2þ
moiety are highlighted in yellow and as a blue ball, respectively. The termini
of the DNA strands are labeled.
A
B
+1
-1
+1
-1
5’
3’
3’
5’
5’
3’
Non-template
DNA
Bridge
Helix
Bridge
Helix
Addition
Site
Addition
Site
5’
V829
A832
RNA
RNA
Template
DNA
Template
DNA
3’
Fig. 2.
Structure of a pol II transcribing complex encountering a site-specific
pyriplatin-dG adduct in DNA. (A) A site-specific pyriplatin-DNA adduct is ac-
commodated in the pol II active site. The view is a standard one, from the
“Rpb2 side,” as described elsewhere (9–11, 39). The RNA transcript, template
DNA strand, and nontemplate DNA strand are depicted in red, cyan, and
green, respectively. Parts of the bridge helix (Rpb1 825–848) are shown in
gray. The pyriplatin-damaged guanosine is colored magenta. The platinum
atom of the cis-fPtðNH3Þ2ðpyÞg2þ moiety is denoted as a yellow ball and
the two ammine groups are in blue. The dashed oval represents the empty
nucleotide addition site in the post-translocation state. The positions of the
RNA strand are labeled. (B) cis-fPtðNH3Þ2ðpyÞg2þ-dG in the pol II active site
adopts a different configuration in comparison with its conformation in
the
structure
of
pyriplatin-modified
duplex
DNA.
The
superimposed
geometry of the cis-fPtðNH3Þ2ðpyÞg2þ-guanosine unit from the DNA duplex
structure (3CO3) is shown in light blue. Side chains of Val 829 and Ala 832 are
depicted in orange. The remainder of the figure is the same as in A.
Wang et al.
PNAS
∣
May 25, 2010
∣
vol. 107
∣
no. 21
∣
9585
BIOCHEMISTRY
of the enzyme in complex with a platinated DNA using an RNA-
containing CTP matched against the damaged guanosine residue.
In this structure, pol II is in pre-translocation state, with the newly
added CMP still occupying the addition site without transloca-
tion. The platinated guanosine residue forms Watson–Crick base
pairs with the newly added CMP (Fig. 4 A and B and Fig. S4). The
cis-fPtðNH3Þ2ðpyÞg2þ moiety is surrounded by the bridge helix at
the bottom, part of the Rpb2 fork region (528–534) on the left
side, and the sugar-phosphate backbone connecting template
DNA positions þ1 and þ2 on the right side (Fig. 4B). Interest-
ingly, upon CMP incorporation, the cis-fPtðNH3Þ2ðpyÞg2þ moiety
adopts a different conformation. The pyridine group of this unit
now faces toward 30-direction of template DNA (Fig. 4 A and B).
The ammine group trans to pyridine is directed toward the bridge
helix and forms hydrogen bonds with main chain atoms of Ala 828
and the side chain of Thr 831 (Fig. 4B). The residues in the bridge
helix are highly conserved from yeast to humans. Because Thr 831
and Ala 828 are absolutely conserved between S. cerevisiae and
humans, the interactions we observe in the S. cerevisiae pol II
structure will also occur in human pol II.
These structural results provide important insights into the
transcription stalling process at a monofunctional pyriplatin–
DNA adduct. The adduct adopts a significantly different confor-
mation within the pol II active site compared to that in duplex
DNA (7). The present structural and biochemical evidence
reveals that pol II stalls after efficient incorporation of CTP
against the damaged guanosine residue. The conformation of
the pyriplatin–DNA adduct changes significantly upon incorpora-
tion of CTP. The modified guanosine rotates into the pol II active
site and serves as a template for base pairing with the matched
substrate, and the cis-fPtðNH3Þ2ðpyÞg2þ moiety is now directed
toward 30-end of the platinated DNA.
Table 1. RNA and DNA scaffold of pol II transcribing complexes
Complex A: (Damaged template 29mer with 9mer RNA)
RNA: 5′
AUGGAGAGG
3′
DNA: 3′ CTACCTCTCCTG*CCCCACCAATACCCATC 5′
DNA: 5′
GTGGTTATGGGTAG 3′
Complex A′: (Nondamaged template 29mer with 9mer RNA)
RNA: 5′
AUGGAGAGG
3′
DNA: 3′ CTACCTCTCCTGCCCCACCAATACCCATC
5′
DNA: 5′
GTGGTTATGGGTAG
3′
Complex B: (Damaged template 29mer with 10mer RNA)
RNA: 5′
AUGGAGAGGA 3′
DNA: 3′ CTACCTCTCCTG*CCCCACCAATACCCATC 5′
DNA: 5′
GTGGTTATGGGTAG 3′
Complex B′: (Nondamaged template 29mer with 10mer RNA)
RNA: 5′
AUGGAGAGGA 3′
DNA: 3′ CTACCTCTCCTGCCCCACCAATACCCATC
5′
DNA: 5′
GTGGTTATGGGTAG
3′
Complex C: (Damaged template 29mer with 11mer RNA)
RNA: 5′
AUGGAGAGGAC3′
DNA: 3′ CTACCTCTCCTG*CCCCACCAATACCCATC 5′
DNA: 5′
GTGGTTATGGGTAG 3′
Complex C′: (Non-damaged Template 29mer with 11mer RNA)
RNA: 5′
AUGGAGAGGAC3′
DNA: 3′ CTACCTCTCCTGCCCCACCAATACCCATC
5′
DNA: 5′
GTGGTTATGGGTAG
3′
G*: cDPCP-dG.
Fig. 3.
Pol II transcription elongation blocked by a site-specific pyriplatin-
DNA adduct. (A) In vitro transcription with preformed pol II elongation com-
plexes A and A0 incubated with a mixture of ATP, CTP, and GTP (25 μM each).
Time points were taken after 0, 0.5, 1, 2, 3, 4, 8, 16, 32, or 64 min incubation.
The RNA transcripts in lanes 1–10 were taken from reactions of the pol II com-
plex with a nondamaged DNA template, whereas the RNA transcripts in lanes
11–20 were taken from reactions of the pol II complex with a site-specifically
damaged DNA template. The stalled RNA transcript is indicated by a black
arrow (Right), and the extended RNA transcript is visible to the left. The
length and sequences of RNA transcripts are given at the left margin of
the gel. (B) In vitro transcription with preformed pol II elongation complexes
C and C’ incubated with 25 μM GTP. The remainder of gel is the same as in A.
(C) In vitro transcription with preformed pol II elongation complexes C and C’
incubated with 25 μM of 30-dGTP. The rest of gel is same as in A. (D) In vitro
transcription with preformed pol II elongation complexes B and B’ incubated
with a mixture of 25 μM CTP and 30-dGTP. Time points were taken after 0, 0.5,
1, 2, 4, 8, 16, 32, 64 min of incubation. The rest of gel is the same as in A. (E) In
vitro transcription with preformed pol II elongation complexes B and B’
incubated with 25 μM CTP. The remainder of the gel is the same as in A.
9586
∣
www.pnas.org/cgi/doi/10.1073/pnas.1002565107
Wang et al.
The result is that the RNA transcript fails to extend beyond the
site of damage, subsequent translocation and nucleotide addition
being strongly inhibited. Several factors contribute to such
translocation inhibition, including (i) stabilization of the initial
pre-translocation state by interaction between the platinated
guanosine and pol II residues (Fig. 4B); (ii) a high translocation
energy barrier; and (iii) an unfavorable subsequent post-translo-
cation state induced by the DNA lesion. Hydrogen bonding inter-
actions between an ammine group of the cis-fPtðNH3Þ2ðpyÞg2þ
moiety with bridge helix partially help to stabilize the initial pre-
translocation state (Fig. 4B). To address the factors ii and iii, we
modeled the pyriplatin-damaged guanosine residue at the −1 po-
sition to mimic the state following translocation of the pyriplatin-
modified guanosine from the þ1 to −1 position. The structure
clearly reveals that the cis-fPtðNH3Þ2ðpyÞg2þ moiety serves as
a strong steric block, narrowing the space between the DNA
nucleotide base (−1) and the bridge helix and preventing the
downstream undamaged nucleoside base on the DNA template
strand from rotating into the canonical þ1 position (Fig. 5A).
Moreover, the fact that the cis-fPtðNH3Þ2ðpyÞg2þ moiety at
the −1 position sterically clashes with the downstream nucleotide
base at the þ1 position suggests that this final state is unfavorable
(Fig. 5 A and B). In summary, our results indicate that pyriplatin–
DNA adducts inhibit pol II transcription elongation by prevent-
ing subsequent translocation and nucleotide addition beyond the
site of damage.
Discussion
Insights into Structure-Activity Relationships (SARs) for the Monofunc-
tional Platinum Drug Family. The original SARs pertaining to
bifunctional platinum compounds such as cisplatin (8) were for-
mulated to explain why anticancer activity appeared to require
neutral, cis-[PtA2X2] compositions, in which A is an amine ligand
and X is a monoanionic leaving group. These rules are clearly
violated by cationic, monofunctional platinum compounds such
as pyriplatin (4, 5). Other monofunctional platinum complexes,
including ½PtðdienÞClþ, ½PtðNH3Þ3Clþ, and trans-½PtðNH3Þ2
ðpyÞClþ, are inactive and do not arrest pol II transcription,
whereas the cis-fPtðNH3Þ2ðpyÞg2þ unit bound to guanosine
blocks pol II transcription and has significant anticancer proper-
ties in mice when administered as pyriplatin (4, 5, 8, 24–32).
The present structure of pol II in complex with DNA site-
specifically modified by pyriplatin provides unique insight into
SARs to be expected for monofunctional platinum drug candi-
dates. We constructed models of potential stalled transcription
complexes containing DNA modified by the following three
representative
units,
fPtðNH3Þ3g2þ, trans-fPtðNH3Þ2ðpyÞg2þ,
and cis-fPtðNH3Þ2ðpyÞg2þ bound to guanosine in DNA and posi-
tioned in either the −1 or þ1 site of pol II, in order to mimic the
A
B
+1
-1
+1
-1
Bridge Helix
Bridge
Helix
Rpb2 528-534
+1
5’
-1
3’
5’
3’
T831
A828
5’
3’
5’
3’
Non-template
DNA
5’
3’
+2
RNA
Template
DNA
Template
DNA
RNA
3.9 Å
3.9 Å
Fig. 4.
Structure of pol II transcribing complex stalled at a site-specific
pyriplatin-DNA adduct after CMP incorporation. (A) The newly incorporated
matched CMP is highlighted in yellow. Other colors are as in Fig. 2. Interac-
tions of the damaged nucleotide and pol II residues are highlighted in (B).
The view is taken roughly from an ∼90 degree clockwise rotation along
the RNA/DNA helix axis from A. Nitrogen and oxygen atoms are depicted
in blue and red, respectively. Hydrogen bonds between ammine group of
the cis-fPtðNH3Þ2ðpyÞg2þ moiety and bridge helix residues are shown as black
dashed lines. The loop of Rpb2 828–834 is shown in green.
X
A
+1
-1
-2
X
+1
-1
-2
RNA
Template
DNA
Bridge
Helix
Non-template
DNA
Addition
Site
3’
5’
5’
5’
3’
3’
+1
-1
-1
+1
+2
-2
Bridge Helix
Addition
Site
3’
3’
5’
Template
DNA
RNA
5’
3’
B
Fig. 5.
Pol II translocation following CMP incorporation is inhibited by a site-
specific pyriplatin-DNA adduct. (A) The cis-fPtðNH3Þ2ðpyÞg2þ-guanosine unit
is superimposed with a nucleoside in −1 position shown in magenta and as a
surface view. In the latter, the nitrogen and oxygen atoms are highlighted in
blue and red, respectively. CMP at the 30-end of RNA chain is highlighted in
yellow. The bridge helix is shown in gray as a surface view. The nucleosides at
the þ1 and þ2 position of the template DNA are drawn in wheat and orange,
respectively. The rotation of the downstream nucleoside base during trans-
location, from the þ2 position to the þ1 position, is blocked by the cis-
fPtðNH3Þ2ðpyÞg2þ moiety, as indicated. Other colors are as in Fig. 2. (B)
The cis-fPtðNH3Þ2ðpyÞg2þ moiety of pyriplatin-dG adduct modeled at −1 posi-
tion clashes with the base at the þ1 position. Colors are as in A, and the view
as in Fig. 4B.
Wang et al.
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post- and pre-translocation states, respectively (Fig. S1). For
each modeled structure, we rotated the platinum unit about
the Pt-N7 bond by 360° and computed van der Waals energies
arising from contacts between platinum ligands and the rest
of the pol II complex (Figs. S5–S9). The fPtðNH3Þ3g2þ and
trans-fPtðNH3Þ2ðpyÞg2þ moieties could be readily accommo-
dated within the pol II active site over wide energy minima.
The lack of a significant steric clash for these two groups, in either
the −1 or þ1 position of the pol II transcribing complex, indicates
the absence of a barrier to transcriptional bypass (Figs. S6–S9).
This finding agrees with experiment. In contrast, the energy bar-
rier is prohibitively high for cis-fPtðNH3Þ2ðpyÞg2þ platinated
DNA modeled at −1 position, which is consistent with its ability
to block pol II bypass and the failure of pol II to reach the sub-
sequent post-translocation state (Figs. S5 and S8). The presence
of a pyridine or other bulky group in the cis configuration is
important
for
restricting
the
rotation
range
of
the
cis-
fPtðNH3Þ2ðpyÞg2þ moiety and thus rendering it a strong steric
block to translocation. For a fPtðNH3Þ3g2þ or trans-fPtðNH3Þ2
ðpyÞg2þ adduct at the −1 position, such a steric clash can be
avoided by rotation about the Pt-N7 bond, facilitating subsequent
pol II translocation. These results are fully consistent with
previous biochemical studies revealing that the latter two
DNA adducts are inactive and fail to block transcription (5, 7,
12, 26–33).
A Unique Molecular Mechanism of Pol II Transcription Inhibition. The
stalling mechanism of monofunctional platinum drugs of the
pyriplatin family is dramatically different from transcription inhi-
bition by cisplatin and UV-induced 1,2-intrastrand cross-links.
For the latter two DNA-modifications, a translocation barrier
prevents delivery of damaged bases to the active site and/or leads
to misincorporation of NTPs against the damage site, respectively
(19, 34). Monofunctional platinum-damaged residues, on the
other hand, can cross over the bridge helix and be accommodated
in the pol II active site. For Pt–dG adducts, the correct CMP
nucleotide can be efficiently incorporated against the damaged
guanosine. It is blockage of the subsequent translocation from
this position after incorporation of the cytosine nucleotide that
leads to inhibition of the RNA polymerase, but only when a bulky
pyridine ligand is present in the cis coordination site.
In conclusion, we report here the structure of a pol II transcri-
bing complex stalled at a site-specific monofunctional DNA
adduct, revealing a unique mechanism of transcription inhibition
by this kind of genome damage. The results establish a basis for
SARs that govern the anticancer drug potential of monofunc-
tional platinum-based DNA-damaging agents. Specific inter-
actions between pol II active site residues and the platinum
ligands are revealed, providing a structural framework for
rational design of more potent monofunctional pyriplatin analo-
gues. Because the spectrum of activity of pyriplatin is dramatically
different from that of cisplatin against an extensive panel of can-
cer cell lines but with reduced potency (7), this information will
be valuable for increasing the anticancer drug potential of this
family of compounds based on pol II stalling with concomitant
induction of apoptosis.
Methods
Preparation of Pol II Transcribing Complexes. Ten-subunit S. cerevisiae pol II
was purified as described (35). RNA oligonucleotides were purchased from
Dharmacon and DNA oligonucleotides were obtained from IDT. cis-
½PtðNH3Þ2ðpyÞClCl was prepared by Ryan Todd at MIT. The site-specifically
platinated template DNA was obtained as described (7).
Pol II transcribing complexes were assembled with the use of synthetic oli-
gonucleotides (10). Briefly, DNA and RNA oligonucleotides were annealed
and mixed with pol II in 20 mM Tris (pH 7.5), 40 mM KCl, and 5 mM DTT.
The final mixture contained 2 μM pol II, 10 μM site-specific pyriplatin-
damaged template DNA strand, and 20 μM nontemplate DNA and RNA oli-
gonucleotides. The mixture was kept for 1 h at room temperature, and excess
oligonucleotides were removed by ultrafiltration. Crystals were obtained
from solutions containing 390 mM ðNH4Þ2HPO4∕NaH2PO4, pH 5.9–6.3,
50 mM dioxane, 10 mM DTT, and 9–11% PEG6000. Crystals of transcribing
complexes were transferred in a stepwise manner to cryobuffer as described
(10, 11). For the structure of the pol II complex with CTP incorporation, 10 mM
CTP was added to the cryobuffer (10, 11).
Crystal Structure Determination and Analysis. Diffraction data were collected
on beam line 11-1 at the Stanford Synchrotron Radiation Laboratory. Data
were processed in DENZO and SCALEPACK (HKL2000) (36). Model building
was performed with the program Coot (37), and refinement was done with
REFMAC with TLS (CCP4i) (Table S1). In the structure of pol II complex with a
CTP incorporation against damaged guanosine residue, we also observed
additional weaker density within the second channel in comparison to the
nucleoside residue at the þ1 position, which may correspond to nonspecific
binding of a second CTP molecule through the soaking process. All structure
models in the figures were superimposed with nucleoside residues near the
active site using PYMOL (38).
Transcription Assay. Transcription assays were performed essentially as de-
scribed (11). In a typical reaction, 32P-labeled RNA oligonucleotide (10 pmol)
was annealed with template DNA 29mer (20 pmol, damaged or nondamaged
template) and nontemplate DNA 14mer (20 pmol) in elongation buffer
(20 mM Tris-HCl, pH 7.5, 40 mM KCl, 0.5 mM MgCl2) in a final volume of
20 μL. An aliquot of the annealed RNA/DNA hybrid was incubated with a five-
fold excess of pol II (final concentration of pol II 1.1 μM, of RNA oligonucleo-
tide 0.22 μM, and of DNA oligonucleotides 0.44 μM) for 10 min at room tem-
perature. Equal volumes of the NTP mixture solution were added (final
concentrations 25 μM) and the mixture was then incubated for 0, 0.5, 1,
2, 3, 4, 8, 16, 32, or 64 min at room temperature before addition of stop solu-
tion (final concentrations 5 M urea, 44.5 mM Tris-HCl, 44.5 mM boric acid,
26 mM EDTA, pH 8.0, Xylene Cyanol and Bromophenol Blue dyes). RNA pro-
ducts were analyzed by PAGE in the presence of urea. Visualization and
quantification of products were performed with the use of a PhosphorIma-
ger (Molecular Dynamics).
Computer
Modeling
Analysis.
Three
representative
platinum
units,
fPtðNH3Þ3g2þ, trans-fPtðNH3Þ2ðpyÞg2þ, and cis-fPtðNH3Þ2ðpyÞg2þ bound to
guanosine in DNA and positioned in either the −1 or þ1 site of pol II were
modeled to mimic the post- and pre-translocation states, respectively. The
vdW interaction energies between the three ligands at different orientations
and the rest of the pol II complex were systematically computed and taken as
direct indicators of steric effects.
The structure of the cis-fPtðNH3Þ2ðpyÞg2þ fragment on DNA in pol II is
available from the current study. Initial configurations for the other two
units, fPtðNH3Þ3g2þ and trans-fPtðNH3Þ2ðpyÞg2þ, were obtained by modeling.
Briefly, the same configuration of pol II, DNA, and RNA as found in the struc-
ture containing cis-fPtðNH3Þ2ðpyÞg2þ was used for these two complexes. The
geometry of the fPtðNH3Þ3g2þ moiety was taken from a previous structure
where it binds to a B-DNA dodecamer (PDB ID: 5BNA) (39). Docking was
achieved by aligning the damaged guanosine base of the two structures.
Finally, the trans-ammine group in fPtðNH3Þ3g2þ was replaced with a pyridine
ligand, and the Pt-N bond length was appropriately adjusted to obtain the
structure for trans-fPtðNH3Þ2ðpyÞg2þ. The same procedure was used to
generate structures at both þ1 and −1 positions.
The vdW energies were computed for different configurations generated
by rotating about the Pt-N7 bond from −180° to 180° for each platinum modi-
fication (see Figs. S5–S7). The rotation angle (φ) was defined to be positive
when rotating in the anticlockwise direction. In the configuration with
φ ¼ 0°, the plane composed of two Pt-N bonds of the ligand which are per-
pendicular to the Pt-N7 bond was set to be parallel to the damaged guano-
sine base. We noticed that, for fPtðNH3Þ3g2þ and trans-fPtðNH3Þ2ðpyÞg2þ, two
trans ammine groups were accommodated at slightly different configura-
tions, with φ ¼ 0° due to the different local environment, which leads to
slightly different energies between conformations with φ and φ 180°.
Because the purpose of our modeling study is to identify major steric clashes
instead of accurately computing free energy changes associated with rota-
tion of the ligand, which requires extensive conformational sampling, we
performed a simple average of the two energies (E1ðφÞ and E2ðφ 180°Þ)
based on their Boltzmann weights (T 298 K), eq 1,
¯E ¼ ðe−βE1E1 þ e−βE2E2Þ∕ðe−βE1 þ e−βE2Þ
[1]
to get a better estimate of vdW energy profiles.
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Wang et al.
The GROMACS simulation package was used to compute vdW energies
between the ligands and the pol II complex (40). A 20-Å cutoff was adopted
for computing the vdW interactions. The AMBER03 force field was used
for the pol II complex including protein, RNA, and DNA (41). The vdW force
field (Leonard–Jones potential) parameters for ligands were generated from
the AMTECHAMBER module of the AMBER 9 package (42) using the general
AMBER force field (GAFF) (43) developed for rational drug design. Since
the Pt atom is not in direct contact with the pol II complex and does not con-
tribute significantly to any steric effects, we excluded it from our vdW energy
calculations.
ACKNOWLEDGMENTS. This research was supported by the National Institute of
General Medical Sciences (NIH Pathway to Independence Award GM085136
to D.W. and GM49985 to R.D. Kornberg) and by the National Cancer Institute
(Grant CA034992 to S.J.L.). Portions of the research were carried out at the
Stanford Synchrotron Radiation Laboratory, a national user facility operated
by Stanford University on behalf of the U.S. Department of Energy, Office of
Basic Energy Sciences. The SSRL Structural Molecular Biology Program is sup-
ported by the Department of Energy, Office of Biological and Environmental
Research, and by the National Institutes of Health, National Center for
Research Resources, Biomedical Technology Program, and the National Insti-
tute of General Medical Sciences.
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|
3M4U
|
Crystal Structure of Trypanosoma brucei Protein Tyrosine Phosphatase TbPTP1
|
The Trypanosoma brucei Life Cycle Switch TbPTP1 Is
Structurally Conserved and Dephosphorylates the Nucleolar
Protein NOPP44/46*
Received for publication,January 28, 2010, and in revised form, April 19, 2010 Published, JBC Papers in Press,May 5, 2010, DOI 10.1074/jbc.M110.108860
Seemay Chou‡, Bryan C. Jensen§, Marilyn Parsons§¶, Tom Alber‡, and Christoph Grundner§¶1
From the ‡Department of Molecular and Cell Biology and QB3 Institute, University of California, Berkeley, California 94720-3200,
the §Seattle Biomedical Research Institute, Seattle, Washington 98109-5219, and the ¶Department of Global Health, University of
Washington, Seattle, Washington 98195-5065
Trypanosoma brucei adapts to changing environments as it
cycles through arrested and proliferating stages in the human
and tsetse fly hosts. Changes in protein tyrosine phosphoryla-
tion of several proteins, including NOPP44/46, accompany
T. brucei development. Moreover, inactivation of T. brucei pro-
tein-tyrosine phosphatase 1 (TbPTP1) triggers differentiation
of bloodstream stumpy forms into tsetse procyclic forms
through unknown downstream effects. Here, we link these
events by showing that NOPP44/46 is a major substrate of
TbPTP1. TbPTP1 substrate-trapping mutants selectively enrich
NOPP44/46 from procyclic stage cell lysates, and TbPTP1 effi-
ciently and selectively dephosphorylates NOPP44/46 in vitro.
To provide insights into the mechanism of NOPP44/46 recog-
nition, we determined the crystal structure of TbPTP1. The
TbPTP1 structure, the first of a kinetoplastid protein-tyrosine
phosphatase (PTP), emphasizes the conservation of the PTP
fold, extending to one of the most diverged eukaryotes. The
structure reveals surfaces that may mediate substrate specificity
and affords a template for the design of selective inhibitors to
interfere with T. brucei transmission.
Trypanosoma brucei causes human African trypanosomiasis
or African sleeping sickness, which is marked by debilitating
neurologic symptoms ranging from sensory impairment to the
characteristic aberrant sleeping patterns that progress to coma.
If untreated, human African trypanosomiasis is fatal. With
30,000 deaths a year and 60 million people living at risk (1),
human African trypanosomiasis is a major disease burden in
sub-Saharan Africa. Current drugs are ineffective and toxic,
and drug resistance is becoming a growing hurdle for treatment
(2).
T. brucei alternates between human and tsetse fly hosts,
requiring extensive and rapid physiologic adaptations. In
humans, the major T. brucei population consists of the extra-
cellular, proliferative slender form in the bloodstream, which
irreversibly differentiates into the G1-arrested stumpy form
poised for transmission to the tsetse fly. Taken up by the tsetse
fly, the stumpy form differentiates into the proliferative procy-
clic form in the insect midgut. Eventually, the tsetse salivary
gland becomes populated with metacyclic forms, which infect
the human host (3). This differentiation cycle requires survival
in a diverse set of environments and forms the basis for infec-
tivity and transmission.
The molecular signals, regulators, and effectors underlying
this complex sequence of events are not well understood but
could provide novel targets for therapeutic interference. Dis-
tinct patterns of protein tyrosine phosphorylation accompany
and often precede stage progression (4), suggesting that tyro-
sine phosphorylation is a key mechanism of developmental reg-
ulation. Studies of the T. brucei dual specificity kinases also pro-
vide evidence that tyrosine phosphorylation regulates the
trypanosome life cycle (5–7). Moreover, NOPP44/46,2 a nucle-
olar RNA-binding protein required for ribosome biogenesis (8),
exhibits dramatic changes in tyrosine phosphorylation in con-
cert with the T. brucei life cycle transitions (9). NOPP44/46 is
tyrosine-phosphorylated in both proliferating procyclic and
non-proliferating stumpy forms, but not in proliferating slen-
der forms, indicating a complex interplay between life cycle and
cell cycle in modulating tyrosine phosphorylation.
Recently, TbPTP1, a PTP with sequence similarity to classi-
cal human PTPs, was identified as a central molecular switch
for the stumpy-to-procyclic progression (10). TbPTP1 activity
arrests stumpy bloodstream forms, suggesting a model in which
TbPTP1 inactivation in the fly midgut releases the arrest and
triggers development into the procyclic form (10). Thus,
TbPTP1 might function downstream of the recently described
proteins associated with differentiation (PAD) transporters,
which represent the first known step in the pathway that allows
the differentiation signals citrate or cis-aconitate to trigger
developmental changes (11). However, the substrates and
downstream effects of TbPTP1 remain unknown.
By sequence comparison, TbPTP1 is similar to human clas-
sical PTPs such as the prototypical PTP1B. TbPTP1 has an
ortholog in Trypanosoma cruzi and Leishmania major and con-
tains six regions that appear to be specific to trypanosomatids
(10). The Kinetoplastida, including T. brucei, constitute some
* Thisworkwassupported,inwholeorinpart,byNationalInstitutesofHealth
Grant R01 AI31077 (to M. P.).
The atomic coordinates and structure factors (code 3M4U) have been deposited
in the Protein Data Bank, Research Collaboratory for Structural Bioinformat-
ics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).
1 To whom correspondence should be addressed: 307 Westlake Ave. N, Ste.
500, Seattle, WA 98109-5219. Tel.: 206-256-7295; Fax: 206-256-7229;
E-mail: christoph.grundner@sbri.org.
2 The abbreviations used are: NOPP44/46, nucleolar phosphoprotein 44/46;
TbPTP1, T. brucei protein-tyrosine phosphatase 1; PTP, protein-tyrosine
phosphatase; r.m.s., root mean square; CHES, 2-(cyclohexylamino)ethane-
sulfonic acid.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 29, pp. 22075–22081, July 16, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
JULY 16, 2010•VOLUME 285•NUMBER 29
JOURNAL OF BIOLOGICAL CHEMISTRY 22075
of the most diverged eukaryotes. The evolutionary distance and
unique sequences raise the question whether the overall struc-
tural conservation of the PTP fold is maintained in these distant
eukaryotes.
Here, we identify NOPP44/46 as a major substrate of the life
cycle switch TbPTP1. We also describe the TbPTP1 crystal
structure, revealing strong conformational similarity to other
eukaryotic PTPs and surface characteristics that help ratio-
nalize NOPP44/46 binding. Trypanosome-specific sequence
motifs follow the canonical PTP fold, and all major functional
elements are structurally conserved. These data establish the
structural correlates of kinetoplastid PTPs within the PTP fam-
ily and provide a new link in the signaling pathway controlling
the stumpy-to-procyclic transition.
EXPERIMENTAL PROCEDURES
Cloning, Protein Expression, and Purification—The full-
length TbPTP1 (systematic ID Tb10.70.0070) gene was ampli-
fied from genomic T. brucei DNA (kindly provided by Dr.
Christian Klotz) and cloned into the pET28b expression vector
in-frame with the N-terminal six-histidine tag. Point mutants
were generated according to the QuikChange protocol (Strat-
agene). BL21 (DE3)-CodonPlus cells were transformed, and
protein expression was induced at A600 of 0.6 by adding 100 M
isopropyl-1-thio--D-galactopyranoside. After 20 h of induc-
tion at 20 °C, cells were harvested, resuspended in 20 mM Tris,
pH 7.5, 100 mM NaCl, and lysed by sonication. The lysate was
centrifuged for 1 h at 20,000 g, and the supernatant was
loaded on a metal-chelating affinity column. Fractions contain-
ing TbPTP1 were identified by measuring the hydrolysis of
p-nitrophenyl phosphate (12). Fractions were pooled, loaded
on a gel filtration column, and eluted in 20 mM Tris, pH 7.5,
100 mM NaCl. Recombinant TbPTP1 was concentrated to
10 mg/ml.
In Vitro Dephosphorylation—NOPP44/46 (Genbank acces-
sion number HM44803) was amplified from T. brucei strain
29.13 (13) genomic DNA and cloned into pLEW-MHTAP (14)
for expression in procyclic form T. brucei 29.13. Expression of
the tagged protein was induced with tetracycline for 24 h, and
the protein was purified using a modified tandem affinity puri-
fication protocol with 1 mM sodium orthovanadate in the lysis
buffer (15, 16). The purified preparation was treated with 10 mM
dithiothreitol for 15 min to inactivate the sodium orthovanadate.
Dephosphorylation reactions were carried out at room tempera-
ture for 15 min in 20 mM Tris, pH 7.5, 100 mM NaCl buffer,
with varying amounts of TbPTP1. For dephosphorylation of
NOPP44/46 with other PTPs, PTP input was normalized to the
activity of 100 nM TbPTP1 at a saturating concentration of the
non-cognate substrate p-nitrophenyl phosphate. Reactions were
stopped by the addition of SDS-PAGE loading buffer, separated
by SDS-PAGE, and detected by Western blot using the 4G10 anti-
Tyr(P) antibody. Blots were stripped and reprobed with mono-
clonal anti-NOPP44/46 1D2 (9).
Substrate Trapping—TbPTP1 resin was prepared by cou-
pling TbPTP1 to NHS-activated SepharoseTM 4 fast flow (GE
Healthcare) according to the manufacturer’s protocol. Wild-
type or the D199A mutant TbPTP1 was coupled at a concen-
tration of 1 mg/ml followed by an incubation in 0.1 M Tris
blocking buffer. Lysates were prepared from procyclic form
T. brucei grown for 16 h in medium containing 1.5 M sodium
orthovanadate. Cells were extracted in lysis buffer (50 mM Tris-
HCl, pH 7.5, 150 mM NaCl, 2 mM EGTA, 1% Triton X-100)
containing 1 mM sodium orthovanadate, complete protease
inhibitor mixture (Roche Applied Science), and 5 mM iodoac-
etamide to inhibit endogenous PTP activity. Iodoacetamide
and orthovanadate were inactivated by the addition of 10 mM
dithiothreitol. To capture substrates of TbPTP1, 10 l of wild-
type or D199A TbPTP1 resin was incubated for 2 h at 4 °C with
500 l of lysate corresponding to 0.5 109 T. brucei cells. The
resin was washed five times in high salt buffer (20 mM HEPES,
pH 8.0, 2 M NaCl, 15% glycerol, 0.5% Nonidet P-40) followed by
five washes alternating in guanidine-HCl buffer (20 mM HEPES,
pH 8.0, 200 mM guanidine-HCl, 15% glycerol, 0.5% Nonidet
P-40) and low salt buffer (20 mM HEPES, pH 8.0, 300 mM NaCl,
15% glycerol). The resin was boiled in reducing SDS-PAGE
buffer for 15 min, run on a 12% Tris-glycine gel, and transferred
to nitrocellulose membrane for Western analysis with 4G10
anti-Tyr(P) (GE Healthcare) and anti-NOPP44/46 antibody (9).
To control for input of recombinant wild-type and trapping
TbPTP1, TbPTP1 was released from the column material by
boiling in SDS sample buffer prior to trapping and analyzed by
SDS-PAGE.
Crystallization, Structure Determination, and Structure
Analysis—Initial crystals were obtained by sitting drop vapor
diffusion trials at 18 °C from a 1:1 mixture of TbPTP1 at 10
mg/ml and 10% polyethylene glycol 3000, 100 mM CHES, pH
9.5. Diffraction quality crystals were obtained by hanging drop
vapor diffusion after introducing mutations E138A, E139A, and
E140A, predicted to reduce the surface entropy of TbPTP1 (17),
the addition of 10 mM tris(2-carboxyethyl)phosphine, and in-
drop trypsin cleavage of the His6 tag using a trypsin:TbPTP1
ratio of 1:1,000 (w/w). Crystals were immersed in mother liquor
containing 10% glycerol, mounted, and flash frozen in liquid
nitrogen.
Diffraction data were collected at the Lawrence Berkeley
National Laboratory Advanced Light Source Beamline 8.3.1.
Data were reduced using the HKL2000 program suite (18).
Phases were obtained by molecular replacement using MolRep
(19) and the search model PTPN3 (Protein Data Bank (PDB)
accession 2B49) modified by CHAINSAW (20). After auto-
mated model building in PHENIX (21), the final model was
built by alternating manual model building using Coot (22) and
maximum likelihood refinement using PHENIX. The Rfree was
determined using a random 5% of the data. The structure was
validated using MOLProbity (23). Images were generated in
PyMOL, and structure comparisons were performed using the
DALI server and PDBsum. The crystal structure was deposited
in the Protein Data Bank under accession number 3M4U.
RESULTS
TbPTP1 Binds NOPP44/46—To identify cellular substrates
of TbPTP1, we generated a substrate-trapping mutant by
replacing the general acid Asp199 with Ala. This mutant is cat-
alytically inactive but retains substrate binding, thus allowing
for stable trapping and isolation of substrates (24). Based on
previous studies suggesting that TbPTP1 is inactive and tyro-
TbPTP1 Structure and Substrate
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VOLUME 285•NUMBER 29•JULY 16, 2010
sine phosphorylation most pronounced in procyclic forms (4,
10), we used cell lysates from T. brucei procyclic forms for trap-
ping experiments. Tyrosine phosphorylation was preserved
throughout cell lysis by the addition of iodoacetamide and
sodium orthovanadate to inhibit endogenous PTPs. The wild-
type and D199A TbPTP1 variants were covalently linked to
NHS-Sepharose beads and incubated with T. brucei lysate.
After high salt, guanidinium hydrochloride, and detergent
washes, bound proteins were eluted by boiling in SDS-PAGE
loading buffer. Western blotting of eluates using anti-Tyr(P)
antibody showed the enrichment of several tyrosine-phosphor-
ylated proteins by the D199A mutant relative to the wild-type
TbPTP1 (Fig. 1A). Two major bands migrated at a molecular
mass of 45 and 70 kDa, and a minor band migrated at 40
kDa. The enrichment of these bands was selective as other
Tyr(P) proteins apparent in the total lysate were not trapped by
TbPTP1.
Because the molecular mass of the 45-kDa species corre-
sponded to that of NOPP44/46, a protein known to be tyrosine-
phosphorylated in procyclic forms in vivo (4, 9), we explored
the possibility that NOPP44/46 was a trapped substrate of
TbPTP1. Experimental replicates of trapping eluates were
probed with anti-NOPP44/46, resulting in signal at the position
identical to the band detected with anti-Tyr(P) antibodies (Fig.
1B). Some phospho-independent binding of NOPP44/46 to
wild-type TbPTP1 was also observed. The specific enrichment
of phosphorylated NOPP44/46 with the trapping TbPTP1
identifies NOPP44/46 as a potential in vivo substrate of
TbPTP1.
TbPTP1 Dephosphorylates NOPP44/46 in Vitro—To con-
firm the observed interaction between TbPTP1 and NOPP44/
46, we tested dephosphorylation of NOPP44/46 by TbPTP1 in
vitro. Although the pH optimum of TbPTP1 is 6 (10), this pref-
erence is unlikely to reflect physiologic function rather than the
generally higher nucleophilicity of cysteine residues at low pH.
In vitro dephosphorylation assays were therefore performed at
pH 7.5, more similar to the pH at which TbPTP1 likely func-
tions. Phosphorylated NOPP44/46 was obtained by overex-
pressing a C-terminally TAP-tagged version of the full-length
protein in procyclic forms. Rapid dephosphorylation of
NOPP44/46 was observed at the lowest TbPTP1 concentration
tested (3.7 nM) and was complete at TbPTP1 concentrations at
or above 33 nM (Fig. 2A). To test whether dephosphorylation of
NOPP44/46 by TbPTP1 is specific, we tested the activity of a
panel of unrelated microbial PTPs on NOPP44/46. Of the five
tested PTPs, only TbPTP1 and Yersinia YopH dephosphory-
lated NOPP44/46; no activity was observed using Mycobacte-
rium tuberculosis PtpA and PtpB, Staphylococcus aureus
SaPtpA, or Listeria monocytogenes lmo1935 (Fig. 2B). With
YopH known to have broad substrate specificity (25), these data
show complete, efficient, and selective dephosphorylation of
NOPP44/46 by TbPTP1 in vitro.
NOPP44/46 Is Phosphorylated on Tyr181—To define the
phosphorylation site(s) on NOPP44/46 recognized by TbPTP1,
we assessed the phosphorylation state of NOPP44/46 variants
in which each of the five Tyr residues was replaced individually
with Phe. Y181F completely abrogated tyrosine phosphoryla-
tion of NOPP44/46, as shown by anti-Tyr(P) Western analysis
(Fig. 3). None of the other mutations reduced the level of
NOPP44/46 Tyr phosphorylation, indicating that Tyr181 is the
only phosphorylated Tyr and the target of TbPTP1. The
NOPP44/46 phosphorylation site is located in a 40-residue
acidic loop encompassing residues 167–207. The sequence of
this segment, 167DAGDEDDNDDDDEAYDEDDSDDDDDD-
DDDDDDDDDDDDDDE207, indicates that phosphorylation
adds additional negative charges to a nearly uninterrupted
acidic sequence. This acidic region containing the target Tyr is
found only in T. brucei homologs.
TbPTP1 Has a Classical PTP Fold—To explore the basis for
recognition of the unusual substrate target sequence and the
architecture of this diverged kinetoplastid PTP, we determined
the crystal structure of TbPTP1 at 2.4 Å resolution (Table 1).
The asymmetric unit contains two TbPTP1 molecules with a
root mean square (r.m.s.) deviation of all atoms of 0.3 Å. The
structure comprises residues Ser6–Thr301 in chain A and
residues Met1–Leu298 in chain B (Fig. 4), as well as one phos-
phate per TbPTP1 and 192 water molecules. No clear elec-
tron density was visible for residues Leu66–Gln73 and
Ala160–Ala162 of chain A and residues Lys67–Arg75, Ala140,
and Gln147–His151 of chain B. The protein contains four
changes. The catalytic cysteine is changed to alanine, possibly
through desulfurization by the phosphine tris(2-carboxyethyl)-
FIGURE 1. Substrate trapping identifies NOPP44/46 as a major TbPTP1
substrate. A, anti-Tyr(P) (-pTyr) Western blot after TbPTP1 substrate trap-
ping. The trapping mutant (trap) selectively enriches three major tyrosine-
phosphorylated proteins. WT, wild type. B, anti-NOPP44/46 Western blot of
proteins bound to wild type and trapping mutant. The Western blot in B is an
experimental replicate of A and identifies the 45-kDa band as NOPP44/46.
Some phospho-independent binding is also apparent. C, Coomassie Blue
staining showing equivalent amounts of TbPTP1 wild type and mutant elute
from the resin prior to trapping.
TbPTP1 Structure and Substrate
JULY 16, 2010•VOLUME 285•NUMBER 29
JOURNAL OF BIOLOGICAL CHEMISTRY 22077
phosphine present at high concentrations in the protein drop, and
Glu138-140 were mutated to alanine to improve the crystallization
properties of the protein (17).
The overall fold of TbPTP1 resembles that of other classical
PTPs, with an extended, twisted -sheet at the center and
-helices surrounding it (Fig. 4A). The catalytic loop, or P-loop,
is situated at the center of the active site and comprises the
invariant PTP signature motif Cys-Xaa5-Arg. A phosphate
binds in the position similar to that of Tyr(P) substrate phos-
phate in PTP-peptide substrate structures (26) (Fig. 4B). The
active site cavity is further delineated by the Tyr(P) loop that
deepens the cavity to 9 Å, thus
excluding Ser(P) and Thr(P) resi-
dues. The WPD loop containing the
general
acid
Asp199
assumes
a
closed conformation, similar to that
seen in other PTP structures with
small ligands bound (27). TbPTP1
contains 9 of 10 PTP sequence
motifs (28) in the same spatial
organization as human PTPs. The
six trypanosome-specific sequence
motifs of TbPTP1 follow the canon-
ical PTP fold and do not give rise
to new structural features. Con-
sistent with a role in substrate
recognition
or
regulation,
the
trypanosome-specific
sequences
predominantly map to the surface
of TbPTP1 (Fig. 5A). The phos-
phate engages in the typical inter-
actions with the P-loop, hydrogen-
bonding
with
six
main
chain
amides and the invariant Arg235 side chain.
The closest structural homologs of TbPTP1 found by the
DALI server are the prototypical human PTP1B and PTPRO
(Glepp1), with a C r.m.s. deviation of 2.1 and 2.2 Å, respec-
tively. The superposition of TbPTP1 with PTPRO (PDB ID
2G59) shows the overall large similarity, with major differences
only at the termini and surface loops (Fig. 5B). The TbPTP1
loop from 62 to 79, although mostly invisible in the structure,
has shifted at the base when compared with the equivalent
PTPRO loop and contains a single-residue insertion when com-
pared with PTPRO and up to seven residues when compared
with other human PTPs. The TbPTP1 loop 138–154 contain-
ing a PEST sequence shows the most divergence from the
PTPRO structure.
TbPTP1 forms weak interactions between the two molecules
in the asymmetric unit in the crystal (data not shown). The
interactions comprise two symmetric salt bridges between
Lys123 and Glu201, hydrogen bonds between Gly127 and Glu201,
as well as 44 non-bonded interactions resulting in an interface
of 500 Å2. PTPs such as PTP form dimers in the crystal and
in solution (27). However, TbPTP1 migrates as a monomer in
size exclusion chromatography (data not shown), suggesting
that these contacts do not reflect a physiologic state but are a
result of crystal packing.
Although the three-dimensional organization of the PTP
active site is highly similar in all PTPs across families, PTP
surface properties vary widely and produce large diversity
(27). The TbPTP1 electrostatic surface shows distinct and
continuous electronegative and positive areas (Fig. 6). The
active site shows moderately electropositive potential, with
the closed WPD loop burying additional electropositive
regions of the phosphate binding pocket. A continuous elec-
tropositive stretch runs across the active site and along one
side of the molecule. This stretch includes the side chains of
Arg15, -23, -30, -50, -125, -175, -276, and Lys113 and -131 (Fig. 6A).
The electrostatic surface of PTP1B with a closed WPD loop
FIGURE 2. TbPTP1 efficiently and selectively dephosphorylates NOPP44/46 in vitro. A TAP-tagged allele of
NOPP44/46 was expressed in procyclic forms and affinity-purified for dephosphorylation reactions. A, TbPTP1
dephosphorylates NOPP44/46 in a dose-dependent manner. -pTyr, anti-Tyr(P) antibody. B, TbPTP1, but not
unrelated phosphatases from M. tuberculosis (PtpA and PtpB), S. aureus (SaPtpA), and L. monocytogenes
(lmo1935), dephosphorylates NOPP44/46.
FIGURE 3. NOPP44/46 is phosphorylated on Tyr181. A, schematic of
NOPP44/46 indicating domain organization and position of the five tyrosine
residues. U, unique region; J, junction; A, acidic region; R, RGG repeat region.
B, all five NOPP44/46 tyrosines were individually changed to Phe, and the
phosphorylation of NOPP44/46 was detected by anti-Tyr(P) antibody (-pTyr)
(upper panel) and anti-NOPP44/46 control Western (lower panel). WT, wild
type.
TbPTP1 Structure and Substrate
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VOLUME 285•NUMBER 29•JULY 16, 2010
also shows extended electropositive areas (Fig. 6B). These
surfaces likely complement the sequence of the highly elec-
tronegative PTP1B substrate, the insulin receptor. PTP1B
binds the triply phosphorylated peptide sequence pYETD-
pYpY, which also concentrates a large number of charges
around the PTP1B active site. In contrast, PTPRO shows few
electropositive areas outside of the direct active site vicinity
and a predicted second Tyr(P) binding site (Fig. 6B, right
panel).
DISCUSSION
T. brucei requires stringent control of developmental pro-
grams to successfully infect its human and tsetse fly hosts,
and a molecular hallmark of life cycle transitions in T. brucei
is the coordinated change in tyrosine phosphorylation.
Recently, the tyrosine phosphatase TbPTP1 was identified as
a key regulator of the trypanosome life cycle. TbPTP1 has an
ortholog in T. cruzi with 61.3% sequence identity. Although
an intracellular parasite, T. cruzi shares the bloodstream-to-
insect route of transmission controlled by TbPTP1 in T. bru-
cei, suggesting that the function of the two PTPs may be
conserved.
Despite the evolutionary distance between humans and
trypanosomes, our crystal structure of TbPTP1 shows a high
degree of structural conservation of the conventional PTP
fold. The 24% sequence identity of TbPTP1 to human PTP1B
translates into a C r.m.s. deviation of 2.1 Å. The conser-
vation of the PTP fold thus extends not only to bacteria but
also to distant eukaryotes and underscores the utility and
evolutionarysuccessofthisscaffoldfortyrosinedephosphory-
lation. TbPTP1 shares 9 of 10 signature motifs with the
human PTPs and has six additional trypanosome-specific
motifs that may play roles in func-
tional regulation or substrate rec-
ognition. The folding of these
motifs suggests that although not
giving rise to new structural ele-
ments, their position mostly on
the surface is consistent with a role
in substrate recognition and/or
regulation.
The TbPTP1 structure also pro-
vides the basis for inhibitor design.
The strong similarity to human
PTPs highlights the need for struc-
tural
information
to
guide
the
design of selective TbPTP1 inhibi-
tors as tools and potential therapeu-
tics. TbPTP1 prevents premature
differentiation of stumpy blood-
stream forms to procyclic forms,
which lack immune evasion mecha-
nisms that allow survival in the
mammalian host. Thus, inhibition
of TbPTP1 would reduce the pool of
tsetse-infective parasites within the
mammalian host, potentially atten-
uating transmission, an approach
that has gained acceptance for the
reduction of malaria (29). Blocking
transmission could be particularly
advantageous for controlling animal
trypanosomiasis, which affects live-
stock and remains a major hurdle
to economic development in sub-
FIGURE 4. Overall structure of TbPTP1. A, TbPTP1 shares the canonical PTP fold. The catalytic motifs P-loop
and WPD loop are highlighted in orange and yellow, respectively. No electron density for residues 65–74 was
visible in chain A (dotted line). N-term, N terminus; C-term, C terminus. B, 2Fo Fc electron density map of the
active site showing phosphate (center), contoured at 1.0 .
FIGURE 5. TbPTP1 has a similar fold to human PTPs. A, trypanosome-specific sequence motifs (green) map on
the surface outside of the active site (P-loop in orange). B, superposition of the C chain in ribbon representation
showing overall strong similarity to human PTPRO.
TABLE 1
Data collection and refinement statistics for TbPTP1
Parentheses denote values for the highest resolution shell.
Data collection
Crystal symmetry
P212121
Unit cell
a, b, c (Å)
76.63, 77.38, 117,2
, , (°)
90, 90, 90
Resolution (Å)
2.4
Rmerge (%)
10
Completeness ( %)
99.46 (97)
Multiplicity
5 (4.9)
I/I
54 (2.5)
Refinement statistics
Resolution (Å)
47-2.4
Reflections
28,040
Rwork/Rfree (%)
20/26
r.m.s. bonds (Å)
0.008
r.m.s. angles (°)
1.065
Average B-factor (Å2)
42.4
Main chain dihedral angles
Most favored (%)
96.6
Allowed (%)
3.2
TbPTP1 Structure and Substrate
JULY 16, 2010•VOLUME 285•NUMBER 29
JOURNAL OF BIOLOGICAL CHEMISTRY 22079
Saharan Africa. Moreover, T. brucei rhodesiense infects both
humans and animals, providing a parasite reservoir for human
infection.
Although tyrosine phosphorylation is emerging as a key reg-
ulator of the trypanosome life cycle, little is known about the
molecular pathways that lead to downstream developmental
changes. Identification of TbPTP1 substrates is essential to
understanding the mechanisms by which TbPTP1 regulates
T. brucei differentiation. Among the phosphoproteins selec-
tively enriched using a TbPTP1 trapping mutant, we identified
NOPP44/46 and yet unidentified 70- and 40-kDa phospho-
proteins as substrates of this PTP. Other substrate phosphopro-
teins might be associated with the insoluble fraction and not
detected by our methods. The functional interaction of
TbPTP1 and NOPP44/46 is supported by efficient and selective
in vitro dephosphorylation. Furthermore, the phosphorylation
pattern of NOPP44/46 during developmental stages, unlike
that of other major tyrosine-phosphorylated species, matches
the proposed activity profile of TbPTP1 in slender and procy-
clic forms (4, 9). The presence of phosphorylated NOPP44/46
in stumpy forms may reflect decreasing TbPTP1 activity or
changes in the activity of the cognate kinase(s) in combination
with a large increase in NOPP44/46 protein levels observed in
stumpy forms (9).
The
TbPTP1
crystal
structure
allows
rationalizing
NOPP44/46 substrate binding. The distinct, continuous elec-
tropositive area running across the TbPTP1 surface and the
active site might be a footprint of electronegative regions of
its substrate(s), such as the acidic stretch harboring the
NOPP44/46 Tyr(P). Phosphorylation sites are usually found in
flexible loop regions, suggesting that linear sequences rather
than conformational sites serve as dephosphorylation sub-
strates. This is consistent with the NOPP44/46 dephosphory-
lation site, which is predicted to be highly disordered. More-
over, the kinetics of substrate peptide turnover by PTPs are
often approaching the limits of diffusion, suggesting that the
selectivity and binding determinants of peptides are contained
within the primary peptide sequence (30).
A phosphoproteomic study of tyrosine-phosphorylated pro-
teins in T. brucei procyclic forms identified 34 phosphoproteins
(31). NOPP44/46 Tyr181, however, was not identified, likely due
to experimental limitations of that study. By immunofluores-
cence, phosphotyrosine proteins mostly associated with the
cytoskeleton and the nucleolus, which is also the site of
NOPP44/46 (32). A physiological interaction between TbPTP1
and NOPP44/46 would require cellular co-localization, and
consistent with this tenet, TbPTP1 was found to associate with
the cytoskeletal and nuclear fractions (10). However, it remains
possible that NOPP44/46 is dephosphorylated outside of the
nucleus as a recent study suggests that a pool of NOPP44/46 is
exported out of the nucleus via exportin 1 (33).
TbPTP1 is a molecular switch for the stumpy-to-procyclic
transition as both genetic and pharmacological inhibition of the
phosphatase lead to spontaneous differentiation of committed
stumpy forms to procyclic forms in vitro (10). Because trypano-
somatids do not use transcriptional control to regulate ex-
pression of protein-coding genes (with a few exceptions), the
key substrates of TbPTP1 that mediate this effect are likely to
modulate mRNA stability, translation, or protein turnover. The
identification of a known ribosome biogenesis protein,
NOPP44/46 (8), as a potential in vivo substrate of TbPTP1
points toward a possible role in translational control. For exam-
ple, the tyrosine phosphorylation state of NOPP44/46 may
modulate ribosome biogenesis, which would in turn affect
translational capacity. This possibility will be examined in
future studies. Alternatively, NOPP44/46 may have uncharac-
terized cellular functions in addition to its essential role in ribo-
some biogenesis that could play specific roles in differentiation.
In vivo, NOPP44/46 acts as a structural scaffold for several
nucleolar proteins (34, 35), and its phosphorylation state may
modulate these interactions to promote specific cellular
changes. Studies in other organisms suggest the existence of
functional links between proteins involved in ribosome biogen-
esis and development, highlighting complex yet conserved
modes of developmental regulation. In zebrafish, Pescadillo, an
essential gene required for nucleolar assembly and 60 S biogen-
esis, was originally discovered in a screen for regulators of
embryonic development (36–39). In yeast, the Pescadillo
ortholog, Yph1p, is found in two distinct multiprotein com-
plexes with different functions in ribosome biogenesis and
DNA replication (40). Interestingly, depletion of NOPP44/46
(44) and Yph1p both lead to cell cycle arrest with defective
S-phase progression (40, 41). Furthermore, the ribosomal bio-
genesis factors such as nucleolin and nucleophosmin play roles
in the cytosol or nucleus distinct from their function in ribo-
some biogenesis (42, 43). As other substrates of TbPTP1 are
identified and tools for the study of TbPTP1 in vivo are refined,
we will be better able to determine how these processes act
together or apart to influence trypanosomatid differentiation
and potentially provide insight into a novel signaling mecha-
nism conserved in eukaryotic development.
FIGURE 6. Electrostatic surface potential of TbPTP1. A, left, the TbPTP1 sur-
face shows distinct electronegative (red) and positive (blue) regions, with a
continuous electropositive stretch across the active site. The entry to the
active site is indicated by the circle. Right, schematic representation in the
sameorientationasleftpanel.B,electrostaticsurfacerepresentationofPTP1B
(1SUG, left) and PTPRO (2G59, right) in the same orientation as TbPTP1. PTP1B
also binds a highly negatively charged substrate and shows large electropos-
itive areas.
TbPTP1 Structure and Substrate
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VOLUME 285•NUMBER 29•JULY 16, 2010
Acknowledgments—We thank Christine L. Gee for help with model
building and refinement, the staff at the Advanced Light Source
Beamline 8.3.1 for help with data collection, and Carolina Vega and
Charles Kifer for technical assistance.
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e678
TbPTP1 Structure and Substrate
JULY 16, 2010•VOLUME 285•NUMBER 29
JOURNAL OF BIOLOGICAL CHEMISTRY 22081
|
3M4W
|
Structural basis for the negative regulation of bacterial stress response by RseB
|
PROTEIN STRUCTURE REPORT
Structural basis for the negative regulation
of bacterial stress response by RseB
Dong Young Kim, Eunju Kwon, JongKeun Choi, Hye-Yeon Hwang,
and Kyeong Kyu Kim*
Department of Molecular Cell Biology, Sungkyunkwan University School of Medicine, Suwon 440-746, Korea
Received 29 December 2009; Revised 13 March 2010; Accepted 16 March 2010
DOI: 10.1002/pro.393
Published online 29 March 2010 proteinscience.org
Abstract: The rE-dependent stress response in bacterial cells is initiated by the DegS- and
RseP-regulated intramembrane proteolysis of a membrane-spanning antisigma factor, RseA. RseB
binds to RseA and inhibits its sequential cleavage, thereby functioning as a negative modulator of
this response. In the crystal structure of the periplasmic domain of RseA bound to RseB, the DegS
cleavage site of RseA is unstructured, however, its P1 residue is buried in the hydrophobic pocket
of RseB, which suggests that RseB binding blocks the access of DegS to the cleavage site.
Keywords: RseA; RseB; RseP; stress response; sigma factor; crystal
Introduction
Regulated intramembrane proteolysis (RIP) is a con-
trol mechanism underlying transmembrane signal
transfer and performs a key role in the initiation of
the
essential
signal
transduction
pathways
in
diverse organisms.1 For example, the Notch signal-
ing pathway, which is critical for a variety of cell–
cell
communications
in
multicellular
organisms,
is controlled by the RIP of the Notch receptor
by
ADAM-family
metalloprotease
and
gamma-
secretase.1 In Gram-negative bacteria, the sequen-
tial cleavage of RseA, a membrane-spanning anti-rE
factor, modulates the initiation of the envelope-stress
response.2,3 RseA forms a tight complex with rE
using its N-terminal cytoplasmic domain, thereby
inhibiting the transcription of rE-dependent genes.
Under stress conditions that include the misfolding
of periplasmic proteins, two membrane proteases,
DegS and RseP, sequentially degrade RseA to liber-
ate rE (Supporting Information Fig. S1). DegS,
which is activated when its PDZ domain is bound to
the C-terminal peptide of unfolded outer membrane
porins (OMPs), cleaves the C-terminal periplasmic
domain
of
RseA.4
Subsequently,
RseP
cleavage
within the membrane domain of RseA releases the
cytoplasmic domain of RseA (associated with the rE)
from the membrane. In the final step, the cyto-
plasmic domain of RseA is degraded such that the
released rE can interact with RNA polymerase.5
RseB
also
participates
in
the
regulation
of
rE-dependent envelope-stress response by inhibiting
Additional Supporting Information may be found in the online
version of this article.
Dong
Young
Kim’s
current
address
is
Department
of
Pharmaceutical
Chemistry,
University
of
California,
San
Francisco, 600 16th Street, San Francisco, CA 94107, USA.
Grant sponsor: 21C Frontier Functional Proteomics Program;
Grant number: FPR08B2-270; Grant sponsor: Korea Healthcare
technology
R&D
Project;
Grant
number:
A092006;
Grant
sponsor: Ubiquitome Research Program; Grant number: M105
33010001-05N3301-00100; Grant sponsor: National Research
Laboratory Program; Grant number: NRL-2006-02287.
*Correspondence to: Kyeong Kyu Kim, Department of Molecular
Cell Biology, Sungkyunkwan University School of Medicine,
Suwon 440-746, Korea. E-mail: kkim@med.skku.ac.kr
1258
PROTEIN SCIENCE 2010 VOL 19:1258—1263
Published by Wiley-Blackwell. V
C 2010 The Protein Society
the intramembrane proteolysis of RseA.6 RseB has
been previously demonstrated to suppress the pro-
teolytic activity of DegS for RseA,6,7 independently
of the activation mechanism of DegS.8 Accordingly,
either the deletion of the rseB gene or the release of
RseB from RseA results in a more rapid degradation
of RseA and increased activity of rE.6,9 In this
study, we attempted to characterize the regulatory
role of RseB in the proteolytic cleavage of RseA and
determined the crystal structure of RseB in complex
with RseAperi (the periplasmic domain of RseA, resi-
dues 121–216) at a resolution of 2.3 A˚ by molecular
replacement using apo-RseB (PDB ID: 2P4B) as a
template (Table I).
Results and Discussion
Structure determination
The RseAperiRseB structure was determined at a
resolution of 2.3 A˚ by molecular replacement using
the large domain (residues 26–200) of E.coli RseB
(PDB ID: 2P4B) as a template.10 Although the asym-
metric unit of the crystal harbors four RseAperiRseB
complexes (Supporting Information Fig. S2), the di-
meric structure of the complex has been known from
size exclusion and SAXS (Small Angle X-ray Scatter-
ing) data.11 Each complex (Com1–Com 4 in Support-
ing Information Fig. S2) is composed of one RseB
monomer and one RseAperi monomer. Com1 is in
contact with two other complexes, Com2 and Com3.
The Com1:Com2 interaction, which is mediated by
hydrogen bonds between relatively well-conserved
residues, buries the 1190 A˚ 2 surface area of each
complex. RseB:RseB, RseAperi:RseAperi, and RseAper-
i:RseB interfaces contribute to this burial of surface
area by 889 A˚ 2, 57 A˚ 2, and 244 A˚ 2, respectively. The
Com1:Com3 interaction results in the burial of 370
A˚ 2, which is primarily a RseB:RseB contact and
involves a zinc ion that was added for the purposes
of crystallization (Supporting Information Fig. S3).
Moreover, the dimeric interaction between the N-ter-
minal regions of the two RseAs in the Com1:Com2
dimer and their proximity to the transmembrane
region demonstrate the involvement of the trans-
membrane
domain
of
RseA
in
dimeric
contacts
[Fig.
1(a)
and
Supporting
Information
Fig.
S4].
There is no biologically relevant higher-order oligo-
meric form that can be generated by symmetry
operations.
Accordingly,
the
Com1:Com2
dimer
(or Com3:Com4 dimer) was considered biologically
relevant
[Fig.
1(a)
and
Supporting
Information
Fig. S2]. In this manuscript, we used Com1 and
Com1:Com2 to describe the monomeric and dimeric
RseAperiRseB complexes, respectively. The RMSD
between
Com1:Com2
and
Com3:Com4
complexes
was 0.43 A˚ for 626 Ca atoms.
Overall structure
We were able to model most residues in RseB, with
the exception of the first N-terminal residue, resi-
dues 240–246, and three C-terminal residues. The
loop connecting the large (RseB25–209) and small
(RseB217–315) domains was disordered in the apo-
RseB model10,12; however, it was well ordered in the
RseAperiRseB complex due to the interaction with
the bound RseAperi, which results in the stabilization
of the loop (Fig. 1). By way of contrast, the 96-resi-
due periplasmic domain of RseA (residues 121–216)
was largely unstructured, and only two regions
involved in RseB binding, RseA132–151 and RseA169–
190, were modeled (Fig. 1 and Supporting Informa-
tion Fig. S5). RseB evidenced similar structures in
their apo- (Chain A of 2P4B or Chain A of 2V43)
and RseAperi-bound states, with an RMSD of 0.85 A˚
for 276 Ca atoms (2P4B) or 1.61 A˚ for 261 Ca atoms
(2V43), thereby indicating that its overall conforma-
tion is largely maintained upon RseA binding. The
major local conformational change was found in two
b-strands (b5 and b6; residues 88–104) in the small
domain of RseB (2P4B), which binds directly to the
C-terminus of RseA132–151 [Fig. 1 and Supporting
Information Fig. S6(a)]. The conformational changes
are more drastic when the RseA-bound RseB was
compared with another crystal structure of apo-
RseB (PDB ID: 2V43) where four b-strands (b3–b6;
residues
68–104)
exhibit
large
conformational
Table I. Data Collection and Refinement Statistics
RseAperiRseB
Data collection
Space group
P212121
Cell dimensions
a, b, c (A˚ )
87.05, 119.58, 150.67
a, b, c ()
90.00, 90.00, 90.00
Resolution (A˚ )
30.00–2.30 (2.38–2.30)a
Rsym or Rmerge
6.0 (20.8)
I/rI
20.0 (3.5)
Completeness (%)
93.0 (81.5)
Redundancy
4.6
Refinement
Resolution (A˚ )
20.00–2.30
No. reflections, working/free
62725/3320
Rwork/Rfree
23.9/27.1
No. atoms
Protein
10108
Zn2þ
6
Water
421
B-factors
Protein
52.1
Zn
82.3
Water
50.0
R.m.s. deviations
Bond lengths (A˚ )
0.008
Bond angles ()
1.482
Ramachandran plot
Most favored (%)
87.2
Additionally allowed (%)
12.7
Generously allowed (%)
0.2
Disallowed (%)
0.0
a Values in parentheses are for highest-resolution shell.
Kim et al.
PROTEIN SCIENCE VOL 19:1258—1263
1259
changes after binding to the C-terminus of RseA
[Fig. 1 and Supporting Information Fig. S6(b)].
Interaction between RseAperi and RseB
RseA binds to a broad area of the RseB groove that
is formed between the large and small domains of
RseB (Figs. 1 and 2). RseA132–151 mostly forms a
random coil rather than a regular secondary struc-
ture and interacts with residues in the large domain
of RseB. The residues in RseA132–151 form hydropho-
bic interactions with the hydrophobic residues or ali-
phatic carbons of bulky residues of RseB, with the
exception of Lys144, which forms a salt-bridge with
Glu181 of RseB [Fig. 2(a)]. The results of the histi-
dine pull-down assay verified that an RseAperi mu-
tant featuring Ala substitutions at Gly143, Lys144,
and Pro147 was still capable of binding to RseB
(data not shown), thereby indicating that the electro-
static interaction attributable to Lys144 is not crit-
ically important to the association between RseB
and
RseA132–151.
Consistent
with
this
finding,
Lys144 is not conserved among RseA homologues in
Gram-negative bacteria (Fig. 3).
RseA169–190 exhibits a helical conformation and
binds principally to the small domain of RseB. The
charged residues in RseA169–190 are well-conserved
in the RseA homologues and are important to RseB
binding
(Figs.
2
and
3).
Most
notably,
Arg172,
Asp179, Glu181, and Arg184/Arg185 in RseA169–190
form salt bridges with Glu293, Arg239, Arg282, and
Asp109 of RseB, respectively [Figs. 2(a,b)]. It has
been demonstrated that RseA169–185 is the minimum
fragment necessary for RseB binding8,10; addition-
ally, the mutation of the conserved Arg residues in
this fragment (Arg172, Arg184, and Arg185) abol-
ishes RseB binding activity10,13 (Fig. 3). Therefore,
RseA132–151 does not appear to be the primary deter-
minant in RseB binding, but it may perform other
additional functions, such as recruiting RseB or
sterically inhibiting the access of proteases. Two
RseAperiRseB complexes in a dimer (Com1:Com2)
are
also
stabilized
via
intercomplex
interactions
[Fig. 1(a) and Supporting Information Fig. S2]. The
Val135, Phe136, and Thr138 residues of RseA in
Com1
are
in
contact
with
Ile50,
Asn51/Thr179/
Gln182,
and
Arg169/Arg184
of
RseB
in
Com2,
respectively, and vice versa [Figs. 1(a),2(b)].
Structural implication of the binding of
RseA to RseB
The DegS cleavage site (Val148-Ser149; P1-P10) at
the C-terminal end of RseA132–151, is located within
the RseB groove in the RseAperiRseB complex [Figs.
1,2(c)]. Val148 is buried in the hydrophobic pocket
formed by Phe100 and Leu102 of RseB and Leu182
of RseA169–190. Ser149, which is located near the he-
lix in RseA169–190, forms a hydrogen bond with
Gln178 of RseA. As a result, the DegS cleavage site
is almost completely hidden by RseB and RseA169–
190, such that DegS access is restricted in the RseA-
periRseB complex, and probably also in the RseAR-
seB complex (Fig. 1). From this perspective, it has
been theorized that the binding of RseA132–151 to
RseB contributes to locating the cleavage site deep
inside of the RseB groove, thereby rendering it re-
sistant to DegS cleavage. This mechanism is consist-
ent with the model proposed in Ref. 8.
In the proteolytic cascade of RseA, the cleavage
by RseP requires prior periplasmic cleavage by DegS
and the release of RseA149–216.
5 It was reported
Figure 1. Structure of the RseAperiRseB complex. (a) Dimer model of the RseAperiRseB complex. Each RseAperi is depicted
in magenta or yellow, and RseB is depicted in green or slate. Ribbon diagram (b) and surface model (c) of the monomeric
RseAperiRseB complex. The regions important for their binding, RseB25–209, RseB210–216, RseB217–315, RseA132–151, and
RseA169–190, are colored slate, purple, red, green, and yellow, respectively. The DegS cleavage site is shown in blue.
1260
PROTEINSCIENCE.ORG
Crystal Structure of RseB in Complex with RseA
recently that the interaction of the newly exposed
C-terminal residue of RseA1–148, Val 148, with the
second PDZ domain of RseP is critically important
for the cleavage.14 It is expected that the interaction
between RseB and RseA1–148 is not very strong due
to the lower binding affinity of RseA121–173 for
RseB detected in previous biochemical studies.10
These findings suggest that RseB in complex with
RseA149–216
will
dissociate
from
RseA1–148
after
DegS
cleavage
and
that
RseB
is
unlikely
to
Figure 2. RseAperiRseB interaction. (a) Schematic drawing of RseAperiRseB interaction. RseA residues located in random
coils and helices are shown as orange circles and green pentagons, respectively. RseB residues in the same complex and
from the other RseAperiRseB complex are marked as white and cyan boxes, respectively. Charge interactions, hydrogen
bonds, and hydrophobic contacts are shown as red, blue, and black lines, respectively. (b) Binding interface between RseA
and RseB. RseB, RseA132–151, and RseA169–190 are colored purple, green, and yellow, respectively. Dotted lines indicate
charge interactions and the involved residues are depicted by stick models. (c) The DegS cleavage site (Val148-Ser149)
bound to the RseB groove in the RseAperiRseB complex is drawn in a ribbon model with the same color scheme as in
Fig. 1(b). The residues near the cleavage site are drawn as stick models and labeled. The cleavage site is indicated by a
black arrow and labeled.
Figure 3. Multiple sequence alignment of the periplasmic domain of RseAs from Gram-negative bacteria. Identical and similar
residues are boxed in blue and yellow, respectively. Species abbreviations are as follows: Ec, Escherichia coli; Sf, Shigella
flexneri; Se, Salmonella enterica; Yp, Yersinia pestis; Eca, Erwinia carotovora; Vc, Vibrio cholerae; So, Shewanella oneidensis;
Hi, Haemophilus influenzae; Ms, Mannheimia succiniciproducens.
Kim et al.
PROTEIN SCIENCE VOL 19:1258—1263
1261
reassociate with RseA1–148. Therefore, RseP
will
interact with the C-terminal end of the DegS-cleaved
RseA (RseA1–148).
Materials and Methods
Protein expression and purification
E.coli RseAperi (periplasmic domain containing resi-
dues 121–216) and RseB (residues 24–318) were
expressed separately in E. coli BL21(DE3) as previ-
ously
described.10,11
His-Trx-RseAperi-
and
RseB-
expressing cells were harvested and mixed at a ratio
of 1:3 (wet weight) to ensure the formation of the
complex. The cells were then sonicated in buffer
A (20 mM Tris-HCl pH 7.5 and 0.1M NaCl). The
RseAperiRseB complex was then purified by nickel-
affinity chromatography and size exclusion chroma-
tography. The cleared lysates were loaded onto a
metal-chelating column (GE Healthcare, Princeton,
NJ) and the proteins were eluted with 50–500 mM
imidazole gradient. The fractions containing His-
Trx-RseAperiRseB were pooled and dialyzed twice
against buffer A. The His-Trx tag was removed with
thrombin at room temperature, and RseAperiRseB
was purified further using a Superdex-200 column
(GE
Healthcare,
Princeton,
NJ)
pre-equilibrated
with buffer A, then concentrated to 15 mg/mL.
Crystallization and data collection
The crystallization of the RseAperiRseB complex
was performed using the microbatch method at
14C. The crystallization drop was prepared by mix-
ing 1 lL protein solution (8–10 mg/mL) and 1 lL
crystallization reagent (28% PEG550MME, 10 mM
ZnSO4, and 100 mM MES pH 6.5) under a layer of
Al’s oil (Hampton Research, Aliso Viejo, CA). The
crystals in a drop were flash-frozen in a cold nitro-
gen stream at 100 K without the addition of a cryo-
protectant, and the diffraction data were collected at
PLS-BL4A (Beam line 4A, Pohang Light Source,
South Korea; wavelength 1.0000 A˚ ). The diffraction
images were recorded to an ADSC Quantum 210
CCD detector. The diffraction data were indexed and
integrated
using
HKL2000
and
scaled
using
SCALEPACK.15
Structure determination
The RseAperiRseB structure was determined by
molecular replacement using PHASER16 with the
large domain (residues 26–200) of E.coli RseB10
(PDB 2P4B) as a template. Three large domains
were initially identified, and the small domains were
added to the model. The fourth RseB was generated
by a noncrystallographic symmetry operation. Sev-
eral
cycles
of
rigid
body,
positional,
simulated
annealing
and
B-factor
refinements,
and
model
rebuilding were conducted at a resolution of 2.3 A˚
using the CNS and COOT programs.17,18 The RseA
model was placed on the additional electron density.
The RseAperiRseB structure was refined further
using REFMAC.19 The final refinement with sol-
vents resulted in R and Rfree values of 23.8% and
27.0%, respectively. The data collection and refine-
ment statistics are summarized in Table I. The fig-
ures were drawn using PyMOL.20 The protein–pro-
tein interface was calculated using Protorp.21 The
coordinates and structure factors for the RseAperi
RseB complex have been deposited under accession
code 3M4W.
Summary
In this study, we characterized the inhibitory mecha-
nisms of RseB in the regulated proteolysis of RseA.
The C-terminal helix in RseA169–190 is a major con-
tributor to the formation of a stable complex with
RseB, whereas the N-terminal region of the periplas-
mic domain of RseA is necessary for the burial of
the DegS cleavage site within an inaccessible pocket
in the RseARseB complex. In the regulation of the
envelope-stress response, RseB functions by blocking
the access of DegS protease rather than converting
RseA into a compact structure that is resistant to
proteolysis, as the random coil structure around the
cleavage site is maintained in the complex. Accord-
ingly, in this study we explain why the release of
RseB is a prerequisite for the degradation of RseA
and the activation of the rE-dependent envelope
stress response at the atomic level. In the crystal
structure
of
the
RseARseB
complex,
RseB
is
unlikely to interfere with the further degradation of
the periplasmically cleaved RseA. Therefore, the
newly exposed Val148 will be readily recognized by
RseP for subsequent cleavage, which results in the
activation of the envelope-stress response.
References
1. Brown MS, Ye J, Rawson RB, Goldstein JL (2000)
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PROTEIN SCIENCE VOL 19:1258—1263
1263
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3M53
|
SET7/9 in complex with TAF10 peptide and AdoHcy
|
SET7/9 Catalytic Mutants Reveal the Role of Active Site Water
Molecules in Lysine Multiple Methylation*□
S
Received for publication,February 15, 2010, and in revised form, July 14, 2010 Published, JBC Papers in Press,August 1, 2010, DOI 10.1074/jbc.M110.114587
Paul A. Del Rizzo‡, Jean-Franc¸ois Couture‡§1, Lynnette M. A. Dirk¶, Bethany S. Strunk‡, Marijo S. Roiko‡,
Joseph S. Brunzelle, Robert L. Houtz¶, and Raymond C. Trievel‡2
From the ‡Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109, the §Ottawa Institute of
Systems Biology, Department of Biochemistry, Microbiology, and Immunology, University of Ottawa, Ottawa, Ontario K1H 8M5,
Canada, the ¶Department of Horticulture, Plant Physiology/Biochemistry/Molecular Biology Program, University of Kentucky,
Lexington, Kentucky 40546, and the Department of Molecular Pharmacology and Biological Chemistry, Northwestern University
Center for Synchrotron Research, Life Sciences Collaborative Access Team, Argonne, Illinois 60439
SET domain lysine methyltransferases (KMTs) methylate
specific lysine residues in histone and non-histone substrates.
These enzymes also display product specificity by catalyzing dis-
tinct degrees of methylation of the lysine -amino group. To
elucidate the molecular mechanism underlying this specificity,
we have characterized the Y245A and Y305F mutants of the
human KMT SET7/9 (also known as KMT7) that alter its prod-
uct specificity from a monomethyltransferase to a di- and a tri-
methyltransferase, respectively. Crystal structures of these
mutants in complex with peptides bearing unmodified, mono-,
di-, and trimethylated lysines illustrate the roles of active site
water molecules in aligning the lysine -amino group for methyl
transfer with S-adenosylmethionine. Displacement or dissocia-
tion of these solvent molecules enlarges the diameter of the
active site, accommodating the increasing size of the methylated
-amino group during successive methyl transfer reactions.
Together, these results furnish new insights into the roles of
active site water molecules in modulating lysine multiple meth-
ylation by SET domain KMTs and provide the first molecular
snapshots of the mono-, di-, and trimethyl transfer reactions
catalyzed by these enzymes.
SET domain enzymes represent a family of S-adenosylmethi-
onine (AdoMet)3-dependent methyltransferases that catalyze
the site-specific methylation of protein lysyl residues in a host
of proteins, including histones, transcription factors, chroma-
tin-modifying enzymes, ribosomal subunits, and other sub-
strates (1–3). In many instances, these modifications serve to
recruit effector proteins that recognize methyl-lysyl residues in
a sequence-dependent fashion (4). In addition, SET domain
KMTs exhibit product specificity, defined as their ability to cat-
alyze mono-, di-, or trimethylation of the lysine -amino group.
This specificity is biologically relevant because many methyl-
lysine-binding proteins can discriminate among different
degrees of lysine methylation (4). Thus, both the site and degree
of lysine methylation are critical to recognition by effector
proteins.
Structural and functional studies have identified a Phe/Tyr
switch in the active site of SET domain KMTs that governs their
respective product specificities (5, 6). According to this model,
KMTs that possess a tyrosine in the Phe/Tyr switch site are
limited to catalyzing lysine monomethylation, whereas en-
zymes that possess a phenylalanine or another hydrophobic
residue in this position display di- or trimethyltransferase activ-
ity. Mutational analysis of various SET domain KMTs, includ-
ing DIM-5 (KMT1) (6), SET7/9 (6), G9A (KMT1C) (5), and
SET8 (also known as PR-SET7 and KMT5A) (7, 8), has demon-
strated that substitutions in the Phe/Tyr switch result in pre-
dictable changes in product specificity. Several models have
been proposed to explain the mechanism by which the Phe/Tyr
switch site governs this specificity, including variations in the
diameter of the active site due to the size of Phe/Tyr switch
residue and steric hindrance by the tyrosine hydroxyl group (6,
9–11). However, our recent studies of the Phe/Tyr switch
mutant Y334F in the human histone H4 Lys-20 (H4K20) meth-
yltransferase SET8 indicate that the Phe/Tyr switch regulates
product specificity via a more subtle mechanism (8). Specifi-
cally, the switch modulates the binding of an active site water
molecule that in turn regulates the transition from mono-
methylation to multiple methylation.
Among the KMTs that have been structurally characterized,
SET7/9 has emerged as an archetypal model for studying the
catalytic mechanism and product specificity of the SET domain
family. Although initially isolated as a histone H3 Lys-4
(H3K4)-specific methyltransferase, this KMT has been shown
to regulate the functions of numerous non-histone substrates
through site-specific methylation (12–21). Early structural and
functional studies of SET7/9 identified two active site mutants,
* Thisworkwassupported,inwholeorinpart,byNationalInstitutesofHealth
Grant R01 GM073839 (to R. C. T.) and National Institutes of Health Admin-
istrative Supplement GM073839-04S1 (to R. C. T.) funded through the
American Recovery and Reinvestment Act. This work was also supported
by Department of Energy Grant DE-FG02-92ER20075 (to R. L. H.).
□
S The on-line version of this article (available at http://www.jbc.org) contains
supplemental Table 1.
The atomic coordinates and structure factors (codes 3M53, 3M54, 3M55, 3M56,
3M57,3M58,3M59,and3M5A)havebeendepositedintheProteinDataBank,
Research Collaboratory for Structural Bioinformatics, Rutgers University, New
Brunswick, NJ (http://www.rcsb.org/).
1 Supported by a Canadian Institutes of Health Research postdoctoral
fellowship.
2 To whom correspondence should be addressed: Dept. of Biological Chem-
istry, University of Michigan Medical School, 1150 West Medical Center Dr.,
5301 MSRB III, Ann Arbor, MI 48109. Tel.: 734-647-0889; Fax: 734-763-4581;
E-mail: rtrievel@umich.edu.
3 The abbreviations used are: AdoMet, S-adenosylmethionine; AdoHcy, S-ad-
enosylhomocysteine; KMT, lysine methyltransferase; ITC, isothermal titra-
tion calorimetry; CH–O, carbon-oxygen hydrogen bond; BisTris, 2-[bis(2-
hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 41, pp. 31849–31858, October 8, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31849
Y245A and Y305F, which change its product specificity. The
Phe/Tyr switch mutant Y305F alters SET7/9 product specificity
from a mono- to dimethyltransferase (6), whereas the Y245A
substitution converts the enzyme into a trimethyltransferase
with weak monomethyltransferase activity (11). These mutants
have been the subjects of numerous molecular modeling simu-
lations that have led to various models to explain their distinct
product specificities (22–26). However, the lack of structural
data for the SET7/9 Y245A and Y305F mutants in complex with
cognate methylated peptides has hindered our understanding
of the mechanisms that define the respective product specific-
ities of these mutants. Moreover, these structures would yield a
framework for visualizing the mono-, di-, and trimethylation
reactions catalyzed by SET domain KMTs.
To gain insight into the molecular basis of their product
specificities, we have determined high resolution crystal
structures of the SET7/9 Y245A and Y305F mutants in com-
plex with peptides of the TATA box-binding protein-associ-
ated factor TAF10 bearing the Lys-189 methylation site in
unmodified (K189), monomethylated (K189me1), dimethyl-
ated (K189me2), and trimethylated (K189me3) states. The
structures and accompanying biochemical data support a
model whereby changes in the occupancy or position of water
molecules in the active site are critical in establishing the prod-
uct specificities of the SET7/9 Y245A and Y305F mutants.
Together, our results provide new insights into the mechanisms
that govern SET domain product specificity and provide step-
wise snapshots of the lysine mono-, di-, and trimethyl transfer
reactions catalyzed by KMTs.
EXPERIMENTAL PROCEDURES
Cloning, Expression, and Purification of the SET7/9 Mutants—
The Y245A and Y305F mutants were introduced into the pHIS2
SET7/9 expression vector encoding residues 110–366 (27) via
QuikChange site-directed mutagenesis (Stratagene) and were
verified by dideoxy DNA sequencing. The plasmids encoding
wild type (WT) SET7/9 and the Y245A and Y305F mutants
were transformed into Rosetta2 DE3 cells (Novagen) and were
expressed as described previously (27, 28). In the course of
characterizing WT SET7/9, we observed that the enzyme co-
purified with AdoMet or another contaminant that resulted in
technical difficulties in the isothermal titration calorimetry
(ITC) experiments and co-crystallization trials with the TAF10
peptides. To overcome this problem, a denaturation and refold-
ing step was inserted in the purification scheme. The denatur-
ation and refolding protocol involved adding 6 M guanidine
HCl, 50 mM sodium phosphate, pH 7.5, and 500 mM NaCl to the
protein while it was immobilized on a nickel-Sepharose column
(GE Healthcare). The column was washed with this buffer, fol-
lowed by a solution of 6 M urea, 50 mM sodium phosphate, pH
7.5, 150 mM NaCl, and 10 mM 2-mercaptoethanol to remove the
cofactor from the denatured enzyme. A reverse gradient from 6
to 0 M urea was then performed in the same buffer to refold the
protein, which was subsequently eluted from the column using
a linear gradient of 0–500 mM imidazole in 50 mM sodium
phosphate, pH 7.5, 500 NaCl, and 10 mM 2-mercaptoethanol.
The refolded protein was digested with tobacco etch virus pro-
tease (29) during dialysis against 20 mM Tris, pH 7.5, 150 mM
NaCl, and 5 mM 2-mercaptoethanol and then purified using a
Superdex 200 gel filtration column (GE Healthcare). Protein
concentration was determined by its absorbance at 280 nm.
Synthetic Peptides—The TAF10 peptides bearing K189,
K189me1,
K189me2,
and
K189me3
(sequence,
acetyl-
SKSK189DRKYTL-amide) and a biotinylated TAF10 peptide
(sequence,
acetyl-SKSK189DRKYTLT(K-EZLink-S-S-biotin)-
amide) were synthesized and purified by New England Peptide,
Inc. Peptide concentrations were measured using the absorb-
ance of their tyrosine residue at 274 nm.
Crystallization and Data Collection—Crystals were pro-
duced by hanging drop vapor diffusion by mixing the crystalli-
zation solution in a 1:1 ratio with 10–20 mg/ml SET7/9, 1 mM
S-adenosylhomocysteine (AdoHcy), and 1.0–3.0 mM unmodi-
fied or methylated TAF10-K189 peptide in 20 mM Tris, pH 8.0,
100 mM NaCl, and 2 mM tris(2-carboxyethyl)phosphine. Crys-
tals were obtained at 20 °C in either 1.8–2.0 M (NH4)2SO4, with
0.1 M BisTris, pH 6.2–6.6, or in 0.95–1.1 M sodium citrate with
100 mM imidazole pH 8.0–8.4. In both crystallization condi-
tions, the final pH values were between pH 8.0 and 9.0. Crystals
in the (NH4)2SO4 condition were typically flash-frozen in the
mother liquor containing 25–30% glycerol, and the crystals in
the citrate condition were frozen in 1.6 M sodium citrate. Data
were collected at the Advanced Photon Source beamlines
21-IDG (LS-CAT) and 23-IDD (GM/CA-CAT). Images were
indexed, integrated, and scaled using HKL2000 (30). Structures
of the mutants were solved by molecular replacement using
MOLREP (31) with the coordinates of a previously reported
SET7/9 ternary complex used as the search model (Protein
Data Bank code 2F69). Successive rounds of model building and
refinement were carried out using Coot (32) and REFMAC (33),
respectively. The geometry of the models were verified by Mol-
Probity (34). Simulated annealing omit maps were calculated
using CNS (35) with the peptide and cofactor removed to elim-
inate model bias in the active site. Structural figures were ren-
dered using PyMOL (Schro¨dinger, LLC.).
Fluorescent Methyltransferase Assay—A coupled fluorescent
methyltransferase assay was used to measure the kinetic
parameters of WT SET7/9 and the Y245A and Y305F mutants
as reported previously, with the exception that 50–150 nM
enzyme, 100 M AdoMet, and varying concentrations of TAF10
peptide substrate were used (27, 36). Assays were performed in
triplicate, and a homocysteine calibration curve was used to
calculate the initial velocities. Kinetic parameters were calcu-
lated by plotting the velocities versus peptide concentration and
by fitting the Michaelis-Menten equation to the data via non-
linear regression using Prism 5.0 (GraphPad). In cases where
the Km value was beyond the measurable range of the assay, the
kcat/Km value was determined as described previously (7).
Calorimetry Experiments—ITC was performed at 20 °C using
a MicroCal VP-ITC calorimeter (GE Healthcare) with 0.12 mM
protein and 4 mM AdoHcy in 20 mM sodium phosphate, pH 7,
and 100 mM NaCl with 1.5 mM peptide as the injectant. Data
were processed, and equilibrium dissociation constants (KD)
and curve fitting errors were calculated from the binding iso-
therms using Origin 7.0 (OriginLab Corp.). WT SET7/9 and the
Y245A and Y305F mutants displayed ligand:protein binding
stoichiometries (N values) between 0.8 and 1.0, demonstrating
Lysine Methylation by SET7/9 Mutants
31850
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
that WT SET7/9 and its mutants were properly refolded due to
their ability to bind peptides in an 1:1 molar ratio.
TLC Product Analysis—Methyltransferase assays were per-
formed in triplicate at 37 °C with the biotinylated TAF10 pep-
tide (0.5 mM) for 30 min in the presence of WT SET7/9 (3
pmol), Y305F mutant (6 pmol), or the Y245A mutant (100
pmol). Assays contained 100 mM HEPES, pH 8.0, 50 mM NaCl,
1.0 mM [methyl-3H]AdoMet (diluted to a specific activity of 0.2
Ci/nmol with purified AdoMet (37)), 15 M Sulfolobus solfa-
taricus AdoHcy hydrolase (36), and 2 units of adenosine deami-
nase (Roche Applied Science) in a final volume of 20 l. The
reactions were terminated by addition of an equal volume of
200 mM MES, pH 5.1, prior to addition of a 2-fold molar excess
of immobilized avidin resin (UltraLink; Pierce). Biotinylated
peptides were allowed to bind at room temperature for 30 min,
and the resin was then collected by centrifugation (9000 g).
The resin was washed three times with 300 mM NaCl, and the
peptide was eluted overnight from the avidin resin by cleavage
of the disulfide bond in the linker of the peptide using 10 mM
tris(2-carboxyethyl)phosphine (Thermo Scientific). The resin
was incubated with additional 10 mM tris(2-carboxyethyl)phos-
phine the following day until the radiolabel was essentially
removed from the resin. The recovered peptides were hydro-
lyzed by incubation with 6 M HCl at 110 °C for 24 h. Subsequent
steps in measuring the radiolabel incorporated into the mono-,
di-, and trimethyl-lysine products were performed as reported
previously (8).
RESULTS
Functional Analysis of the SET7/9
Y305F Mutant—Prior studies of
SET7/9 by Zhang et al. (6) reported
that mutation of the Phe/Tyr switch
residue Tyr-305 to a phenylalanine
alters its product specificity from a
mono- to dimethyltransferase. We
verified these findings by demon-
strating that WT SET7/9 mono-
methylated the TAF10-K189 pep-
tide, whereas the Y305F mutant
mono- and dimethylated this sub-
strate, as demonstrated by mass
spectrometry (data not shown). We
next examined whether the Y305F
substitution altered the affinity of SET7/9 for the TAF10-K189
peptides using ITC (Fig. 1). A comparison of the KD values
revealed that SET7/9 Y305F bound the TAF10-K189 and
TAF10-K189me1 peptides 4- and 6-fold more tightly, respec-
tively, than the WT enzyme, whereas this mutant displayed a
substantially diminished affinity for the TAF10-K189me2 pep-
tide (Table 1). Although the WT enzyme and the Y305F mutant
exhibited discernable differences in their affinities for the
unmodified and monomethylated peptides, these variations are
modest and cannot account for their distinct product specific-
ities, suggesting that a kinetic effect during methylation may be
responsible.
To investigate this possibility, we characterized the kinetic
parameters of WT SET7/9 and the Y305F mutant using the
TAF10 peptides as substrates. Both enzymes methylated the
unmodified peptide with comparable kcat and Km values (Table
2). In analyzing the kinetic parameters for the methylation
of the monomethylated peptide by SET7/9 Y305F, we found
that this substrate displayed an elevated Km value that was
beyond the measurable range of the assay due to its limited
solubility. In this case, we measured the catalytic efficiency
(kcat/Km) for the methylation of this peptide and found that it
was methylated 15-fold less efficiently than the unmodified
peptide by SET7/9 Y305F. Given the fact that the Y305F mutant
exhibited a higher binding affinity for the TAF10-K189me1
peptide than the WT enzyme (Table 1), the kinetic data suggest
that a step in the reaction pathway following substrate binding
limits the catalytic efficiency of this mutant.
We next examined whether the Y305F mutant dimethylated
the TAF10-K189 peptide via a processive or a distributive
mechanism. In a processive mechanism, the methyl-lysine
substrate would remain bound to the enzyme during successive
methyl transfer reactions; thus, the concentration of an inter-
mediate, such as monomethyl-lysine, cannot exceed the en-
zyme concentration during the assay. In a distributive mecha-
nism, the intermediates are released into solution where they
accumulate prior to the next round of methylation, resulting in
an intermediate concentration that is greater than that of the
enzyme. Using a radiometric TLC assay and a biotinylated
TAF10 peptide, we quantified the amounts of monomethylated
products generated by the WT SET7/9 and the Y305F mutant
FIGURE 1. ITC analysis of WT SET7/9 and the Y245A and Y305F mutants. Representative titrations and
binding isotherms for the calorimetry experiments are shown for the unmodified TAF10-K189 peptide titrated
into WT SET7/9 (left plot), SET7/9 Y305F (middle plot), and SET7/9 Y245A (right plot). In each panel, the ITC
titration experiment (upper panel) is shown with the binding isotherms (lower panel) fit to a single binding site
model.
TABLE 1
Analysis of the binding affinity of WT SET7/9 and its catalytic
mutants for unmodified and methylated TAF10 peptides
SET7/9
TAF10 peptide
KD
a
M
WT
K189
4.9 0.20
WT
K189me1
4.0 0.36
Y305F
K189
1.3 0.10
Y305F
K189me1
0.62 0.065
Y305Fb
K189me2
70
Y245A
K189
4.0 0.25
Y245A
K189me1
3.3 0.10
Y245A
K189me2
5.8 0.22
Y245A
K189me3
11 0.28
a Curve fitting errors were calculated from the binding isotherms.
b An estimate of the affinity is reported due to weak peptide binding.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31851
(Table 3). The data demonstrate that comparable amounts of
monomethyl-lysine were generated when the quantity of
enzyme usedistakenintoaccount,inagreementwiththeirsimilar
turnover numbers for the TAF10-K189 peptide (Table 2). The
Y305F mutant also produced small but measurable quantities of
radiolabeled dimethyl-lysine product that were substantially
smaller than the amount of monomethyl-lysine generated.
Therefore, the TLC data are consistent with a distributive
mechanism for dimethylation by the Y305F mutant because the
amount of monomethyl-lysine produced exceeded the quantity
of enzyme used in the assay.
Structures of WT SET7/9 and the Y305F Mutant in Complex
with Unmodified and Methylated TAF10 Peptides—To deter-
mine the mechanism by which the Y305F substitution alters the
product specificity of SET7/9, we determined the crystal struc-
tures of this mutant bound to AdoHcy and TAF10-K189,
TAF10-K189me1, and TAF10-K189me2 peptides and com-
pared these to the structures of the WT SET7/9AdoHcy
TAF10-K189 complex (supplemental Table 1). The structures
of these complexes were determined to 1.85 Å or higher reso-
lution, permitting unambiguous modeling of the K189 side
chains in the active site of the enzyme based on simulated
annealing omit maps (Fig. 2). The ternary complexes of the WT
and the Y305F mutant superimpose with overall root mean
square differences of less than 0.3 Å for all aligned atoms, indi-
cating that neither the Y305F mutation nor the binding of the
various TAF10-K189 peptides results in substantial changes in
its overall structure.
An inspection of the active sites of the SET7/9 WT and
Y305F complexes illustrates the binding modes of the unmod-
ified and methylated forms of K189 in the TAF10 peptides (Fig.
2, A–D). The K189 side chain binds in an extended all trans
conformation in a deep pocket, termed the lysine binding chan-
nel, that is composed of Tyr-245, Thr-266, Leu-267, Ser-268,
Tyr-305, Tyr-335, and Tyr-337 (not shown for clarity) (Fig. 2A).
These residues interact with the aliphatic portion of the K189
side chain primarily through van der Waals contacts. The lysine
binding channel connects to the AdoMet-binding site on the
opposite face of the catalytic domain via an oxygen-lined
methyl transfer pore (38). During catalysis, the methyl group of
the cofactor is positioned within the methyl transfer pore
for the SN2 reaction with the -amino group of the lysine or
methyl-lysine substrate (see below).
To lower the activation barrier for this reaction, the lysine
-amine nucleophile is aligned for methyl transfer through a
hydrogen bond network within the active site. In the WT
enzyme, the K189 -amino group hydrogen bonds to the
hydroxyl group of Tyr-245 as well as to two water molecules
(Fig. 2A). One of the water molecules (termed water 1), is coor-
dinated in a solvent pocket, through hydrogen bonds to the
carbonyl oxygens of Gly-292 and Ala-295 and to the hydroxyl
group of the Phe/Tyr switch residue Tyr-305. This solvent
pocket is structurally conserved in SET domain KMTs and has
an important role in defining product specificity through the
adjacent Phe/Tyr switch residue, as shown in our prior studies
of the human H4K20 methyltransferase SET8 (8). The other
water molecule is bound within the methyl transfer pore
between the lysine substrate and the thioether sulfur atom of
AdoHcy through hydrogen bonds to Tyr-245, Asn-265, and
His-293 in SET7/9 and the TAF10-K189 -amino group. This
water is not observed in other structures of SET7/9 ternary
complexes and may represent the approximate position that
the AdoMet methyl group occupies in the methyl transfer pore
in the Michaelis complex.
In structures of the Y305F ternary complexes, the K189,
K189me1, and K189me2 side chains also adopt extended trans
side chain geometries within the lysine binding channel that are
stabilized via hydrogen bonding to Tyr-245 in the enzyme (Fig.
2, B–D). The orientations of the K189me1 and K189me2 side
chains are further maintained through carbon-oxygen (CH–O)
hydrogen bonding between the methyl groups and oxygen
atoms within the vicinity of the methyl transfer pore, as
reported previously in other SET domain KMT structures (8,
10, 38). A superimposition of the SET7/9 WT and Y305F com-
plexes underscores the similarity of the lysyl binding conforma-
tions (Fig. 2E). However, there are notable differences in the
hydrogen bond patterns and occupancy of water 1 within the
solvent pocket in the Y305F mutant compared with the WT
enzyme. Specifically, the Y305F substitution results in the loss
of one hydrogen bond to water 1 in the structures of the TAF10-
K189 and TAF10-K189me1 complexes (Fig. 2, B and C). In con-
trast, water 1 is absent in TAF10-K189me2 complex, and the
vacated solvent pocket is occupied by one of the methyl groups
TABLE 2
Kinetic parameters of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
TAF10 peptide substrate
Km
a
kcat
a
kcat/Km
a
M
min1
M1 min1 103
WT
K189
160 17
17 0.62
110 17
Y305F
K189
88 5.0
17 0.30
190 11
Y305Fb
K189me1
11 0.50
Y245A
K189
200 35
0.53 0.04
2.6 0.47
Y245A
K189me1
210 23
5.9 0.23
28 3.3
Y245A
K189me2
400 29
6.5 0.16
15 1.2
a Curve fitting errors were calculated from the hyperbolic fits for the Michaelis-Menten equation.
b Km and kcat were not determined because the Km value was beyond the measurable range; therefore, the kcat/Km value is reported.
TABLE 3
Product analysis of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
Quantity
of enzyme
Measured
product
Amount of
product formeda
nmol
nmol
WT
0.003
Kme1
0.65 0.07
Y305F
0.006
Kme1
1.5 0.49
Kme2
0.033 0.009
Y245A
0.100
Kme1
0.80 0.22
Kme2
0.39 0.021
Kme3
0.076 0.019
a Standard deviation was calculated from triplicate measurements.
Lysine Methylation by SET7/9 Mutants
31852
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
of the dimethyl -amine (Fig. 2D). This methyl group forms a
3.5-Å CH–O hydrogen bond to the carbonyl oxygen of Ala-295,
further stabilizing the binding of the dimethyl-lysine side chain.
A homologous dimethyl-lysine-binding mode and CH–O
hydrogen bond was observed in our prior structural studies of
the SET8 Y334F Phe/Tyr switch mutant that confers an analo-
gous change in product specificity from a mono- to a dimeth-
yltransferase (8). A structural alignment of the active sites of the
SET7/9 Y305F and SET8 Y334F mutants bound to cognate
dimethylated peptides illustrates that the coordinates of the
dimethyl-lysyl side chains are virtually superimposable, with
one methyl group oriented toward the methyltransfer pore and
the second positioned within the vacant solvent pocket (Fig.
2F). Taken together, the structures of the SET7/9 Y305F com-
plexes and the similarities in the dimethyl-lysine conforma-
tions in the SET7/9 Y305F and SET8 Y334F mutants imply that
the Phe/Tyr switch governs product specificity through a con-
served mechanism whereby it indirectly influences the binding
modes of the methyl-lysine side chain by modulating the affin-
ity of the water molecule (water 1) bound in the solvent pocket.
Biochemical Characterization of
the SET7/9 Y245A Mutant—Previ-
ous studies by Xiao et al. (11)
reported that the Y245A mutation
yields an unusual change in the
product
specificity
of
SET7/9,
converting the enzyme to a trimeth-
yltransferase with weak monometh-
yltransferase activity. We deter-
mined that the SET7/9 Y245A could
mono-, di-, and trimethylate the
TAF10-K189 peptide by mass spec-
trometry (data not shown) and TLC
(Table 3), confirming the earlier
studies of Xiao et al. (11). ITC
analysis revealed that the Y245A
mutant displayed comparable KD
values
for
the
unmodified
and
methylated TAF10-K189 peptides
(Fig. 1), although its affinity for the
trimethylated peptide was modestly
diminished in comparison with the
other peptides (Table 1). The ITC
data demonstrate that the Y245A
mutant
bound
the
unmodified,
mono-,
and
dimethylated
sub-
strates with equivalent affinities,
suggesting that a kinetic effect or
a structural alteration in the active
site may be responsible for its
diminished
activity
toward
un-
modified substrates.
To gain further insight into its
peculiar
product
specificity,
we
characterized the kinetic properties
of
the
SET7/9
Y245A
mutant.
Steady state analysis demonstrated
that this mutant displayed similar
Km values for the unmodified, mono- and dimethylated TAF10
peptides (Table 2). However, the turnover number for the
TAF10-K189 peptide was diminished over 10-fold versus the
methylated peptides and was reduced 30-fold versus the WT
enzyme, in agreement with the weak monomethyltransferase
activity reported by Xiao et al. (11). In addition, we investigated
whether this mutant catalyzes lysine trimethylation via a pro-
cessive or distributive mechanism as described for SET7/9
Y305F. The TLC data illustrate that the mono- and dimethyl-
lysine intermediates accumulated at quantities greater than
that of the enzyme used in the assay, indicating that SET7/9
Y245A obeys a distributive mechanism, analogous to the Y305F
mutant (Table 3).
Structures of SET7/9 Y245A Bound to Unmodified and Meth-
ylated TAF10 Peptides—To elucidate the mechanism underly-
ing its unusual product specificity, we determined the crystal
structures of SET7/9 Y245A in complex with AdoHcy and
unmodified, mono-, di-, and trimethylated TAF10 peptides
(supplemental Table 1). These complexes superimpose with
the structure of the WT SET7/9AdoHcyTAF10-K189 com-
FIGURE 2. Crystal structures of WT SET7/9 and the Y305F mutant in complex with AdoHcy and unmodi-
fied and methylated TAF10 peptides. A, active site of WT SET7/9 bound to AdoHcy and TAF10-K189 and the
active site of SET7/9 Y305F bound to AdoHcy and TAF10-K189 (B), TAF10-K189me1 (C), and TAF10-K189me2
peptides (D). A–D, the K189 residue in the TAF10 peptide is depicted with yellow carbon atoms with the
corresponding Fo – Fc simulated-annealing omit maps (contoured at 2.5 ) shown. AdoHcy is colored with
green carbon atoms, and SET7/9 carbon atoms are colored gray, with the exception of Tyr-305/Y305F
(magenta) and Tyr-245 (cyan). Gly-264 to Ser-268 are shown, and the main chain atoms of Gly-292 to Ala-295
are depicted for clarity. Red dashed lines indicate conventional hydrogen bonds, and blue dashed lines indicate
CH–O hydrogen bonds. Cyan dashed lines indicate weak CH–O hydrogen bonds between 3.5 and 3.7 Å in
length. The water molecule in the solvent binding pocket is numbered 1. E, overlay of the active sites of the WT
enzyme (light blue carbon atoms) and the Y305F mutant (green, yellow, and pink carbon atoms correspond to
K189, K189me1, and K189me2, respectively). Red dashed lines indicate coordination of the waters in the struc-
turesoftheY305Fmutant,andthelightbluedashedlineindicateshydrogenbondingthatonlyoccursintheWT
enzyme. F, superimposition of the active sites of the SET7/9 Y305FAdoHcyTAF10-K189me2 and SET8
Y334FAdoHcyH4K20me2 (Protein Data Bank code 3F9X) complexes shown with green and yellow carbon
atoms, respectively.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31853
plex with root mean squared differences of less than 0.4 Å for all
aligned atoms, indicating that the Y245A mutant does not per-
turb the overall structure of the enzyme. Simulated annealing
omit maps illustrate that K189 side chains are bound within the
lysine binding channel through hydrogen bonds and van der
Waals contacts (Fig. 3, A–D), although the interactions and
binding modes are distinct from those in the complexes of WT
SET7/9 and the Y305F mutant (Fig. 2, A–D). In the unmodified
TAF10 peptide complex, the K189 -amino group forms a weak
hydrogen bond with the carbonyl oxygen of Gly-264 (Fig. 3A),
whereas the -amino groups of K189me1 and K189me2 hydro-
gen bond to the hydroxyl group of Tyr-305 in the mono- and
dimethylated peptide complexes (Fig. 3, B and C). The confor-
mations of the K189me1 and K189me2 side chains are further
stabilized by water-mediated hydrogen bonding and through
CH–O hydrogen bonding to their methyl groups. In the
TAF10-K189me3 peptide complex, the trimethyl-lysine side
chain is coordinated exclusively through direct and water-me-
diated CH–O hydrogen bonds to its methyl groups because the
quaternary -ammonium cation cannot engage in hydrogen
bonding (Fig. 3D).
A structural alignment of the four SET7/9 Y245A complexes
illustrates distinct binding modes for the unmodified versus the
methylated K189 side chains, highlighting the selectivity of this
mutant for methylated substrates. The side chains of K189me1,
K189me2, and K189me3 roughly overlay with their respective
-amino groups superimposed and adopt slightly kinked con-
formations (Fig. 3E), as opposed to the extended trans geome-
try of the unmodified and methylated lysines in the complexes
of the WT enzyme and the Y305F mutant (Fig. 2, A–D). Con-
versely, the unmodified K189 side chain does not superimpose
with its methylated counterparts and is oriented in an alterna-
tive configuration due to its hydrogen bonding to Gly-264 (Fig.
3, A and E). An overlay of the structures of the WT enzyme and
Y245A mutant bound to the unmodified TAF10 peptide illus-
trates that the side chains of K189 do not superimpose and that
the K189 -amino group appears to be misaligned with AdoHcy
in the Y245A complex (Fig. 3F). This suboptimal alignment
may explain the diminished kcat value of SET7/9 Y245A mutant
toward substrates with unmodified lysines (Table 2).
A comparison of the structures of the SET7/9 Y245A and
Y305F complexes yields a molecular explanation for the differ-
ent product specificities of these two mutants. In the SET7/9
Y305F complexes, Tyr-245 aligns the K189 -amino group for
methyl transfer through hydrogen bonding to its hydroxyl
group (Fig. 2, B–D). Conversely, in the Y245A mutant, the
K189me1 and K189me2 -amino groups are oriented through
hydrogen bonding to Tyr-305 (Fig. 3, B and C). These distinct
hydrogen bond patterns impart differences in the conforma-
tions of the lysyl side chains due to the relative orientations of
Tyr-245 and Tyr-305 in the lysine binding channel. Specifically,
the kinked conformation adopted by the K189me1 and
K189me2 side chains in the Y245A complexes (Fig. 3, B and C)
may contribute to the differences in the turnover numbers of
this mutant versus those of the WT enzyme and the Y305F
mutant (Table 2). In addition, the dimethyl -amino group of
the K189me2 side chain binds in distinct orientations in the
Y245A and Y305F mutants due to their hydrogen bonding to
FIGURE 3. Crystal structures of the SET7/9 Y245A mutant in complex with
AdoHcy and unmodified and methylated TAF10 peptides. Active site of
SET7/9 Y245A bound to AdoHcy and TAF10-K189 (A), TAF10-K189me1 (B),
TAF10-K189me2 (C), and TAF10-K189me3 peptides (D). A–D, Fo – Fc simulated-
annealing omit maps (contoured at 2.5 ) for the unmodified and methylated
K189 side chains are illustrated. The residues and hydrogen bonds in each com-
plexarecoloredasinFig.2.Thewatermoleculesinthelysinebindingchannelsof
the Y245A complexes are numbered 1–4, as described in the text. E, superimpo-
sition of the active sites of the Y245A complexes bound to the four methylated
statesofTAF10-K189(unmodified,mono-,di-,andtrimethylatedshowningreen,
yellow, pink, and blue, respectively). F, overlay of WT SET7/9 (blue carbons) and
SET7/9 Y245A (green carbons) in complex with TAF10-K189. Waters correspond-
ing to the WT and Y245A structures are colored cyan and green, respectively.
G, alignment of the active sites of the Y245A (yellow carbons) and Y305F (green
carbons) mutants in complex with the TAF10-K189me2 peptide. Hydrogen
bonds from the Y305F structure are shown as green dashed lines, and waters and
hydrogen bonds in the Y245A structure are shown in yellow and orange,
respectively.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
Tyr-305 and Tyr-245, respectively (Fig. 3G). In the Y305F
mutant, hydrogen bonds to the dimethyl -amino group cou-
pled with steric constraints in the lysine binding channel pre-
vent the K189me2 side chain from undergoing a conforma-
tional change that is conducive to trimethylation (Fig. 2D),
consistent with its dimethyltransferase activity. However, in the
Y245A mutant, the alanine substitution enlarges the diameter
of the lysine binding channel, accommodating trimethyl-lysine
(Fig. 3D). In addition, the larger diameter would permit the
dimethyl-lysine substrate to undergo the conformational reor-
ganization necessary to align the -amino group in a productive
geometry for trimethylation.
A major difference in the active site of the Y245A mutant
versus the other SET7/9 structures is the presence of several
water molecules bound in the cavity generated by the Y245A
mutation. In the structure of the Y245A mutant bound to
TAF10-K189, three water molecules (waters 2–4) occupy this
cavity and are arranged in a triangular geometry (Fig. 3A). In
addition, water 1 shifts 1.6 Å from its position in the solvent
pocket toward water 2 to which it forms a hydrogen bond (Fig.
3, A and E). The shift in water 1 was unexpected given its con-
served orientation in the solvent pocket of the SET7/9 WT and
Y305F complexes (Fig. 2, A–C) as well as in the structures of
other SET domain KMTs (8). This displacement is presumably
related to the alternative conformation of the K189 side chain
whose -amino group is too distant (4.3 Å) to form a productive
hydrogen bond to water 1. Conversely, in the Y245A complexes
bound to TAF10-K189me1 and TAF10-K189me2, water 1
remains tightly bound in the solvent pocket through hydrogen
bonds to Tyr-305 hydroxyl group, the carbonyl oxygens of Gly-
292 and Ala-295, and the K189 -amino group (Fig. 3, B, C, and
E), analogous to its binding in the WT enzyme (Fig. 2A). How-
ever, in the TAF10-K189me3 complex, one of the methyl
groups of the trimethyl -ammonium cation is oriented into the
solvent pocket (Fig. 3D), similar to the dimethyl-lysine binding
mode observed in the Y305F mutant (Fig. 2D). The binding of
the methyl group in the solvent pocket displaces water 1 by 3.2
Å relative to its position in the TAF10-K189me1 complex (Fig.
3E), thereby avoiding a steric clash with the trimethylated
-ammonium group. Variations in the occupancy of water 2 are
also seen in the different Y245A structures. Water 2 is bound in
similar orientations in the active site of the unmodified and
monomethylated peptide complexes but is absent in the di- and
trimethylated peptide complexes due to the binding of a methyl
group in this position (Fig. 3, A–E). In summary, the changes in
the positions or occupancies of waters 1 and 2 correlate with the
binding modes of the unmodified and methylated K189 within
the active site of the Y245A mutant.
Catalytic Models of Lysine Multiple Methylation by SET7/9
Y245A, and Y305F—The structures of the SET7/9 complexes
reported here offer a prime opportunity to generate stepwise
models for lysine mono-, di-, and trimethylation by a SET
domain KMT. We modeled the AdoMet-bound Michaelis
complexes by superimposing the SET7/9 product complexes
with the previously reported structure of the SET7/9-AdoMet
binary complex (Fig. 4) (39). The conformations of the mono-
and dimethyl -amino groups in the Michaelis complexes were
inferred from the coordinates of the corresponding dimethyl-
and trimethyl-lysine products, respectively. In addition, we
modeled the -amino group in a deprotonated state with its
hydrogen atoms oriented toward the hydrogen bond acceptors
that align the lysyl side chain for methylation. As a basis for this
comparison, we first modeled the monomethylation reaction
catalyzed by WT SET7/9 (Fig. 4A). In the substrate ternary
complex, the lysine -amine is aligned with the methyl group
and sulfonium cation of AdoMet through a hydrogen bond to
the Tyr-245 hydroxyl group and water 1 in the solvent pocket.
The values of the reaction distance and angle are 2.8 Å and 153°,
respectively, in approximate agreement with the linear geome-
try of a SN2 methyl transfer reaction calculated in other mod-
eled substrate complexes (8, 10). In the product complex, the
monomethyl-lysine side chain is bound in an extended confor-
mation with its methyl group oriented within the methyl trans-
fer pore, thereby obstructing AdoMet binding. Furthermore,
water 1 remains tightly coordinated in the solvent pocket
through four hydrogen bonds to Gly-292, Ala-295, Tyr-305,
and the monomethyl -amino group. These interactions hinder
the dissociation of water 1 and the related rearrangement of the
monomethyl-lysine side chain required for a second methyl
transfer reaction, explaining why the WT enzyme cannot cata-
lyze di- and trimethylation. These findings concur with the
FIGURE 4. Catalytic models of the methyl transfer reactions catalyzed by
WT SET7/9 and the Y305F mutant. A, monomethylation of TAF10-K189 by
the WT enzyme. The reaction scheme depicts the modeled substrate ternary
complex (left) and the product complex (right) for the transfer of the methyl
group from AdoMet (green carbon atoms) to K189 in TAF10 (yellow carbons),
yielding AdoHcy and K189me1. The red arrow indicates the direction of the
nucleophilic attack of the deprotonated -amino group on the AdoMet
methyl group. The transferred methyl group is colored green, and the white
atoms represent the hydrogens of the -amino group. Hydrogen bonds
and residues in the enzyme active site are illustrated as in Fig. 2. The
reaction distance and angle are labeled in red. B and C, models of the
Y305F mutant for the first methyl transfer reaction with TAF10-K189 (B)
and second methyl transfer reaction with TAF10-K189me1 (C). Color
schemes are the same as in A.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31855
model for SET7/9 product specificity reported in previous
structural and functional studies (6, 11).
Similar reaction geometry is observed in the model for the
monomethyl transfer reaction catalyzed by SET7/9 Y305F.
Hydrogen bonds from the Tyr-245 hydroxyl group and water 1
align the lysine -amino group with the AdoMet methyl group
at a distance of 2.1 Å and an angle of 160°, equivalent to those
measured in the Michaelis complex of the WT enzyme (Fig. 4, A
and B). In the product complex, monomethyl-lysine adopts an
extended trans configuration analogous to that in the WT
enzyme. For dimethylation to occur, the monomethyl-lysine
must undergo a conformational change in which its methyl
group is rotated out of the methyl transfer path with AdoMet.
The structure of the Y305F mutant bound to the dimethylated
TAF10 peptide (Fig. 2D) implies that this rearrangement occurs
through the dissociation of water 1 due to the loss of the Tyr-
305 hydrogen bond in the solvent pocket. The dissociation of
water 1 would enable the monomethyl-lysine side chain to
adopt an alternative conformation through a rotation about its
C–N bond, projecting the methyl group into the solvent
pocket (Fig. 4C). This rotation reorients the methyl group out of
the methyl transfer path while realigning the monomethyl -
amino group for a second methylation reaction through a direct
hydrogen bond to the Tyr-245 hydroxyl group and a CH–O
hydrogen bond between its methyl group and Ala-295. The
modeled reaction geometry for monomethyl-lysine substrate
complex of the Y305F mutant (2.7 Å, 160°) is equivalent to that
of the first methyl transfer reaction in SET7/9 Y305F. These
geometries concur with our previous models for mono- and
dimethylation catalyzed by SET8 Y334F (8), illustrating that the
orientation of a methyl group into the solvent pocket is a con-
served feature of SET domain KMTs that catalyze multiple
methylation.
In addition, we modeled the methyl transfer reactions cata-
lyzed by SET7/9 Y245A (Fig. 5). In the model of the lysine sub-
strate complex, the -amino group is aligned for methyl transfer
by a hydrogen bond to the carbonyl oxygen of Gly-264, result-
ing in a short reaction distance (2.3 Å) and a suboptimal reac-
tion angle (141°) with the methyl group of AdoMet (Fig. 5A).
This misalignment appears to be a direct consequence of the
Y245A mutation that abolishes hydrogen bonding to the -
amino group, illustrating that the suboptimal orientation of the
-amine likely contributes to the diminished activity of this
mutant toward unmodified substrates (11). Conversely, in the
modeled monomethyl-lysine substrate complex for SET7/9
Y245A, Tyr-305 hydrogen bonds to the -amino group, aligning
it for methyl transfer (Fig. 5B). In addition, CH–O hydrogen
bonds to the monomethyl-lysine methyl group and the dissoci-
ation of water 2 from the active site also contribute to reposition-
ing the -amino group for dimethylation. Collectively, these inter-
actions orient the -amine in a reaction angle of 165° that is more
conducive to methyl transfer. However, the reaction distance for
dimethylation is 0.6 Å longer than that in the corresponding
Y305F model because Tyr-305 is positioned further from
AdoMet than Tyr-245 (Figs. 4C and 5B).
In the third methyl transfer reaction catalyzed by SET7/9
Y245A, the lone pair of electrons of the dimethyl-lysine -
amino group acts as the nucleophile and thus cannot engage in
hydrogen bonding. The structure of the trimethyl-lysine prod-
uct complex (Fig. 3D) implies that the dimethyl -amine is
aligned via CH–O hydrogen bonds to its methyl groups, as
shown in the model of the Michaelis complex for this reaction
(Fig. 5C). These CH–O hydrogen bonds restrain the orienta-
tion of the -amino group and position one of the methyl
groups into the solvent pocket, displacing water 1 as discussed
earlier (Fig. 3, D and E). These interactions cumulatively align
the -amino group and AdoMet methyl group with a reaction
distance of 3.0 Å and angle of 162° (Fig. 5C). Taken together, the
models of the substrate complexes for SET7/9 Y245A suggest
that CH–O hydrogen bonds play an increasingly important role
in aligning the methylated -amino group in successive rounds
of methyl transfer.
DISCUSSION
The structural and functional characterization of the
SET7/9 Y245A and Y305F mutants presented here yields
new insights into the mechanism underlying the product
specificity of SET domain KMTs. Importantly, it resolves a
general paradox concerning this specificity. How does the
active site constrain the motion of the lysine -amino group
to align it for methyl transfer with AdoMet, while providing
adequate volume to accommodate the mono-, di-, and tri-
methylated lysine side chain generated during multiple
methyl transfer reactions? The structures of the Y305F and
Y245A mutants resolve this paradox, illustrating that alter-
ations in the positions or occupancies of water molecules
within their active sites generate the space required to
FIGURE 5. Models for the methyl transfer reactions catalyzed by the
SET7/9 Y245A mutant. Models of the Y245A mutant for the first methyl
transfer reaction with TAF10-K189 (A), the second methyl transfer reaction
with TAF10-K189me1 (B), and the third methyl transfer reaction with TAF10-
K189me2 (C). Residues and hydrogen bonds are depicted as in Fig. 3.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
accommodate the multiply methylated -amine produced
during successive catalytic cycles. Minor perturbations in
the side chains of certain active site residues, such as Tyr-
305, are also observed in alignments of the WT enzyme and
the Y245A and Y305F complexes, although these changes are
modest compared with the displacement or dissociation of
the water molecules in the active site. These findings suggest
that the waters function as transient place holders that facil-
itate the SN2 methyl transfer reaction. During monomethy-
lation, they function to constrain the movement of the lysine
-amino group by mediating hydrogen bonds between the
substrate and enzyme, thereby promoting the linear align-
ment with the methyl group and sulfonium cation of
AdoMet (Fig. 4, A and B). During di- and trimethylation, the
water molecules either relocate within the lysine binding
channel or dissociate from the enzyme, yielding the space
required to rotate the methyl group away from the methyl
transfer pore and to realign the -amine in productive geom-
etry for the next methyl transfer reaction (Figs. 4C and 5, B
and C). These findings agree with our prior analysis of the
SET8 Phe/Tyr switch mutant in which we demonstrated that
the Y334F substitution attenuates hydrogen bonding to the
water molecule bound in the solvent pocket, promoting its
dissociation and the conformational changes necessary for
lysine dimethylation (8). Indeed, there is a nearly identical
alignment of the dimethyl-lysine side chains in the structures of
SET7/9Y305FandSET8Y334Fcomplexes,despitethedifferences
in the orientations of the Phe-305 and Phe-334 side chains in each
structure (Fig. 2F). Finally, both the SET8 Y334F (8) and SET7/9
Y305F mutants (Table 2) displayed diminished catalytic efficien-
cies for lysine dimethylation versus monomethylation. These dif-
ferences may reflect the kinetics of the reorganization within the
active site, including the dissociation of the water molecule from
the solvent pocket and the concomitant realignment of the
monomethyl-lysine into a productive geometry for dimethylation.
In addition to their place-holding role, the active site waters
may also facilitate the deprotonation of the lysine -amino
group between methyl transfer reactions. For methylation to
occur, the -amino group must be deprotonated to function as
the nucleophile in the SN2 methyl transfer reaction with
AdoMet (Figs. 4 and 5). Although the pKa value of the lysine
-amine in solution is 10.5, molecular dynamics simulations
by Zhang and Bruice (25, 26) indicate that this value diminishes
to 8.2 upon formation of the SET7/9 Michaelis complex due to
the proximity of the AdoMet sulfonium cation and the low
dielectric constant of the active site. Furthermore, their simu-
lations show that a chain of water molecules facilitates the dep-
rotonation of the -amino group prior to methyl transfer, trans-
ferring the proton to bulk solvent. Although these water
molecule chains are not evident in our crystal structures, the
Y305F and Y245A complexes suggest another potential mech-
anism for deprotonation. In the dimethyl-lysine complexes of
the Y305F and Y245A mutants, the dissociation of water 1 and
2, respectively, from the lysine binding channel requires that
the solvent-mediated hydrogen bond to the -amino group is
broken (Figs. 2D and 3C). It is conceivable that these waters
dissociate from the active site as hydronium ions, promoting
the realignment and deprotonation of the methyl -amino
group for the next methyl transfer reaction.
A comparison of the SET7/9 Y305F and SET8 Y334F com-
plexes yields insights into the mechanism by which the Phe/Tyr
switch influences water binding within the solvent pocket. The
phenylalanine substitution in the Phe/Tyr switch results in the
loss of a single hydrogen bond to the water molecule (water 1) in
the solvent pocket compared with the four hydrogen bonds that
coordinate the solvent molecule in WT SET7/9 (Fig. 2, A and B)
and SET8 (7, 8). Although this attenuation in hydrogen bonding
may appear insignificant, this difference is nonetheless impor-
tant for at least two reasons. First, theoretical calculations indi-
cate that, on average, water molecules form 3.5 hydrogen
bonds in solutions (40, 41). This value is greater than the num-
ber of hydrogen bonds coordinating water 1 in the solvent
pocket in SET7/9 Y305F (Fig. 2, B and C) as well as in SET8
Y334F and other di- and trimethyltransferases that possess a
hydrophobic residue in the Phe/Tyr switch site (8). From the
perspective of the water molecule, the greater hydrogen bond-
ing potential in solution would tend to thermodynamically
favor its dissociation from the solvent pocket in SET domain
KMTs that lack a tyrosine in the Phe/Tyr switch position. Sec-
ond, the ordered binding of water molecules observed in the
active sites of SET domain ternary complexes represents an
unfavorable entropy compared with their diffusion in bulk sol-
vent. In WT SET7/9 (Fig. 2A) and SET8 (7, 8), this entropic
penalty can be partially offset through the favorable enthalpy of
binding associated with the four hydrogen bonds that coordi-
nate the water within the solvent pocket. It is conceivable that
the loss of the tyrosine-mediated hydrogen bond in the Phe/Tyr
switch shifts the equilibrium in favor of dissociation of the
water molecule from the solvent pocket, thereby facilitating
dimethylation in SET7/9 Y305F, SET8 Y334F, and other di- and
trimethyltransferases.
The structures of the SET7/9 Y245A and Y305F complexes
illustrate the interactions that align the lysine -amino group
during the methyl transfer reactions in each enzyme. In the WT
enzyme and the Y305F mutant, hydrogen bonding to the
hydroxyl group of Tyr-245 appears to be critical in properly
aligning the -amine for methyl transfer (Fig. 4). Tyr-245 is
conserved in the sequences of many SET domain KMTs (8, 42),
and substitutions of this residue generally impair or abolish
activity, indicating its importance in catalysis (8, 43). However,
SET7/9 appears to be an exception to this rule, as the Y245A
mutant is not only active but is capable of catalyzing lysine
trimethylation. In this mutant, Tyr-305 appears to assume the
role of Tyr-245 by hydrogen bonding to the monomethylated
-amino group to align it for methyl transfer with AdoMet, as
illustrated in the modeled substrate complex for the dimethy-
lation reaction (Fig. 5B). Conversely, in the model for trimethy-
lation, the Tyr-305 hydroxyl group does not hydrogen bond to
the -amine but instead participates in a CH–O hydrogen bond
with one of the methyl groups to assist in aligning the dimethy-
lated -amine for the methyl transfer reaction (Fig. 5C). Addi-
tional structural and functional studies of the SET domain tri-
methyltransferases will aid in further illuminating the roles
of CH–O hydrogen bonds in facilitating lysine multiple
methylation.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31857
Acknowledgments—We acknowledge S. Schiebold for assistance in
protein expression, purification, and crystallization and S. Anderson
and R. Sanishvili for their assistance with x-ray data collection. We
also thank S. Bulfer and S. Horowitz for reading the manuscript and
providing useful comments. This work utilized the Protein Structure
Facility of the Michigan Diabetes Research and Training Center, Uni-
versity of Michigan, supported by National Institutes of Health Grant
DK020572, NIDDK. Use of the Advanced Photon Source was sup-
ported by the United States Department of Energy, Basic Energy Sci-
ences, Office of Science, under Contract DE-AC02-06CH11357.
GM/CA CAT has been funded in whole or in part by National
Institutes of Health NCI Grant Y1-CO-1020 and NIGMS Grant
Y1-GM-1104. Use of the LS-CAT Sector 21 was supported by Michi-
gan Economic Development Corporation and the Michigan Technol-
ogy Tri-Corridor Grant 085P1000817 for the support of this research
program.
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Lysine Methylation by SET7/9 Mutants
31858
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
|
3M54
|
SET7/9 Y305F in complex with TAF10 peptide and AdoHcy
|
SET7/9 Catalytic Mutants Reveal the Role of Active Site Water
Molecules in Lysine Multiple Methylation*□
S
Received for publication,February 15, 2010, and in revised form, July 14, 2010 Published, JBC Papers in Press,August 1, 2010, DOI 10.1074/jbc.M110.114587
Paul A. Del Rizzo‡, Jean-Franc¸ois Couture‡§1, Lynnette M. A. Dirk¶, Bethany S. Strunk‡, Marijo S. Roiko‡,
Joseph S. Brunzelle, Robert L. Houtz¶, and Raymond C. Trievel‡2
From the ‡Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109, the §Ottawa Institute of
Systems Biology, Department of Biochemistry, Microbiology, and Immunology, University of Ottawa, Ottawa, Ontario K1H 8M5,
Canada, the ¶Department of Horticulture, Plant Physiology/Biochemistry/Molecular Biology Program, University of Kentucky,
Lexington, Kentucky 40546, and the Department of Molecular Pharmacology and Biological Chemistry, Northwestern University
Center for Synchrotron Research, Life Sciences Collaborative Access Team, Argonne, Illinois 60439
SET domain lysine methyltransferases (KMTs) methylate
specific lysine residues in histone and non-histone substrates.
These enzymes also display product specificity by catalyzing dis-
tinct degrees of methylation of the lysine -amino group. To
elucidate the molecular mechanism underlying this specificity,
we have characterized the Y245A and Y305F mutants of the
human KMT SET7/9 (also known as KMT7) that alter its prod-
uct specificity from a monomethyltransferase to a di- and a tri-
methyltransferase, respectively. Crystal structures of these
mutants in complex with peptides bearing unmodified, mono-,
di-, and trimethylated lysines illustrate the roles of active site
water molecules in aligning the lysine -amino group for methyl
transfer with S-adenosylmethionine. Displacement or dissocia-
tion of these solvent molecules enlarges the diameter of the
active site, accommodating the increasing size of the methylated
-amino group during successive methyl transfer reactions.
Together, these results furnish new insights into the roles of
active site water molecules in modulating lysine multiple meth-
ylation by SET domain KMTs and provide the first molecular
snapshots of the mono-, di-, and trimethyl transfer reactions
catalyzed by these enzymes.
SET domain enzymes represent a family of S-adenosylmethi-
onine (AdoMet)3-dependent methyltransferases that catalyze
the site-specific methylation of protein lysyl residues in a host
of proteins, including histones, transcription factors, chroma-
tin-modifying enzymes, ribosomal subunits, and other sub-
strates (1–3). In many instances, these modifications serve to
recruit effector proteins that recognize methyl-lysyl residues in
a sequence-dependent fashion (4). In addition, SET domain
KMTs exhibit product specificity, defined as their ability to cat-
alyze mono-, di-, or trimethylation of the lysine -amino group.
This specificity is biologically relevant because many methyl-
lysine-binding proteins can discriminate among different
degrees of lysine methylation (4). Thus, both the site and degree
of lysine methylation are critical to recognition by effector
proteins.
Structural and functional studies have identified a Phe/Tyr
switch in the active site of SET domain KMTs that governs their
respective product specificities (5, 6). According to this model,
KMTs that possess a tyrosine in the Phe/Tyr switch site are
limited to catalyzing lysine monomethylation, whereas en-
zymes that possess a phenylalanine or another hydrophobic
residue in this position display di- or trimethyltransferase activ-
ity. Mutational analysis of various SET domain KMTs, includ-
ing DIM-5 (KMT1) (6), SET7/9 (6), G9A (KMT1C) (5), and
SET8 (also known as PR-SET7 and KMT5A) (7, 8), has demon-
strated that substitutions in the Phe/Tyr switch result in pre-
dictable changes in product specificity. Several models have
been proposed to explain the mechanism by which the Phe/Tyr
switch site governs this specificity, including variations in the
diameter of the active site due to the size of Phe/Tyr switch
residue and steric hindrance by the tyrosine hydroxyl group (6,
9–11). However, our recent studies of the Phe/Tyr switch
mutant Y334F in the human histone H4 Lys-20 (H4K20) meth-
yltransferase SET8 indicate that the Phe/Tyr switch regulates
product specificity via a more subtle mechanism (8). Specifi-
cally, the switch modulates the binding of an active site water
molecule that in turn regulates the transition from mono-
methylation to multiple methylation.
Among the KMTs that have been structurally characterized,
SET7/9 has emerged as an archetypal model for studying the
catalytic mechanism and product specificity of the SET domain
family. Although initially isolated as a histone H3 Lys-4
(H3K4)-specific methyltransferase, this KMT has been shown
to regulate the functions of numerous non-histone substrates
through site-specific methylation (12–21). Early structural and
functional studies of SET7/9 identified two active site mutants,
* Thisworkwassupported,inwholeorinpart,byNationalInstitutesofHealth
Grant R01 GM073839 (to R. C. T.) and National Institutes of Health Admin-
istrative Supplement GM073839-04S1 (to R. C. T.) funded through the
American Recovery and Reinvestment Act. This work was also supported
by Department of Energy Grant DE-FG02-92ER20075 (to R. L. H.).
□
S The on-line version of this article (available at http://www.jbc.org) contains
supplemental Table 1.
The atomic coordinates and structure factors (codes 3M53, 3M54, 3M55, 3M56,
3M57,3M58,3M59,and3M5A)havebeendepositedintheProteinDataBank,
Research Collaboratory for Structural Bioinformatics, Rutgers University, New
Brunswick, NJ (http://www.rcsb.org/).
1 Supported by a Canadian Institutes of Health Research postdoctoral
fellowship.
2 To whom correspondence should be addressed: Dept. of Biological Chem-
istry, University of Michigan Medical School, 1150 West Medical Center Dr.,
5301 MSRB III, Ann Arbor, MI 48109. Tel.: 734-647-0889; Fax: 734-763-4581;
E-mail: rtrievel@umich.edu.
3 The abbreviations used are: AdoMet, S-adenosylmethionine; AdoHcy, S-ad-
enosylhomocysteine; KMT, lysine methyltransferase; ITC, isothermal titra-
tion calorimetry; CH–O, carbon-oxygen hydrogen bond; BisTris, 2-[bis(2-
hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 41, pp. 31849–31858, October 8, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31849
Y245A and Y305F, which change its product specificity. The
Phe/Tyr switch mutant Y305F alters SET7/9 product specificity
from a mono- to dimethyltransferase (6), whereas the Y245A
substitution converts the enzyme into a trimethyltransferase
with weak monomethyltransferase activity (11). These mutants
have been the subjects of numerous molecular modeling simu-
lations that have led to various models to explain their distinct
product specificities (22–26). However, the lack of structural
data for the SET7/9 Y245A and Y305F mutants in complex with
cognate methylated peptides has hindered our understanding
of the mechanisms that define the respective product specific-
ities of these mutants. Moreover, these structures would yield a
framework for visualizing the mono-, di-, and trimethylation
reactions catalyzed by SET domain KMTs.
To gain insight into the molecular basis of their product
specificities, we have determined high resolution crystal
structures of the SET7/9 Y245A and Y305F mutants in com-
plex with peptides of the TATA box-binding protein-associ-
ated factor TAF10 bearing the Lys-189 methylation site in
unmodified (K189), monomethylated (K189me1), dimethyl-
ated (K189me2), and trimethylated (K189me3) states. The
structures and accompanying biochemical data support a
model whereby changes in the occupancy or position of water
molecules in the active site are critical in establishing the prod-
uct specificities of the SET7/9 Y245A and Y305F mutants.
Together, our results provide new insights into the mechanisms
that govern SET domain product specificity and provide step-
wise snapshots of the lysine mono-, di-, and trimethyl transfer
reactions catalyzed by KMTs.
EXPERIMENTAL PROCEDURES
Cloning, Expression, and Purification of the SET7/9 Mutants—
The Y245A and Y305F mutants were introduced into the pHIS2
SET7/9 expression vector encoding residues 110–366 (27) via
QuikChange site-directed mutagenesis (Stratagene) and were
verified by dideoxy DNA sequencing. The plasmids encoding
wild type (WT) SET7/9 and the Y245A and Y305F mutants
were transformed into Rosetta2 DE3 cells (Novagen) and were
expressed as described previously (27, 28). In the course of
characterizing WT SET7/9, we observed that the enzyme co-
purified with AdoMet or another contaminant that resulted in
technical difficulties in the isothermal titration calorimetry
(ITC) experiments and co-crystallization trials with the TAF10
peptides. To overcome this problem, a denaturation and refold-
ing step was inserted in the purification scheme. The denatur-
ation and refolding protocol involved adding 6 M guanidine
HCl, 50 mM sodium phosphate, pH 7.5, and 500 mM NaCl to the
protein while it was immobilized on a nickel-Sepharose column
(GE Healthcare). The column was washed with this buffer, fol-
lowed by a solution of 6 M urea, 50 mM sodium phosphate, pH
7.5, 150 mM NaCl, and 10 mM 2-mercaptoethanol to remove the
cofactor from the denatured enzyme. A reverse gradient from 6
to 0 M urea was then performed in the same buffer to refold the
protein, which was subsequently eluted from the column using
a linear gradient of 0–500 mM imidazole in 50 mM sodium
phosphate, pH 7.5, 500 NaCl, and 10 mM 2-mercaptoethanol.
The refolded protein was digested with tobacco etch virus pro-
tease (29) during dialysis against 20 mM Tris, pH 7.5, 150 mM
NaCl, and 5 mM 2-mercaptoethanol and then purified using a
Superdex 200 gel filtration column (GE Healthcare). Protein
concentration was determined by its absorbance at 280 nm.
Synthetic Peptides—The TAF10 peptides bearing K189,
K189me1,
K189me2,
and
K189me3
(sequence,
acetyl-
SKSK189DRKYTL-amide) and a biotinylated TAF10 peptide
(sequence,
acetyl-SKSK189DRKYTLT(K-EZLink-S-S-biotin)-
amide) were synthesized and purified by New England Peptide,
Inc. Peptide concentrations were measured using the absorb-
ance of their tyrosine residue at 274 nm.
Crystallization and Data Collection—Crystals were pro-
duced by hanging drop vapor diffusion by mixing the crystalli-
zation solution in a 1:1 ratio with 10–20 mg/ml SET7/9, 1 mM
S-adenosylhomocysteine (AdoHcy), and 1.0–3.0 mM unmodi-
fied or methylated TAF10-K189 peptide in 20 mM Tris, pH 8.0,
100 mM NaCl, and 2 mM tris(2-carboxyethyl)phosphine. Crys-
tals were obtained at 20 °C in either 1.8–2.0 M (NH4)2SO4, with
0.1 M BisTris, pH 6.2–6.6, or in 0.95–1.1 M sodium citrate with
100 mM imidazole pH 8.0–8.4. In both crystallization condi-
tions, the final pH values were between pH 8.0 and 9.0. Crystals
in the (NH4)2SO4 condition were typically flash-frozen in the
mother liquor containing 25–30% glycerol, and the crystals in
the citrate condition were frozen in 1.6 M sodium citrate. Data
were collected at the Advanced Photon Source beamlines
21-IDG (LS-CAT) and 23-IDD (GM/CA-CAT). Images were
indexed, integrated, and scaled using HKL2000 (30). Structures
of the mutants were solved by molecular replacement using
MOLREP (31) with the coordinates of a previously reported
SET7/9 ternary complex used as the search model (Protein
Data Bank code 2F69). Successive rounds of model building and
refinement were carried out using Coot (32) and REFMAC (33),
respectively. The geometry of the models were verified by Mol-
Probity (34). Simulated annealing omit maps were calculated
using CNS (35) with the peptide and cofactor removed to elim-
inate model bias in the active site. Structural figures were ren-
dered using PyMOL (Schro¨dinger, LLC.).
Fluorescent Methyltransferase Assay—A coupled fluorescent
methyltransferase assay was used to measure the kinetic
parameters of WT SET7/9 and the Y245A and Y305F mutants
as reported previously, with the exception that 50–150 nM
enzyme, 100 M AdoMet, and varying concentrations of TAF10
peptide substrate were used (27, 36). Assays were performed in
triplicate, and a homocysteine calibration curve was used to
calculate the initial velocities. Kinetic parameters were calcu-
lated by plotting the velocities versus peptide concentration and
by fitting the Michaelis-Menten equation to the data via non-
linear regression using Prism 5.0 (GraphPad). In cases where
the Km value was beyond the measurable range of the assay, the
kcat/Km value was determined as described previously (7).
Calorimetry Experiments—ITC was performed at 20 °C using
a MicroCal VP-ITC calorimeter (GE Healthcare) with 0.12 mM
protein and 4 mM AdoHcy in 20 mM sodium phosphate, pH 7,
and 100 mM NaCl with 1.5 mM peptide as the injectant. Data
were processed, and equilibrium dissociation constants (KD)
and curve fitting errors were calculated from the binding iso-
therms using Origin 7.0 (OriginLab Corp.). WT SET7/9 and the
Y245A and Y305F mutants displayed ligand:protein binding
stoichiometries (N values) between 0.8 and 1.0, demonstrating
Lysine Methylation by SET7/9 Mutants
31850
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
that WT SET7/9 and its mutants were properly refolded due to
their ability to bind peptides in an 1:1 molar ratio.
TLC Product Analysis—Methyltransferase assays were per-
formed in triplicate at 37 °C with the biotinylated TAF10 pep-
tide (0.5 mM) for 30 min in the presence of WT SET7/9 (3
pmol), Y305F mutant (6 pmol), or the Y245A mutant (100
pmol). Assays contained 100 mM HEPES, pH 8.0, 50 mM NaCl,
1.0 mM [methyl-3H]AdoMet (diluted to a specific activity of 0.2
Ci/nmol with purified AdoMet (37)), 15 M Sulfolobus solfa-
taricus AdoHcy hydrolase (36), and 2 units of adenosine deami-
nase (Roche Applied Science) in a final volume of 20 l. The
reactions were terminated by addition of an equal volume of
200 mM MES, pH 5.1, prior to addition of a 2-fold molar excess
of immobilized avidin resin (UltraLink; Pierce). Biotinylated
peptides were allowed to bind at room temperature for 30 min,
and the resin was then collected by centrifugation (9000 g).
The resin was washed three times with 300 mM NaCl, and the
peptide was eluted overnight from the avidin resin by cleavage
of the disulfide bond in the linker of the peptide using 10 mM
tris(2-carboxyethyl)phosphine (Thermo Scientific). The resin
was incubated with additional 10 mM tris(2-carboxyethyl)phos-
phine the following day until the radiolabel was essentially
removed from the resin. The recovered peptides were hydro-
lyzed by incubation with 6 M HCl at 110 °C for 24 h. Subsequent
steps in measuring the radiolabel incorporated into the mono-,
di-, and trimethyl-lysine products were performed as reported
previously (8).
RESULTS
Functional Analysis of the SET7/9
Y305F Mutant—Prior studies of
SET7/9 by Zhang et al. (6) reported
that mutation of the Phe/Tyr switch
residue Tyr-305 to a phenylalanine
alters its product specificity from a
mono- to dimethyltransferase. We
verified these findings by demon-
strating that WT SET7/9 mono-
methylated the TAF10-K189 pep-
tide, whereas the Y305F mutant
mono- and dimethylated this sub-
strate, as demonstrated by mass
spectrometry (data not shown). We
next examined whether the Y305F
substitution altered the affinity of SET7/9 for the TAF10-K189
peptides using ITC (Fig. 1). A comparison of the KD values
revealed that SET7/9 Y305F bound the TAF10-K189 and
TAF10-K189me1 peptides 4- and 6-fold more tightly, respec-
tively, than the WT enzyme, whereas this mutant displayed a
substantially diminished affinity for the TAF10-K189me2 pep-
tide (Table 1). Although the WT enzyme and the Y305F mutant
exhibited discernable differences in their affinities for the
unmodified and monomethylated peptides, these variations are
modest and cannot account for their distinct product specific-
ities, suggesting that a kinetic effect during methylation may be
responsible.
To investigate this possibility, we characterized the kinetic
parameters of WT SET7/9 and the Y305F mutant using the
TAF10 peptides as substrates. Both enzymes methylated the
unmodified peptide with comparable kcat and Km values (Table
2). In analyzing the kinetic parameters for the methylation
of the monomethylated peptide by SET7/9 Y305F, we found
that this substrate displayed an elevated Km value that was
beyond the measurable range of the assay due to its limited
solubility. In this case, we measured the catalytic efficiency
(kcat/Km) for the methylation of this peptide and found that it
was methylated 15-fold less efficiently than the unmodified
peptide by SET7/9 Y305F. Given the fact that the Y305F mutant
exhibited a higher binding affinity for the TAF10-K189me1
peptide than the WT enzyme (Table 1), the kinetic data suggest
that a step in the reaction pathway following substrate binding
limits the catalytic efficiency of this mutant.
We next examined whether the Y305F mutant dimethylated
the TAF10-K189 peptide via a processive or a distributive
mechanism. In a processive mechanism, the methyl-lysine
substrate would remain bound to the enzyme during successive
methyl transfer reactions; thus, the concentration of an inter-
mediate, such as monomethyl-lysine, cannot exceed the en-
zyme concentration during the assay. In a distributive mecha-
nism, the intermediates are released into solution where they
accumulate prior to the next round of methylation, resulting in
an intermediate concentration that is greater than that of the
enzyme. Using a radiometric TLC assay and a biotinylated
TAF10 peptide, we quantified the amounts of monomethylated
products generated by the WT SET7/9 and the Y305F mutant
FIGURE 1. ITC analysis of WT SET7/9 and the Y245A and Y305F mutants. Representative titrations and
binding isotherms for the calorimetry experiments are shown for the unmodified TAF10-K189 peptide titrated
into WT SET7/9 (left plot), SET7/9 Y305F (middle plot), and SET7/9 Y245A (right plot). In each panel, the ITC
titration experiment (upper panel) is shown with the binding isotherms (lower panel) fit to a single binding site
model.
TABLE 1
Analysis of the binding affinity of WT SET7/9 and its catalytic
mutants for unmodified and methylated TAF10 peptides
SET7/9
TAF10 peptide
KD
a
M
WT
K189
4.9 0.20
WT
K189me1
4.0 0.36
Y305F
K189
1.3 0.10
Y305F
K189me1
0.62 0.065
Y305Fb
K189me2
70
Y245A
K189
4.0 0.25
Y245A
K189me1
3.3 0.10
Y245A
K189me2
5.8 0.22
Y245A
K189me3
11 0.28
a Curve fitting errors were calculated from the binding isotherms.
b An estimate of the affinity is reported due to weak peptide binding.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31851
(Table 3). The data demonstrate that comparable amounts of
monomethyl-lysine were generated when the quantity of
enzyme usedistakenintoaccount,inagreementwiththeirsimilar
turnover numbers for the TAF10-K189 peptide (Table 2). The
Y305F mutant also produced small but measurable quantities of
radiolabeled dimethyl-lysine product that were substantially
smaller than the amount of monomethyl-lysine generated.
Therefore, the TLC data are consistent with a distributive
mechanism for dimethylation by the Y305F mutant because the
amount of monomethyl-lysine produced exceeded the quantity
of enzyme used in the assay.
Structures of WT SET7/9 and the Y305F Mutant in Complex
with Unmodified and Methylated TAF10 Peptides—To deter-
mine the mechanism by which the Y305F substitution alters the
product specificity of SET7/9, we determined the crystal struc-
tures of this mutant bound to AdoHcy and TAF10-K189,
TAF10-K189me1, and TAF10-K189me2 peptides and com-
pared these to the structures of the WT SET7/9AdoHcy
TAF10-K189 complex (supplemental Table 1). The structures
of these complexes were determined to 1.85 Å or higher reso-
lution, permitting unambiguous modeling of the K189 side
chains in the active site of the enzyme based on simulated
annealing omit maps (Fig. 2). The ternary complexes of the WT
and the Y305F mutant superimpose with overall root mean
square differences of less than 0.3 Å for all aligned atoms, indi-
cating that neither the Y305F mutation nor the binding of the
various TAF10-K189 peptides results in substantial changes in
its overall structure.
An inspection of the active sites of the SET7/9 WT and
Y305F complexes illustrates the binding modes of the unmod-
ified and methylated forms of K189 in the TAF10 peptides (Fig.
2, A–D). The K189 side chain binds in an extended all trans
conformation in a deep pocket, termed the lysine binding chan-
nel, that is composed of Tyr-245, Thr-266, Leu-267, Ser-268,
Tyr-305, Tyr-335, and Tyr-337 (not shown for clarity) (Fig. 2A).
These residues interact with the aliphatic portion of the K189
side chain primarily through van der Waals contacts. The lysine
binding channel connects to the AdoMet-binding site on the
opposite face of the catalytic domain via an oxygen-lined
methyl transfer pore (38). During catalysis, the methyl group of
the cofactor is positioned within the methyl transfer pore
for the SN2 reaction with the -amino group of the lysine or
methyl-lysine substrate (see below).
To lower the activation barrier for this reaction, the lysine
-amine nucleophile is aligned for methyl transfer through a
hydrogen bond network within the active site. In the WT
enzyme, the K189 -amino group hydrogen bonds to the
hydroxyl group of Tyr-245 as well as to two water molecules
(Fig. 2A). One of the water molecules (termed water 1), is coor-
dinated in a solvent pocket, through hydrogen bonds to the
carbonyl oxygens of Gly-292 and Ala-295 and to the hydroxyl
group of the Phe/Tyr switch residue Tyr-305. This solvent
pocket is structurally conserved in SET domain KMTs and has
an important role in defining product specificity through the
adjacent Phe/Tyr switch residue, as shown in our prior studies
of the human H4K20 methyltransferase SET8 (8). The other
water molecule is bound within the methyl transfer pore
between the lysine substrate and the thioether sulfur atom of
AdoHcy through hydrogen bonds to Tyr-245, Asn-265, and
His-293 in SET7/9 and the TAF10-K189 -amino group. This
water is not observed in other structures of SET7/9 ternary
complexes and may represent the approximate position that
the AdoMet methyl group occupies in the methyl transfer pore
in the Michaelis complex.
In structures of the Y305F ternary complexes, the K189,
K189me1, and K189me2 side chains also adopt extended trans
side chain geometries within the lysine binding channel that are
stabilized via hydrogen bonding to Tyr-245 in the enzyme (Fig.
2, B–D). The orientations of the K189me1 and K189me2 side
chains are further maintained through carbon-oxygen (CH–O)
hydrogen bonding between the methyl groups and oxygen
atoms within the vicinity of the methyl transfer pore, as
reported previously in other SET domain KMT structures (8,
10, 38). A superimposition of the SET7/9 WT and Y305F com-
plexes underscores the similarity of the lysyl binding conforma-
tions (Fig. 2E). However, there are notable differences in the
hydrogen bond patterns and occupancy of water 1 within the
solvent pocket in the Y305F mutant compared with the WT
enzyme. Specifically, the Y305F substitution results in the loss
of one hydrogen bond to water 1 in the structures of the TAF10-
K189 and TAF10-K189me1 complexes (Fig. 2, B and C). In con-
trast, water 1 is absent in TAF10-K189me2 complex, and the
vacated solvent pocket is occupied by one of the methyl groups
TABLE 2
Kinetic parameters of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
TAF10 peptide substrate
Km
a
kcat
a
kcat/Km
a
M
min1
M1 min1 103
WT
K189
160 17
17 0.62
110 17
Y305F
K189
88 5.0
17 0.30
190 11
Y305Fb
K189me1
11 0.50
Y245A
K189
200 35
0.53 0.04
2.6 0.47
Y245A
K189me1
210 23
5.9 0.23
28 3.3
Y245A
K189me2
400 29
6.5 0.16
15 1.2
a Curve fitting errors were calculated from the hyperbolic fits for the Michaelis-Menten equation.
b Km and kcat were not determined because the Km value was beyond the measurable range; therefore, the kcat/Km value is reported.
TABLE 3
Product analysis of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
Quantity
of enzyme
Measured
product
Amount of
product formeda
nmol
nmol
WT
0.003
Kme1
0.65 0.07
Y305F
0.006
Kme1
1.5 0.49
Kme2
0.033 0.009
Y245A
0.100
Kme1
0.80 0.22
Kme2
0.39 0.021
Kme3
0.076 0.019
a Standard deviation was calculated from triplicate measurements.
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of the dimethyl -amine (Fig. 2D). This methyl group forms a
3.5-Å CH–O hydrogen bond to the carbonyl oxygen of Ala-295,
further stabilizing the binding of the dimethyl-lysine side chain.
A homologous dimethyl-lysine-binding mode and CH–O
hydrogen bond was observed in our prior structural studies of
the SET8 Y334F Phe/Tyr switch mutant that confers an analo-
gous change in product specificity from a mono- to a dimeth-
yltransferase (8). A structural alignment of the active sites of the
SET7/9 Y305F and SET8 Y334F mutants bound to cognate
dimethylated peptides illustrates that the coordinates of the
dimethyl-lysyl side chains are virtually superimposable, with
one methyl group oriented toward the methyltransfer pore and
the second positioned within the vacant solvent pocket (Fig.
2F). Taken together, the structures of the SET7/9 Y305F com-
plexes and the similarities in the dimethyl-lysine conforma-
tions in the SET7/9 Y305F and SET8 Y334F mutants imply that
the Phe/Tyr switch governs product specificity through a con-
served mechanism whereby it indirectly influences the binding
modes of the methyl-lysine side chain by modulating the affin-
ity of the water molecule (water 1) bound in the solvent pocket.
Biochemical Characterization of
the SET7/9 Y245A Mutant—Previ-
ous studies by Xiao et al. (11)
reported that the Y245A mutation
yields an unusual change in the
product
specificity
of
SET7/9,
converting the enzyme to a trimeth-
yltransferase with weak monometh-
yltransferase activity. We deter-
mined that the SET7/9 Y245A could
mono-, di-, and trimethylate the
TAF10-K189 peptide by mass spec-
trometry (data not shown) and TLC
(Table 3), confirming the earlier
studies of Xiao et al. (11). ITC
analysis revealed that the Y245A
mutant displayed comparable KD
values
for
the
unmodified
and
methylated TAF10-K189 peptides
(Fig. 1), although its affinity for the
trimethylated peptide was modestly
diminished in comparison with the
other peptides (Table 1). The ITC
data demonstrate that the Y245A
mutant
bound
the
unmodified,
mono-,
and
dimethylated
sub-
strates with equivalent affinities,
suggesting that a kinetic effect or
a structural alteration in the active
site may be responsible for its
diminished
activity
toward
un-
modified substrates.
To gain further insight into its
peculiar
product
specificity,
we
characterized the kinetic properties
of
the
SET7/9
Y245A
mutant.
Steady state analysis demonstrated
that this mutant displayed similar
Km values for the unmodified, mono- and dimethylated TAF10
peptides (Table 2). However, the turnover number for the
TAF10-K189 peptide was diminished over 10-fold versus the
methylated peptides and was reduced 30-fold versus the WT
enzyme, in agreement with the weak monomethyltransferase
activity reported by Xiao et al. (11). In addition, we investigated
whether this mutant catalyzes lysine trimethylation via a pro-
cessive or distributive mechanism as described for SET7/9
Y305F. The TLC data illustrate that the mono- and dimethyl-
lysine intermediates accumulated at quantities greater than
that of the enzyme used in the assay, indicating that SET7/9
Y245A obeys a distributive mechanism, analogous to the Y305F
mutant (Table 3).
Structures of SET7/9 Y245A Bound to Unmodified and Meth-
ylated TAF10 Peptides—To elucidate the mechanism underly-
ing its unusual product specificity, we determined the crystal
structures of SET7/9 Y245A in complex with AdoHcy and
unmodified, mono-, di-, and trimethylated TAF10 peptides
(supplemental Table 1). These complexes superimpose with
the structure of the WT SET7/9AdoHcyTAF10-K189 com-
FIGURE 2. Crystal structures of WT SET7/9 and the Y305F mutant in complex with AdoHcy and unmodi-
fied and methylated TAF10 peptides. A, active site of WT SET7/9 bound to AdoHcy and TAF10-K189 and the
active site of SET7/9 Y305F bound to AdoHcy and TAF10-K189 (B), TAF10-K189me1 (C), and TAF10-K189me2
peptides (D). A–D, the K189 residue in the TAF10 peptide is depicted with yellow carbon atoms with the
corresponding Fo – Fc simulated-annealing omit maps (contoured at 2.5 ) shown. AdoHcy is colored with
green carbon atoms, and SET7/9 carbon atoms are colored gray, with the exception of Tyr-305/Y305F
(magenta) and Tyr-245 (cyan). Gly-264 to Ser-268 are shown, and the main chain atoms of Gly-292 to Ala-295
are depicted for clarity. Red dashed lines indicate conventional hydrogen bonds, and blue dashed lines indicate
CH–O hydrogen bonds. Cyan dashed lines indicate weak CH–O hydrogen bonds between 3.5 and 3.7 Å in
length. The water molecule in the solvent binding pocket is numbered 1. E, overlay of the active sites of the WT
enzyme (light blue carbon atoms) and the Y305F mutant (green, yellow, and pink carbon atoms correspond to
K189, K189me1, and K189me2, respectively). Red dashed lines indicate coordination of the waters in the struc-
turesoftheY305Fmutant,andthelightbluedashedlineindicateshydrogenbondingthatonlyoccursintheWT
enzyme. F, superimposition of the active sites of the SET7/9 Y305FAdoHcyTAF10-K189me2 and SET8
Y334FAdoHcyH4K20me2 (Protein Data Bank code 3F9X) complexes shown with green and yellow carbon
atoms, respectively.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31853
plex with root mean squared differences of less than 0.4 Å for all
aligned atoms, indicating that the Y245A mutant does not per-
turb the overall structure of the enzyme. Simulated annealing
omit maps illustrate that K189 side chains are bound within the
lysine binding channel through hydrogen bonds and van der
Waals contacts (Fig. 3, A–D), although the interactions and
binding modes are distinct from those in the complexes of WT
SET7/9 and the Y305F mutant (Fig. 2, A–D). In the unmodified
TAF10 peptide complex, the K189 -amino group forms a weak
hydrogen bond with the carbonyl oxygen of Gly-264 (Fig. 3A),
whereas the -amino groups of K189me1 and K189me2 hydro-
gen bond to the hydroxyl group of Tyr-305 in the mono- and
dimethylated peptide complexes (Fig. 3, B and C). The confor-
mations of the K189me1 and K189me2 side chains are further
stabilized by water-mediated hydrogen bonding and through
CH–O hydrogen bonding to their methyl groups. In the
TAF10-K189me3 peptide complex, the trimethyl-lysine side
chain is coordinated exclusively through direct and water-me-
diated CH–O hydrogen bonds to its methyl groups because the
quaternary -ammonium cation cannot engage in hydrogen
bonding (Fig. 3D).
A structural alignment of the four SET7/9 Y245A complexes
illustrates distinct binding modes for the unmodified versus the
methylated K189 side chains, highlighting the selectivity of this
mutant for methylated substrates. The side chains of K189me1,
K189me2, and K189me3 roughly overlay with their respective
-amino groups superimposed and adopt slightly kinked con-
formations (Fig. 3E), as opposed to the extended trans geome-
try of the unmodified and methylated lysines in the complexes
of the WT enzyme and the Y305F mutant (Fig. 2, A–D). Con-
versely, the unmodified K189 side chain does not superimpose
with its methylated counterparts and is oriented in an alterna-
tive configuration due to its hydrogen bonding to Gly-264 (Fig.
3, A and E). An overlay of the structures of the WT enzyme and
Y245A mutant bound to the unmodified TAF10 peptide illus-
trates that the side chains of K189 do not superimpose and that
the K189 -amino group appears to be misaligned with AdoHcy
in the Y245A complex (Fig. 3F). This suboptimal alignment
may explain the diminished kcat value of SET7/9 Y245A mutant
toward substrates with unmodified lysines (Table 2).
A comparison of the structures of the SET7/9 Y245A and
Y305F complexes yields a molecular explanation for the differ-
ent product specificities of these two mutants. In the SET7/9
Y305F complexes, Tyr-245 aligns the K189 -amino group for
methyl transfer through hydrogen bonding to its hydroxyl
group (Fig. 2, B–D). Conversely, in the Y245A mutant, the
K189me1 and K189me2 -amino groups are oriented through
hydrogen bonding to Tyr-305 (Fig. 3, B and C). These distinct
hydrogen bond patterns impart differences in the conforma-
tions of the lysyl side chains due to the relative orientations of
Tyr-245 and Tyr-305 in the lysine binding channel. Specifically,
the kinked conformation adopted by the K189me1 and
K189me2 side chains in the Y245A complexes (Fig. 3, B and C)
may contribute to the differences in the turnover numbers of
this mutant versus those of the WT enzyme and the Y305F
mutant (Table 2). In addition, the dimethyl -amino group of
the K189me2 side chain binds in distinct orientations in the
Y245A and Y305F mutants due to their hydrogen bonding to
FIGURE 3. Crystal structures of the SET7/9 Y245A mutant in complex with
AdoHcy and unmodified and methylated TAF10 peptides. Active site of
SET7/9 Y245A bound to AdoHcy and TAF10-K189 (A), TAF10-K189me1 (B),
TAF10-K189me2 (C), and TAF10-K189me3 peptides (D). A–D, Fo – Fc simulated-
annealing omit maps (contoured at 2.5 ) for the unmodified and methylated
K189 side chains are illustrated. The residues and hydrogen bonds in each com-
plexarecoloredasinFig.2.Thewatermoleculesinthelysinebindingchannelsof
the Y245A complexes are numbered 1–4, as described in the text. E, superimpo-
sition of the active sites of the Y245A complexes bound to the four methylated
statesofTAF10-K189(unmodified,mono-,di-,andtrimethylatedshowningreen,
yellow, pink, and blue, respectively). F, overlay of WT SET7/9 (blue carbons) and
SET7/9 Y245A (green carbons) in complex with TAF10-K189. Waters correspond-
ing to the WT and Y245A structures are colored cyan and green, respectively.
G, alignment of the active sites of the Y245A (yellow carbons) and Y305F (green
carbons) mutants in complex with the TAF10-K189me2 peptide. Hydrogen
bonds from the Y305F structure are shown as green dashed lines, and waters and
hydrogen bonds in the Y245A structure are shown in yellow and orange,
respectively.
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Tyr-305 and Tyr-245, respectively (Fig. 3G). In the Y305F
mutant, hydrogen bonds to the dimethyl -amino group cou-
pled with steric constraints in the lysine binding channel pre-
vent the K189me2 side chain from undergoing a conforma-
tional change that is conducive to trimethylation (Fig. 2D),
consistent with its dimethyltransferase activity. However, in the
Y245A mutant, the alanine substitution enlarges the diameter
of the lysine binding channel, accommodating trimethyl-lysine
(Fig. 3D). In addition, the larger diameter would permit the
dimethyl-lysine substrate to undergo the conformational reor-
ganization necessary to align the -amino group in a productive
geometry for trimethylation.
A major difference in the active site of the Y245A mutant
versus the other SET7/9 structures is the presence of several
water molecules bound in the cavity generated by the Y245A
mutation. In the structure of the Y245A mutant bound to
TAF10-K189, three water molecules (waters 2–4) occupy this
cavity and are arranged in a triangular geometry (Fig. 3A). In
addition, water 1 shifts 1.6 Å from its position in the solvent
pocket toward water 2 to which it forms a hydrogen bond (Fig.
3, A and E). The shift in water 1 was unexpected given its con-
served orientation in the solvent pocket of the SET7/9 WT and
Y305F complexes (Fig. 2, A–C) as well as in the structures of
other SET domain KMTs (8). This displacement is presumably
related to the alternative conformation of the K189 side chain
whose -amino group is too distant (4.3 Å) to form a productive
hydrogen bond to water 1. Conversely, in the Y245A complexes
bound to TAF10-K189me1 and TAF10-K189me2, water 1
remains tightly bound in the solvent pocket through hydrogen
bonds to Tyr-305 hydroxyl group, the carbonyl oxygens of Gly-
292 and Ala-295, and the K189 -amino group (Fig. 3, B, C, and
E), analogous to its binding in the WT enzyme (Fig. 2A). How-
ever, in the TAF10-K189me3 complex, one of the methyl
groups of the trimethyl -ammonium cation is oriented into the
solvent pocket (Fig. 3D), similar to the dimethyl-lysine binding
mode observed in the Y305F mutant (Fig. 2D). The binding of
the methyl group in the solvent pocket displaces water 1 by 3.2
Å relative to its position in the TAF10-K189me1 complex (Fig.
3E), thereby avoiding a steric clash with the trimethylated
-ammonium group. Variations in the occupancy of water 2 are
also seen in the different Y245A structures. Water 2 is bound in
similar orientations in the active site of the unmodified and
monomethylated peptide complexes but is absent in the di- and
trimethylated peptide complexes due to the binding of a methyl
group in this position (Fig. 3, A–E). In summary, the changes in
the positions or occupancies of waters 1 and 2 correlate with the
binding modes of the unmodified and methylated K189 within
the active site of the Y245A mutant.
Catalytic Models of Lysine Multiple Methylation by SET7/9
Y245A, and Y305F—The structures of the SET7/9 complexes
reported here offer a prime opportunity to generate stepwise
models for lysine mono-, di-, and trimethylation by a SET
domain KMT. We modeled the AdoMet-bound Michaelis
complexes by superimposing the SET7/9 product complexes
with the previously reported structure of the SET7/9-AdoMet
binary complex (Fig. 4) (39). The conformations of the mono-
and dimethyl -amino groups in the Michaelis complexes were
inferred from the coordinates of the corresponding dimethyl-
and trimethyl-lysine products, respectively. In addition, we
modeled the -amino group in a deprotonated state with its
hydrogen atoms oriented toward the hydrogen bond acceptors
that align the lysyl side chain for methylation. As a basis for this
comparison, we first modeled the monomethylation reaction
catalyzed by WT SET7/9 (Fig. 4A). In the substrate ternary
complex, the lysine -amine is aligned with the methyl group
and sulfonium cation of AdoMet through a hydrogen bond to
the Tyr-245 hydroxyl group and water 1 in the solvent pocket.
The values of the reaction distance and angle are 2.8 Å and 153°,
respectively, in approximate agreement with the linear geome-
try of a SN2 methyl transfer reaction calculated in other mod-
eled substrate complexes (8, 10). In the product complex, the
monomethyl-lysine side chain is bound in an extended confor-
mation with its methyl group oriented within the methyl trans-
fer pore, thereby obstructing AdoMet binding. Furthermore,
water 1 remains tightly coordinated in the solvent pocket
through four hydrogen bonds to Gly-292, Ala-295, Tyr-305,
and the monomethyl -amino group. These interactions hinder
the dissociation of water 1 and the related rearrangement of the
monomethyl-lysine side chain required for a second methyl
transfer reaction, explaining why the WT enzyme cannot cata-
lyze di- and trimethylation. These findings concur with the
FIGURE 4. Catalytic models of the methyl transfer reactions catalyzed by
WT SET7/9 and the Y305F mutant. A, monomethylation of TAF10-K189 by
the WT enzyme. The reaction scheme depicts the modeled substrate ternary
complex (left) and the product complex (right) for the transfer of the methyl
group from AdoMet (green carbon atoms) to K189 in TAF10 (yellow carbons),
yielding AdoHcy and K189me1. The red arrow indicates the direction of the
nucleophilic attack of the deprotonated -amino group on the AdoMet
methyl group. The transferred methyl group is colored green, and the white
atoms represent the hydrogens of the -amino group. Hydrogen bonds
and residues in the enzyme active site are illustrated as in Fig. 2. The
reaction distance and angle are labeled in red. B and C, models of the
Y305F mutant for the first methyl transfer reaction with TAF10-K189 (B)
and second methyl transfer reaction with TAF10-K189me1 (C). Color
schemes are the same as in A.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY 31855
model for SET7/9 product specificity reported in previous
structural and functional studies (6, 11).
Similar reaction geometry is observed in the model for the
monomethyl transfer reaction catalyzed by SET7/9 Y305F.
Hydrogen bonds from the Tyr-245 hydroxyl group and water 1
align the lysine -amino group with the AdoMet methyl group
at a distance of 2.1 Å and an angle of 160°, equivalent to those
measured in the Michaelis complex of the WT enzyme (Fig. 4, A
and B). In the product complex, monomethyl-lysine adopts an
extended trans configuration analogous to that in the WT
enzyme. For dimethylation to occur, the monomethyl-lysine
must undergo a conformational change in which its methyl
group is rotated out of the methyl transfer path with AdoMet.
The structure of the Y305F mutant bound to the dimethylated
TAF10 peptide (Fig. 2D) implies that this rearrangement occurs
through the dissociation of water 1 due to the loss of the Tyr-
305 hydrogen bond in the solvent pocket. The dissociation of
water 1 would enable the monomethyl-lysine side chain to
adopt an alternative conformation through a rotation about its
C–N bond, projecting the methyl group into the solvent
pocket (Fig. 4C). This rotation reorients the methyl group out of
the methyl transfer path while realigning the monomethyl -
amino group for a second methylation reaction through a direct
hydrogen bond to the Tyr-245 hydroxyl group and a CH–O
hydrogen bond between its methyl group and Ala-295. The
modeled reaction geometry for monomethyl-lysine substrate
complex of the Y305F mutant (2.7 Å, 160°) is equivalent to that
of the first methyl transfer reaction in SET7/9 Y305F. These
geometries concur with our previous models for mono- and
dimethylation catalyzed by SET8 Y334F (8), illustrating that the
orientation of a methyl group into the solvent pocket is a con-
served feature of SET domain KMTs that catalyze multiple
methylation.
In addition, we modeled the methyl transfer reactions cata-
lyzed by SET7/9 Y245A (Fig. 5). In the model of the lysine sub-
strate complex, the -amino group is aligned for methyl transfer
by a hydrogen bond to the carbonyl oxygen of Gly-264, result-
ing in a short reaction distance (2.3 Å) and a suboptimal reac-
tion angle (141°) with the methyl group of AdoMet (Fig. 5A).
This misalignment appears to be a direct consequence of the
Y245A mutation that abolishes hydrogen bonding to the -
amino group, illustrating that the suboptimal orientation of the
-amine likely contributes to the diminished activity of this
mutant toward unmodified substrates (11). Conversely, in the
modeled monomethyl-lysine substrate complex for SET7/9
Y245A, Tyr-305 hydrogen bonds to the -amino group, aligning
it for methyl transfer (Fig. 5B). In addition, CH–O hydrogen
bonds to the monomethyl-lysine methyl group and the dissoci-
ation of water 2 from the active site also contribute to reposition-
ing the -amino group for dimethylation. Collectively, these inter-
actions orient the -amine in a reaction angle of 165° that is more
conducive to methyl transfer. However, the reaction distance for
dimethylation is 0.6 Å longer than that in the corresponding
Y305F model because Tyr-305 is positioned further from
AdoMet than Tyr-245 (Figs. 4C and 5B).
In the third methyl transfer reaction catalyzed by SET7/9
Y245A, the lone pair of electrons of the dimethyl-lysine -
amino group acts as the nucleophile and thus cannot engage in
hydrogen bonding. The structure of the trimethyl-lysine prod-
uct complex (Fig. 3D) implies that the dimethyl -amine is
aligned via CH–O hydrogen bonds to its methyl groups, as
shown in the model of the Michaelis complex for this reaction
(Fig. 5C). These CH–O hydrogen bonds restrain the orienta-
tion of the -amino group and position one of the methyl
groups into the solvent pocket, displacing water 1 as discussed
earlier (Fig. 3, D and E). These interactions cumulatively align
the -amino group and AdoMet methyl group with a reaction
distance of 3.0 Å and angle of 162° (Fig. 5C). Taken together, the
models of the substrate complexes for SET7/9 Y245A suggest
that CH–O hydrogen bonds play an increasingly important role
in aligning the methylated -amino group in successive rounds
of methyl transfer.
DISCUSSION
The structural and functional characterization of the
SET7/9 Y245A and Y305F mutants presented here yields
new insights into the mechanism underlying the product
specificity of SET domain KMTs. Importantly, it resolves a
general paradox concerning this specificity. How does the
active site constrain the motion of the lysine -amino group
to align it for methyl transfer with AdoMet, while providing
adequate volume to accommodate the mono-, di-, and tri-
methylated lysine side chain generated during multiple
methyl transfer reactions? The structures of the Y305F and
Y245A mutants resolve this paradox, illustrating that alter-
ations in the positions or occupancies of water molecules
within their active sites generate the space required to
FIGURE 5. Models for the methyl transfer reactions catalyzed by the
SET7/9 Y245A mutant. Models of the Y245A mutant for the first methyl
transfer reaction with TAF10-K189 (A), the second methyl transfer reaction
with TAF10-K189me1 (B), and the third methyl transfer reaction with TAF10-
K189me2 (C). Residues and hydrogen bonds are depicted as in Fig. 3.
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JOURNAL OF BIOLOGICAL CHEMISTRY
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accommodate the multiply methylated -amine produced
during successive catalytic cycles. Minor perturbations in
the side chains of certain active site residues, such as Tyr-
305, are also observed in alignments of the WT enzyme and
the Y245A and Y305F complexes, although these changes are
modest compared with the displacement or dissociation of
the water molecules in the active site. These findings suggest
that the waters function as transient place holders that facil-
itate the SN2 methyl transfer reaction. During monomethy-
lation, they function to constrain the movement of the lysine
-amino group by mediating hydrogen bonds between the
substrate and enzyme, thereby promoting the linear align-
ment with the methyl group and sulfonium cation of
AdoMet (Fig. 4, A and B). During di- and trimethylation, the
water molecules either relocate within the lysine binding
channel or dissociate from the enzyme, yielding the space
required to rotate the methyl group away from the methyl
transfer pore and to realign the -amine in productive geom-
etry for the next methyl transfer reaction (Figs. 4C and 5, B
and C). These findings agree with our prior analysis of the
SET8 Phe/Tyr switch mutant in which we demonstrated that
the Y334F substitution attenuates hydrogen bonding to the
water molecule bound in the solvent pocket, promoting its
dissociation and the conformational changes necessary for
lysine dimethylation (8). Indeed, there is a nearly identical
alignment of the dimethyl-lysine side chains in the structures of
SET7/9Y305FandSET8Y334Fcomplexes,despitethedifferences
in the orientations of the Phe-305 and Phe-334 side chains in each
structure (Fig. 2F). Finally, both the SET8 Y334F (8) and SET7/9
Y305F mutants (Table 2) displayed diminished catalytic efficien-
cies for lysine dimethylation versus monomethylation. These dif-
ferences may reflect the kinetics of the reorganization within the
active site, including the dissociation of the water molecule from
the solvent pocket and the concomitant realignment of the
monomethyl-lysine into a productive geometry for dimethylation.
In addition to their place-holding role, the active site waters
may also facilitate the deprotonation of the lysine -amino
group between methyl transfer reactions. For methylation to
occur, the -amino group must be deprotonated to function as
the nucleophile in the SN2 methyl transfer reaction with
AdoMet (Figs. 4 and 5). Although the pKa value of the lysine
-amine in solution is 10.5, molecular dynamics simulations
by Zhang and Bruice (25, 26) indicate that this value diminishes
to 8.2 upon formation of the SET7/9 Michaelis complex due to
the proximity of the AdoMet sulfonium cation and the low
dielectric constant of the active site. Furthermore, their simu-
lations show that a chain of water molecules facilitates the dep-
rotonation of the -amino group prior to methyl transfer, trans-
ferring the proton to bulk solvent. Although these water
molecule chains are not evident in our crystal structures, the
Y305F and Y245A complexes suggest another potential mech-
anism for deprotonation. In the dimethyl-lysine complexes of
the Y305F and Y245A mutants, the dissociation of water 1 and
2, respectively, from the lysine binding channel requires that
the solvent-mediated hydrogen bond to the -amino group is
broken (Figs. 2D and 3C). It is conceivable that these waters
dissociate from the active site as hydronium ions, promoting
the realignment and deprotonation of the methyl -amino
group for the next methyl transfer reaction.
A comparison of the SET7/9 Y305F and SET8 Y334F com-
plexes yields insights into the mechanism by which the Phe/Tyr
switch influences water binding within the solvent pocket. The
phenylalanine substitution in the Phe/Tyr switch results in the
loss of a single hydrogen bond to the water molecule (water 1) in
the solvent pocket compared with the four hydrogen bonds that
coordinate the solvent molecule in WT SET7/9 (Fig. 2, A and B)
and SET8 (7, 8). Although this attenuation in hydrogen bonding
may appear insignificant, this difference is nonetheless impor-
tant for at least two reasons. First, theoretical calculations indi-
cate that, on average, water molecules form 3.5 hydrogen
bonds in solutions (40, 41). This value is greater than the num-
ber of hydrogen bonds coordinating water 1 in the solvent
pocket in SET7/9 Y305F (Fig. 2, B and C) as well as in SET8
Y334F and other di- and trimethyltransferases that possess a
hydrophobic residue in the Phe/Tyr switch site (8). From the
perspective of the water molecule, the greater hydrogen bond-
ing potential in solution would tend to thermodynamically
favor its dissociation from the solvent pocket in SET domain
KMTs that lack a tyrosine in the Phe/Tyr switch position. Sec-
ond, the ordered binding of water molecules observed in the
active sites of SET domain ternary complexes represents an
unfavorable entropy compared with their diffusion in bulk sol-
vent. In WT SET7/9 (Fig. 2A) and SET8 (7, 8), this entropic
penalty can be partially offset through the favorable enthalpy of
binding associated with the four hydrogen bonds that coordi-
nate the water within the solvent pocket. It is conceivable that
the loss of the tyrosine-mediated hydrogen bond in the Phe/Tyr
switch shifts the equilibrium in favor of dissociation of the
water molecule from the solvent pocket, thereby facilitating
dimethylation in SET7/9 Y305F, SET8 Y334F, and other di- and
trimethyltransferases.
The structures of the SET7/9 Y245A and Y305F complexes
illustrate the interactions that align the lysine -amino group
during the methyl transfer reactions in each enzyme. In the WT
enzyme and the Y305F mutant, hydrogen bonding to the
hydroxyl group of Tyr-245 appears to be critical in properly
aligning the -amine for methyl transfer (Fig. 4). Tyr-245 is
conserved in the sequences of many SET domain KMTs (8, 42),
and substitutions of this residue generally impair or abolish
activity, indicating its importance in catalysis (8, 43). However,
SET7/9 appears to be an exception to this rule, as the Y245A
mutant is not only active but is capable of catalyzing lysine
trimethylation. In this mutant, Tyr-305 appears to assume the
role of Tyr-245 by hydrogen bonding to the monomethylated
-amino group to align it for methyl transfer with AdoMet, as
illustrated in the modeled substrate complex for the dimethy-
lation reaction (Fig. 5B). Conversely, in the model for trimethy-
lation, the Tyr-305 hydroxyl group does not hydrogen bond to
the -amine but instead participates in a CH–O hydrogen bond
with one of the methyl groups to assist in aligning the dimethy-
lated -amine for the methyl transfer reaction (Fig. 5C). Addi-
tional structural and functional studies of the SET domain tri-
methyltransferases will aid in further illuminating the roles
of CH–O hydrogen bonds in facilitating lysine multiple
methylation.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31857
Acknowledgments—We acknowledge S. Schiebold for assistance in
protein expression, purification, and crystallization and S. Anderson
and R. Sanishvili for their assistance with x-ray data collection. We
also thank S. Bulfer and S. Horowitz for reading the manuscript and
providing useful comments. This work utilized the Protein Structure
Facility of the Michigan Diabetes Research and Training Center, Uni-
versity of Michigan, supported by National Institutes of Health Grant
DK020572, NIDDK. Use of the Advanced Photon Source was sup-
ported by the United States Department of Energy, Basic Energy Sci-
ences, Office of Science, under Contract DE-AC02-06CH11357.
GM/CA CAT has been funded in whole or in part by National
Institutes of Health NCI Grant Y1-CO-1020 and NIGMS Grant
Y1-GM-1104. Use of the LS-CAT Sector 21 was supported by Michi-
gan Economic Development Corporation and the Michigan Technol-
ogy Tri-Corridor Grant 085P1000817 for the support of this research
program.
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Lysine Methylation by SET7/9 Mutants
31858
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
|
3M55
|
SET7/9 Y305F in complex with TAF10-K189me1 peptide and AdoHcy
|
SET7/9 Catalytic Mutants Reveal the Role of Active Site Water
Molecules in Lysine Multiple Methylation*□
S
Received for publication,February 15, 2010, and in revised form, July 14, 2010 Published, JBC Papers in Press,August 1, 2010, DOI 10.1074/jbc.M110.114587
Paul A. Del Rizzo‡, Jean-Franc¸ois Couture‡§1, Lynnette M. A. Dirk¶, Bethany S. Strunk‡, Marijo S. Roiko‡,
Joseph S. Brunzelle, Robert L. Houtz¶, and Raymond C. Trievel‡2
From the ‡Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109, the §Ottawa Institute of
Systems Biology, Department of Biochemistry, Microbiology, and Immunology, University of Ottawa, Ottawa, Ontario K1H 8M5,
Canada, the ¶Department of Horticulture, Plant Physiology/Biochemistry/Molecular Biology Program, University of Kentucky,
Lexington, Kentucky 40546, and the Department of Molecular Pharmacology and Biological Chemistry, Northwestern University
Center for Synchrotron Research, Life Sciences Collaborative Access Team, Argonne, Illinois 60439
SET domain lysine methyltransferases (KMTs) methylate
specific lysine residues in histone and non-histone substrates.
These enzymes also display product specificity by catalyzing dis-
tinct degrees of methylation of the lysine -amino group. To
elucidate the molecular mechanism underlying this specificity,
we have characterized the Y245A and Y305F mutants of the
human KMT SET7/9 (also known as KMT7) that alter its prod-
uct specificity from a monomethyltransferase to a di- and a tri-
methyltransferase, respectively. Crystal structures of these
mutants in complex with peptides bearing unmodified, mono-,
di-, and trimethylated lysines illustrate the roles of active site
water molecules in aligning the lysine -amino group for methyl
transfer with S-adenosylmethionine. Displacement or dissocia-
tion of these solvent molecules enlarges the diameter of the
active site, accommodating the increasing size of the methylated
-amino group during successive methyl transfer reactions.
Together, these results furnish new insights into the roles of
active site water molecules in modulating lysine multiple meth-
ylation by SET domain KMTs and provide the first molecular
snapshots of the mono-, di-, and trimethyl transfer reactions
catalyzed by these enzymes.
SET domain enzymes represent a family of S-adenosylmethi-
onine (AdoMet)3-dependent methyltransferases that catalyze
the site-specific methylation of protein lysyl residues in a host
of proteins, including histones, transcription factors, chroma-
tin-modifying enzymes, ribosomal subunits, and other sub-
strates (1–3). In many instances, these modifications serve to
recruit effector proteins that recognize methyl-lysyl residues in
a sequence-dependent fashion (4). In addition, SET domain
KMTs exhibit product specificity, defined as their ability to cat-
alyze mono-, di-, or trimethylation of the lysine -amino group.
This specificity is biologically relevant because many methyl-
lysine-binding proteins can discriminate among different
degrees of lysine methylation (4). Thus, both the site and degree
of lysine methylation are critical to recognition by effector
proteins.
Structural and functional studies have identified a Phe/Tyr
switch in the active site of SET domain KMTs that governs their
respective product specificities (5, 6). According to this model,
KMTs that possess a tyrosine in the Phe/Tyr switch site are
limited to catalyzing lysine monomethylation, whereas en-
zymes that possess a phenylalanine or another hydrophobic
residue in this position display di- or trimethyltransferase activ-
ity. Mutational analysis of various SET domain KMTs, includ-
ing DIM-5 (KMT1) (6), SET7/9 (6), G9A (KMT1C) (5), and
SET8 (also known as PR-SET7 and KMT5A) (7, 8), has demon-
strated that substitutions in the Phe/Tyr switch result in pre-
dictable changes in product specificity. Several models have
been proposed to explain the mechanism by which the Phe/Tyr
switch site governs this specificity, including variations in the
diameter of the active site due to the size of Phe/Tyr switch
residue and steric hindrance by the tyrosine hydroxyl group (6,
9–11). However, our recent studies of the Phe/Tyr switch
mutant Y334F in the human histone H4 Lys-20 (H4K20) meth-
yltransferase SET8 indicate that the Phe/Tyr switch regulates
product specificity via a more subtle mechanism (8). Specifi-
cally, the switch modulates the binding of an active site water
molecule that in turn regulates the transition from mono-
methylation to multiple methylation.
Among the KMTs that have been structurally characterized,
SET7/9 has emerged as an archetypal model for studying the
catalytic mechanism and product specificity of the SET domain
family. Although initially isolated as a histone H3 Lys-4
(H3K4)-specific methyltransferase, this KMT has been shown
to regulate the functions of numerous non-histone substrates
through site-specific methylation (12–21). Early structural and
functional studies of SET7/9 identified two active site mutants,
* Thisworkwassupported,inwholeorinpart,byNationalInstitutesofHealth
Grant R01 GM073839 (to R. C. T.) and National Institutes of Health Admin-
istrative Supplement GM073839-04S1 (to R. C. T.) funded through the
American Recovery and Reinvestment Act. This work was also supported
by Department of Energy Grant DE-FG02-92ER20075 (to R. L. H.).
□
S The on-line version of this article (available at http://www.jbc.org) contains
supplemental Table 1.
The atomic coordinates and structure factors (codes 3M53, 3M54, 3M55, 3M56,
3M57,3M58,3M59,and3M5A)havebeendepositedintheProteinDataBank,
Research Collaboratory for Structural Bioinformatics, Rutgers University, New
Brunswick, NJ (http://www.rcsb.org/).
1 Supported by a Canadian Institutes of Health Research postdoctoral
fellowship.
2 To whom correspondence should be addressed: Dept. of Biological Chem-
istry, University of Michigan Medical School, 1150 West Medical Center Dr.,
5301 MSRB III, Ann Arbor, MI 48109. Tel.: 734-647-0889; Fax: 734-763-4581;
E-mail: rtrievel@umich.edu.
3 The abbreviations used are: AdoMet, S-adenosylmethionine; AdoHcy, S-ad-
enosylhomocysteine; KMT, lysine methyltransferase; ITC, isothermal titra-
tion calorimetry; CH–O, carbon-oxygen hydrogen bond; BisTris, 2-[bis(2-
hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 41, pp. 31849–31858, October 8, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31849
Y245A and Y305F, which change its product specificity. The
Phe/Tyr switch mutant Y305F alters SET7/9 product specificity
from a mono- to dimethyltransferase (6), whereas the Y245A
substitution converts the enzyme into a trimethyltransferase
with weak monomethyltransferase activity (11). These mutants
have been the subjects of numerous molecular modeling simu-
lations that have led to various models to explain their distinct
product specificities (22–26). However, the lack of structural
data for the SET7/9 Y245A and Y305F mutants in complex with
cognate methylated peptides has hindered our understanding
of the mechanisms that define the respective product specific-
ities of these mutants. Moreover, these structures would yield a
framework for visualizing the mono-, di-, and trimethylation
reactions catalyzed by SET domain KMTs.
To gain insight into the molecular basis of their product
specificities, we have determined high resolution crystal
structures of the SET7/9 Y245A and Y305F mutants in com-
plex with peptides of the TATA box-binding protein-associ-
ated factor TAF10 bearing the Lys-189 methylation site in
unmodified (K189), monomethylated (K189me1), dimethyl-
ated (K189me2), and trimethylated (K189me3) states. The
structures and accompanying biochemical data support a
model whereby changes in the occupancy or position of water
molecules in the active site are critical in establishing the prod-
uct specificities of the SET7/9 Y245A and Y305F mutants.
Together, our results provide new insights into the mechanisms
that govern SET domain product specificity and provide step-
wise snapshots of the lysine mono-, di-, and trimethyl transfer
reactions catalyzed by KMTs.
EXPERIMENTAL PROCEDURES
Cloning, Expression, and Purification of the SET7/9 Mutants—
The Y245A and Y305F mutants were introduced into the pHIS2
SET7/9 expression vector encoding residues 110–366 (27) via
QuikChange site-directed mutagenesis (Stratagene) and were
verified by dideoxy DNA sequencing. The plasmids encoding
wild type (WT) SET7/9 and the Y245A and Y305F mutants
were transformed into Rosetta2 DE3 cells (Novagen) and were
expressed as described previously (27, 28). In the course of
characterizing WT SET7/9, we observed that the enzyme co-
purified with AdoMet or another contaminant that resulted in
technical difficulties in the isothermal titration calorimetry
(ITC) experiments and co-crystallization trials with the TAF10
peptides. To overcome this problem, a denaturation and refold-
ing step was inserted in the purification scheme. The denatur-
ation and refolding protocol involved adding 6 M guanidine
HCl, 50 mM sodium phosphate, pH 7.5, and 500 mM NaCl to the
protein while it was immobilized on a nickel-Sepharose column
(GE Healthcare). The column was washed with this buffer, fol-
lowed by a solution of 6 M urea, 50 mM sodium phosphate, pH
7.5, 150 mM NaCl, and 10 mM 2-mercaptoethanol to remove the
cofactor from the denatured enzyme. A reverse gradient from 6
to 0 M urea was then performed in the same buffer to refold the
protein, which was subsequently eluted from the column using
a linear gradient of 0–500 mM imidazole in 50 mM sodium
phosphate, pH 7.5, 500 NaCl, and 10 mM 2-mercaptoethanol.
The refolded protein was digested with tobacco etch virus pro-
tease (29) during dialysis against 20 mM Tris, pH 7.5, 150 mM
NaCl, and 5 mM 2-mercaptoethanol and then purified using a
Superdex 200 gel filtration column (GE Healthcare). Protein
concentration was determined by its absorbance at 280 nm.
Synthetic Peptides—The TAF10 peptides bearing K189,
K189me1,
K189me2,
and
K189me3
(sequence,
acetyl-
SKSK189DRKYTL-amide) and a biotinylated TAF10 peptide
(sequence,
acetyl-SKSK189DRKYTLT(K-EZLink-S-S-biotin)-
amide) were synthesized and purified by New England Peptide,
Inc. Peptide concentrations were measured using the absorb-
ance of their tyrosine residue at 274 nm.
Crystallization and Data Collection—Crystals were pro-
duced by hanging drop vapor diffusion by mixing the crystalli-
zation solution in a 1:1 ratio with 10–20 mg/ml SET7/9, 1 mM
S-adenosylhomocysteine (AdoHcy), and 1.0–3.0 mM unmodi-
fied or methylated TAF10-K189 peptide in 20 mM Tris, pH 8.0,
100 mM NaCl, and 2 mM tris(2-carboxyethyl)phosphine. Crys-
tals were obtained at 20 °C in either 1.8–2.0 M (NH4)2SO4, with
0.1 M BisTris, pH 6.2–6.6, or in 0.95–1.1 M sodium citrate with
100 mM imidazole pH 8.0–8.4. In both crystallization condi-
tions, the final pH values were between pH 8.0 and 9.0. Crystals
in the (NH4)2SO4 condition were typically flash-frozen in the
mother liquor containing 25–30% glycerol, and the crystals in
the citrate condition were frozen in 1.6 M sodium citrate. Data
were collected at the Advanced Photon Source beamlines
21-IDG (LS-CAT) and 23-IDD (GM/CA-CAT). Images were
indexed, integrated, and scaled using HKL2000 (30). Structures
of the mutants were solved by molecular replacement using
MOLREP (31) with the coordinates of a previously reported
SET7/9 ternary complex used as the search model (Protein
Data Bank code 2F69). Successive rounds of model building and
refinement were carried out using Coot (32) and REFMAC (33),
respectively. The geometry of the models were verified by Mol-
Probity (34). Simulated annealing omit maps were calculated
using CNS (35) with the peptide and cofactor removed to elim-
inate model bias in the active site. Structural figures were ren-
dered using PyMOL (Schro¨dinger, LLC.).
Fluorescent Methyltransferase Assay—A coupled fluorescent
methyltransferase assay was used to measure the kinetic
parameters of WT SET7/9 and the Y245A and Y305F mutants
as reported previously, with the exception that 50–150 nM
enzyme, 100 M AdoMet, and varying concentrations of TAF10
peptide substrate were used (27, 36). Assays were performed in
triplicate, and a homocysteine calibration curve was used to
calculate the initial velocities. Kinetic parameters were calcu-
lated by plotting the velocities versus peptide concentration and
by fitting the Michaelis-Menten equation to the data via non-
linear regression using Prism 5.0 (GraphPad). In cases where
the Km value was beyond the measurable range of the assay, the
kcat/Km value was determined as described previously (7).
Calorimetry Experiments—ITC was performed at 20 °C using
a MicroCal VP-ITC calorimeter (GE Healthcare) with 0.12 mM
protein and 4 mM AdoHcy in 20 mM sodium phosphate, pH 7,
and 100 mM NaCl with 1.5 mM peptide as the injectant. Data
were processed, and equilibrium dissociation constants (KD)
and curve fitting errors were calculated from the binding iso-
therms using Origin 7.0 (OriginLab Corp.). WT SET7/9 and the
Y245A and Y305F mutants displayed ligand:protein binding
stoichiometries (N values) between 0.8 and 1.0, demonstrating
Lysine Methylation by SET7/9 Mutants
31850
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
that WT SET7/9 and its mutants were properly refolded due to
their ability to bind peptides in an 1:1 molar ratio.
TLC Product Analysis—Methyltransferase assays were per-
formed in triplicate at 37 °C with the biotinylated TAF10 pep-
tide (0.5 mM) for 30 min in the presence of WT SET7/9 (3
pmol), Y305F mutant (6 pmol), or the Y245A mutant (100
pmol). Assays contained 100 mM HEPES, pH 8.0, 50 mM NaCl,
1.0 mM [methyl-3H]AdoMet (diluted to a specific activity of 0.2
Ci/nmol with purified AdoMet (37)), 15 M Sulfolobus solfa-
taricus AdoHcy hydrolase (36), and 2 units of adenosine deami-
nase (Roche Applied Science) in a final volume of 20 l. The
reactions were terminated by addition of an equal volume of
200 mM MES, pH 5.1, prior to addition of a 2-fold molar excess
of immobilized avidin resin (UltraLink; Pierce). Biotinylated
peptides were allowed to bind at room temperature for 30 min,
and the resin was then collected by centrifugation (9000 g).
The resin was washed three times with 300 mM NaCl, and the
peptide was eluted overnight from the avidin resin by cleavage
of the disulfide bond in the linker of the peptide using 10 mM
tris(2-carboxyethyl)phosphine (Thermo Scientific). The resin
was incubated with additional 10 mM tris(2-carboxyethyl)phos-
phine the following day until the radiolabel was essentially
removed from the resin. The recovered peptides were hydro-
lyzed by incubation with 6 M HCl at 110 °C for 24 h. Subsequent
steps in measuring the radiolabel incorporated into the mono-,
di-, and trimethyl-lysine products were performed as reported
previously (8).
RESULTS
Functional Analysis of the SET7/9
Y305F Mutant—Prior studies of
SET7/9 by Zhang et al. (6) reported
that mutation of the Phe/Tyr switch
residue Tyr-305 to a phenylalanine
alters its product specificity from a
mono- to dimethyltransferase. We
verified these findings by demon-
strating that WT SET7/9 mono-
methylated the TAF10-K189 pep-
tide, whereas the Y305F mutant
mono- and dimethylated this sub-
strate, as demonstrated by mass
spectrometry (data not shown). We
next examined whether the Y305F
substitution altered the affinity of SET7/9 for the TAF10-K189
peptides using ITC (Fig. 1). A comparison of the KD values
revealed that SET7/9 Y305F bound the TAF10-K189 and
TAF10-K189me1 peptides 4- and 6-fold more tightly, respec-
tively, than the WT enzyme, whereas this mutant displayed a
substantially diminished affinity for the TAF10-K189me2 pep-
tide (Table 1). Although the WT enzyme and the Y305F mutant
exhibited discernable differences in their affinities for the
unmodified and monomethylated peptides, these variations are
modest and cannot account for their distinct product specific-
ities, suggesting that a kinetic effect during methylation may be
responsible.
To investigate this possibility, we characterized the kinetic
parameters of WT SET7/9 and the Y305F mutant using the
TAF10 peptides as substrates. Both enzymes methylated the
unmodified peptide with comparable kcat and Km values (Table
2). In analyzing the kinetic parameters for the methylation
of the monomethylated peptide by SET7/9 Y305F, we found
that this substrate displayed an elevated Km value that was
beyond the measurable range of the assay due to its limited
solubility. In this case, we measured the catalytic efficiency
(kcat/Km) for the methylation of this peptide and found that it
was methylated 15-fold less efficiently than the unmodified
peptide by SET7/9 Y305F. Given the fact that the Y305F mutant
exhibited a higher binding affinity for the TAF10-K189me1
peptide than the WT enzyme (Table 1), the kinetic data suggest
that a step in the reaction pathway following substrate binding
limits the catalytic efficiency of this mutant.
We next examined whether the Y305F mutant dimethylated
the TAF10-K189 peptide via a processive or a distributive
mechanism. In a processive mechanism, the methyl-lysine
substrate would remain bound to the enzyme during successive
methyl transfer reactions; thus, the concentration of an inter-
mediate, such as monomethyl-lysine, cannot exceed the en-
zyme concentration during the assay. In a distributive mecha-
nism, the intermediates are released into solution where they
accumulate prior to the next round of methylation, resulting in
an intermediate concentration that is greater than that of the
enzyme. Using a radiometric TLC assay and a biotinylated
TAF10 peptide, we quantified the amounts of monomethylated
products generated by the WT SET7/9 and the Y305F mutant
FIGURE 1. ITC analysis of WT SET7/9 and the Y245A and Y305F mutants. Representative titrations and
binding isotherms for the calorimetry experiments are shown for the unmodified TAF10-K189 peptide titrated
into WT SET7/9 (left plot), SET7/9 Y305F (middle plot), and SET7/9 Y245A (right plot). In each panel, the ITC
titration experiment (upper panel) is shown with the binding isotherms (lower panel) fit to a single binding site
model.
TABLE 1
Analysis of the binding affinity of WT SET7/9 and its catalytic
mutants for unmodified and methylated TAF10 peptides
SET7/9
TAF10 peptide
KD
a
M
WT
K189
4.9 0.20
WT
K189me1
4.0 0.36
Y305F
K189
1.3 0.10
Y305F
K189me1
0.62 0.065
Y305Fb
K189me2
70
Y245A
K189
4.0 0.25
Y245A
K189me1
3.3 0.10
Y245A
K189me2
5.8 0.22
Y245A
K189me3
11 0.28
a Curve fitting errors were calculated from the binding isotherms.
b An estimate of the affinity is reported due to weak peptide binding.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31851
(Table 3). The data demonstrate that comparable amounts of
monomethyl-lysine were generated when the quantity of
enzyme usedistakenintoaccount,inagreementwiththeirsimilar
turnover numbers for the TAF10-K189 peptide (Table 2). The
Y305F mutant also produced small but measurable quantities of
radiolabeled dimethyl-lysine product that were substantially
smaller than the amount of monomethyl-lysine generated.
Therefore, the TLC data are consistent with a distributive
mechanism for dimethylation by the Y305F mutant because the
amount of monomethyl-lysine produced exceeded the quantity
of enzyme used in the assay.
Structures of WT SET7/9 and the Y305F Mutant in Complex
with Unmodified and Methylated TAF10 Peptides—To deter-
mine the mechanism by which the Y305F substitution alters the
product specificity of SET7/9, we determined the crystal struc-
tures of this mutant bound to AdoHcy and TAF10-K189,
TAF10-K189me1, and TAF10-K189me2 peptides and com-
pared these to the structures of the WT SET7/9AdoHcy
TAF10-K189 complex (supplemental Table 1). The structures
of these complexes were determined to 1.85 Å or higher reso-
lution, permitting unambiguous modeling of the K189 side
chains in the active site of the enzyme based on simulated
annealing omit maps (Fig. 2). The ternary complexes of the WT
and the Y305F mutant superimpose with overall root mean
square differences of less than 0.3 Å for all aligned atoms, indi-
cating that neither the Y305F mutation nor the binding of the
various TAF10-K189 peptides results in substantial changes in
its overall structure.
An inspection of the active sites of the SET7/9 WT and
Y305F complexes illustrates the binding modes of the unmod-
ified and methylated forms of K189 in the TAF10 peptides (Fig.
2, A–D). The K189 side chain binds in an extended all trans
conformation in a deep pocket, termed the lysine binding chan-
nel, that is composed of Tyr-245, Thr-266, Leu-267, Ser-268,
Tyr-305, Tyr-335, and Tyr-337 (not shown for clarity) (Fig. 2A).
These residues interact with the aliphatic portion of the K189
side chain primarily through van der Waals contacts. The lysine
binding channel connects to the AdoMet-binding site on the
opposite face of the catalytic domain via an oxygen-lined
methyl transfer pore (38). During catalysis, the methyl group of
the cofactor is positioned within the methyl transfer pore
for the SN2 reaction with the -amino group of the lysine or
methyl-lysine substrate (see below).
To lower the activation barrier for this reaction, the lysine
-amine nucleophile is aligned for methyl transfer through a
hydrogen bond network within the active site. In the WT
enzyme, the K189 -amino group hydrogen bonds to the
hydroxyl group of Tyr-245 as well as to two water molecules
(Fig. 2A). One of the water molecules (termed water 1), is coor-
dinated in a solvent pocket, through hydrogen bonds to the
carbonyl oxygens of Gly-292 and Ala-295 and to the hydroxyl
group of the Phe/Tyr switch residue Tyr-305. This solvent
pocket is structurally conserved in SET domain KMTs and has
an important role in defining product specificity through the
adjacent Phe/Tyr switch residue, as shown in our prior studies
of the human H4K20 methyltransferase SET8 (8). The other
water molecule is bound within the methyl transfer pore
between the lysine substrate and the thioether sulfur atom of
AdoHcy through hydrogen bonds to Tyr-245, Asn-265, and
His-293 in SET7/9 and the TAF10-K189 -amino group. This
water is not observed in other structures of SET7/9 ternary
complexes and may represent the approximate position that
the AdoMet methyl group occupies in the methyl transfer pore
in the Michaelis complex.
In structures of the Y305F ternary complexes, the K189,
K189me1, and K189me2 side chains also adopt extended trans
side chain geometries within the lysine binding channel that are
stabilized via hydrogen bonding to Tyr-245 in the enzyme (Fig.
2, B–D). The orientations of the K189me1 and K189me2 side
chains are further maintained through carbon-oxygen (CH–O)
hydrogen bonding between the methyl groups and oxygen
atoms within the vicinity of the methyl transfer pore, as
reported previously in other SET domain KMT structures (8,
10, 38). A superimposition of the SET7/9 WT and Y305F com-
plexes underscores the similarity of the lysyl binding conforma-
tions (Fig. 2E). However, there are notable differences in the
hydrogen bond patterns and occupancy of water 1 within the
solvent pocket in the Y305F mutant compared with the WT
enzyme. Specifically, the Y305F substitution results in the loss
of one hydrogen bond to water 1 in the structures of the TAF10-
K189 and TAF10-K189me1 complexes (Fig. 2, B and C). In con-
trast, water 1 is absent in TAF10-K189me2 complex, and the
vacated solvent pocket is occupied by one of the methyl groups
TABLE 2
Kinetic parameters of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
TAF10 peptide substrate
Km
a
kcat
a
kcat/Km
a
M
min1
M1 min1 103
WT
K189
160 17
17 0.62
110 17
Y305F
K189
88 5.0
17 0.30
190 11
Y305Fb
K189me1
11 0.50
Y245A
K189
200 35
0.53 0.04
2.6 0.47
Y245A
K189me1
210 23
5.9 0.23
28 3.3
Y245A
K189me2
400 29
6.5 0.16
15 1.2
a Curve fitting errors were calculated from the hyperbolic fits for the Michaelis-Menten equation.
b Km and kcat were not determined because the Km value was beyond the measurable range; therefore, the kcat/Km value is reported.
TABLE 3
Product analysis of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
Quantity
of enzyme
Measured
product
Amount of
product formeda
nmol
nmol
WT
0.003
Kme1
0.65 0.07
Y305F
0.006
Kme1
1.5 0.49
Kme2
0.033 0.009
Y245A
0.100
Kme1
0.80 0.22
Kme2
0.39 0.021
Kme3
0.076 0.019
a Standard deviation was calculated from triplicate measurements.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
of the dimethyl -amine (Fig. 2D). This methyl group forms a
3.5-Å CH–O hydrogen bond to the carbonyl oxygen of Ala-295,
further stabilizing the binding of the dimethyl-lysine side chain.
A homologous dimethyl-lysine-binding mode and CH–O
hydrogen bond was observed in our prior structural studies of
the SET8 Y334F Phe/Tyr switch mutant that confers an analo-
gous change in product specificity from a mono- to a dimeth-
yltransferase (8). A structural alignment of the active sites of the
SET7/9 Y305F and SET8 Y334F mutants bound to cognate
dimethylated peptides illustrates that the coordinates of the
dimethyl-lysyl side chains are virtually superimposable, with
one methyl group oriented toward the methyltransfer pore and
the second positioned within the vacant solvent pocket (Fig.
2F). Taken together, the structures of the SET7/9 Y305F com-
plexes and the similarities in the dimethyl-lysine conforma-
tions in the SET7/9 Y305F and SET8 Y334F mutants imply that
the Phe/Tyr switch governs product specificity through a con-
served mechanism whereby it indirectly influences the binding
modes of the methyl-lysine side chain by modulating the affin-
ity of the water molecule (water 1) bound in the solvent pocket.
Biochemical Characterization of
the SET7/9 Y245A Mutant—Previ-
ous studies by Xiao et al. (11)
reported that the Y245A mutation
yields an unusual change in the
product
specificity
of
SET7/9,
converting the enzyme to a trimeth-
yltransferase with weak monometh-
yltransferase activity. We deter-
mined that the SET7/9 Y245A could
mono-, di-, and trimethylate the
TAF10-K189 peptide by mass spec-
trometry (data not shown) and TLC
(Table 3), confirming the earlier
studies of Xiao et al. (11). ITC
analysis revealed that the Y245A
mutant displayed comparable KD
values
for
the
unmodified
and
methylated TAF10-K189 peptides
(Fig. 1), although its affinity for the
trimethylated peptide was modestly
diminished in comparison with the
other peptides (Table 1). The ITC
data demonstrate that the Y245A
mutant
bound
the
unmodified,
mono-,
and
dimethylated
sub-
strates with equivalent affinities,
suggesting that a kinetic effect or
a structural alteration in the active
site may be responsible for its
diminished
activity
toward
un-
modified substrates.
To gain further insight into its
peculiar
product
specificity,
we
characterized the kinetic properties
of
the
SET7/9
Y245A
mutant.
Steady state analysis demonstrated
that this mutant displayed similar
Km values for the unmodified, mono- and dimethylated TAF10
peptides (Table 2). However, the turnover number for the
TAF10-K189 peptide was diminished over 10-fold versus the
methylated peptides and was reduced 30-fold versus the WT
enzyme, in agreement with the weak monomethyltransferase
activity reported by Xiao et al. (11). In addition, we investigated
whether this mutant catalyzes lysine trimethylation via a pro-
cessive or distributive mechanism as described for SET7/9
Y305F. The TLC data illustrate that the mono- and dimethyl-
lysine intermediates accumulated at quantities greater than
that of the enzyme used in the assay, indicating that SET7/9
Y245A obeys a distributive mechanism, analogous to the Y305F
mutant (Table 3).
Structures of SET7/9 Y245A Bound to Unmodified and Meth-
ylated TAF10 Peptides—To elucidate the mechanism underly-
ing its unusual product specificity, we determined the crystal
structures of SET7/9 Y245A in complex with AdoHcy and
unmodified, mono-, di-, and trimethylated TAF10 peptides
(supplemental Table 1). These complexes superimpose with
the structure of the WT SET7/9AdoHcyTAF10-K189 com-
FIGURE 2. Crystal structures of WT SET7/9 and the Y305F mutant in complex with AdoHcy and unmodi-
fied and methylated TAF10 peptides. A, active site of WT SET7/9 bound to AdoHcy and TAF10-K189 and the
active site of SET7/9 Y305F bound to AdoHcy and TAF10-K189 (B), TAF10-K189me1 (C), and TAF10-K189me2
peptides (D). A–D, the K189 residue in the TAF10 peptide is depicted with yellow carbon atoms with the
corresponding Fo – Fc simulated-annealing omit maps (contoured at 2.5 ) shown. AdoHcy is colored with
green carbon atoms, and SET7/9 carbon atoms are colored gray, with the exception of Tyr-305/Y305F
(magenta) and Tyr-245 (cyan). Gly-264 to Ser-268 are shown, and the main chain atoms of Gly-292 to Ala-295
are depicted for clarity. Red dashed lines indicate conventional hydrogen bonds, and blue dashed lines indicate
CH–O hydrogen bonds. Cyan dashed lines indicate weak CH–O hydrogen bonds between 3.5 and 3.7 Å in
length. The water molecule in the solvent binding pocket is numbered 1. E, overlay of the active sites of the WT
enzyme (light blue carbon atoms) and the Y305F mutant (green, yellow, and pink carbon atoms correspond to
K189, K189me1, and K189me2, respectively). Red dashed lines indicate coordination of the waters in the struc-
turesoftheY305Fmutant,andthelightbluedashedlineindicateshydrogenbondingthatonlyoccursintheWT
enzyme. F, superimposition of the active sites of the SET7/9 Y305FAdoHcyTAF10-K189me2 and SET8
Y334FAdoHcyH4K20me2 (Protein Data Bank code 3F9X) complexes shown with green and yellow carbon
atoms, respectively.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31853
plex with root mean squared differences of less than 0.4 Å for all
aligned atoms, indicating that the Y245A mutant does not per-
turb the overall structure of the enzyme. Simulated annealing
omit maps illustrate that K189 side chains are bound within the
lysine binding channel through hydrogen bonds and van der
Waals contacts (Fig. 3, A–D), although the interactions and
binding modes are distinct from those in the complexes of WT
SET7/9 and the Y305F mutant (Fig. 2, A–D). In the unmodified
TAF10 peptide complex, the K189 -amino group forms a weak
hydrogen bond with the carbonyl oxygen of Gly-264 (Fig. 3A),
whereas the -amino groups of K189me1 and K189me2 hydro-
gen bond to the hydroxyl group of Tyr-305 in the mono- and
dimethylated peptide complexes (Fig. 3, B and C). The confor-
mations of the K189me1 and K189me2 side chains are further
stabilized by water-mediated hydrogen bonding and through
CH–O hydrogen bonding to their methyl groups. In the
TAF10-K189me3 peptide complex, the trimethyl-lysine side
chain is coordinated exclusively through direct and water-me-
diated CH–O hydrogen bonds to its methyl groups because the
quaternary -ammonium cation cannot engage in hydrogen
bonding (Fig. 3D).
A structural alignment of the four SET7/9 Y245A complexes
illustrates distinct binding modes for the unmodified versus the
methylated K189 side chains, highlighting the selectivity of this
mutant for methylated substrates. The side chains of K189me1,
K189me2, and K189me3 roughly overlay with their respective
-amino groups superimposed and adopt slightly kinked con-
formations (Fig. 3E), as opposed to the extended trans geome-
try of the unmodified and methylated lysines in the complexes
of the WT enzyme and the Y305F mutant (Fig. 2, A–D). Con-
versely, the unmodified K189 side chain does not superimpose
with its methylated counterparts and is oriented in an alterna-
tive configuration due to its hydrogen bonding to Gly-264 (Fig.
3, A and E). An overlay of the structures of the WT enzyme and
Y245A mutant bound to the unmodified TAF10 peptide illus-
trates that the side chains of K189 do not superimpose and that
the K189 -amino group appears to be misaligned with AdoHcy
in the Y245A complex (Fig. 3F). This suboptimal alignment
may explain the diminished kcat value of SET7/9 Y245A mutant
toward substrates with unmodified lysines (Table 2).
A comparison of the structures of the SET7/9 Y245A and
Y305F complexes yields a molecular explanation for the differ-
ent product specificities of these two mutants. In the SET7/9
Y305F complexes, Tyr-245 aligns the K189 -amino group for
methyl transfer through hydrogen bonding to its hydroxyl
group (Fig. 2, B–D). Conversely, in the Y245A mutant, the
K189me1 and K189me2 -amino groups are oriented through
hydrogen bonding to Tyr-305 (Fig. 3, B and C). These distinct
hydrogen bond patterns impart differences in the conforma-
tions of the lysyl side chains due to the relative orientations of
Tyr-245 and Tyr-305 in the lysine binding channel. Specifically,
the kinked conformation adopted by the K189me1 and
K189me2 side chains in the Y245A complexes (Fig. 3, B and C)
may contribute to the differences in the turnover numbers of
this mutant versus those of the WT enzyme and the Y305F
mutant (Table 2). In addition, the dimethyl -amino group of
the K189me2 side chain binds in distinct orientations in the
Y245A and Y305F mutants due to their hydrogen bonding to
FIGURE 3. Crystal structures of the SET7/9 Y245A mutant in complex with
AdoHcy and unmodified and methylated TAF10 peptides. Active site of
SET7/9 Y245A bound to AdoHcy and TAF10-K189 (A), TAF10-K189me1 (B),
TAF10-K189me2 (C), and TAF10-K189me3 peptides (D). A–D, Fo – Fc simulated-
annealing omit maps (contoured at 2.5 ) for the unmodified and methylated
K189 side chains are illustrated. The residues and hydrogen bonds in each com-
plexarecoloredasinFig.2.Thewatermoleculesinthelysinebindingchannelsof
the Y245A complexes are numbered 1–4, as described in the text. E, superimpo-
sition of the active sites of the Y245A complexes bound to the four methylated
statesofTAF10-K189(unmodified,mono-,di-,andtrimethylatedshowningreen,
yellow, pink, and blue, respectively). F, overlay of WT SET7/9 (blue carbons) and
SET7/9 Y245A (green carbons) in complex with TAF10-K189. Waters correspond-
ing to the WT and Y245A structures are colored cyan and green, respectively.
G, alignment of the active sites of the Y245A (yellow carbons) and Y305F (green
carbons) mutants in complex with the TAF10-K189me2 peptide. Hydrogen
bonds from the Y305F structure are shown as green dashed lines, and waters and
hydrogen bonds in the Y245A structure are shown in yellow and orange,
respectively.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
Tyr-305 and Tyr-245, respectively (Fig. 3G). In the Y305F
mutant, hydrogen bonds to the dimethyl -amino group cou-
pled with steric constraints in the lysine binding channel pre-
vent the K189me2 side chain from undergoing a conforma-
tional change that is conducive to trimethylation (Fig. 2D),
consistent with its dimethyltransferase activity. However, in the
Y245A mutant, the alanine substitution enlarges the diameter
of the lysine binding channel, accommodating trimethyl-lysine
(Fig. 3D). In addition, the larger diameter would permit the
dimethyl-lysine substrate to undergo the conformational reor-
ganization necessary to align the -amino group in a productive
geometry for trimethylation.
A major difference in the active site of the Y245A mutant
versus the other SET7/9 structures is the presence of several
water molecules bound in the cavity generated by the Y245A
mutation. In the structure of the Y245A mutant bound to
TAF10-K189, three water molecules (waters 2–4) occupy this
cavity and are arranged in a triangular geometry (Fig. 3A). In
addition, water 1 shifts 1.6 Å from its position in the solvent
pocket toward water 2 to which it forms a hydrogen bond (Fig.
3, A and E). The shift in water 1 was unexpected given its con-
served orientation in the solvent pocket of the SET7/9 WT and
Y305F complexes (Fig. 2, A–C) as well as in the structures of
other SET domain KMTs (8). This displacement is presumably
related to the alternative conformation of the K189 side chain
whose -amino group is too distant (4.3 Å) to form a productive
hydrogen bond to water 1. Conversely, in the Y245A complexes
bound to TAF10-K189me1 and TAF10-K189me2, water 1
remains tightly bound in the solvent pocket through hydrogen
bonds to Tyr-305 hydroxyl group, the carbonyl oxygens of Gly-
292 and Ala-295, and the K189 -amino group (Fig. 3, B, C, and
E), analogous to its binding in the WT enzyme (Fig. 2A). How-
ever, in the TAF10-K189me3 complex, one of the methyl
groups of the trimethyl -ammonium cation is oriented into the
solvent pocket (Fig. 3D), similar to the dimethyl-lysine binding
mode observed in the Y305F mutant (Fig. 2D). The binding of
the methyl group in the solvent pocket displaces water 1 by 3.2
Å relative to its position in the TAF10-K189me1 complex (Fig.
3E), thereby avoiding a steric clash with the trimethylated
-ammonium group. Variations in the occupancy of water 2 are
also seen in the different Y245A structures. Water 2 is bound in
similar orientations in the active site of the unmodified and
monomethylated peptide complexes but is absent in the di- and
trimethylated peptide complexes due to the binding of a methyl
group in this position (Fig. 3, A–E). In summary, the changes in
the positions or occupancies of waters 1 and 2 correlate with the
binding modes of the unmodified and methylated K189 within
the active site of the Y245A mutant.
Catalytic Models of Lysine Multiple Methylation by SET7/9
Y245A, and Y305F—The structures of the SET7/9 complexes
reported here offer a prime opportunity to generate stepwise
models for lysine mono-, di-, and trimethylation by a SET
domain KMT. We modeled the AdoMet-bound Michaelis
complexes by superimposing the SET7/9 product complexes
with the previously reported structure of the SET7/9-AdoMet
binary complex (Fig. 4) (39). The conformations of the mono-
and dimethyl -amino groups in the Michaelis complexes were
inferred from the coordinates of the corresponding dimethyl-
and trimethyl-lysine products, respectively. In addition, we
modeled the -amino group in a deprotonated state with its
hydrogen atoms oriented toward the hydrogen bond acceptors
that align the lysyl side chain for methylation. As a basis for this
comparison, we first modeled the monomethylation reaction
catalyzed by WT SET7/9 (Fig. 4A). In the substrate ternary
complex, the lysine -amine is aligned with the methyl group
and sulfonium cation of AdoMet through a hydrogen bond to
the Tyr-245 hydroxyl group and water 1 in the solvent pocket.
The values of the reaction distance and angle are 2.8 Å and 153°,
respectively, in approximate agreement with the linear geome-
try of a SN2 methyl transfer reaction calculated in other mod-
eled substrate complexes (8, 10). In the product complex, the
monomethyl-lysine side chain is bound in an extended confor-
mation with its methyl group oriented within the methyl trans-
fer pore, thereby obstructing AdoMet binding. Furthermore,
water 1 remains tightly coordinated in the solvent pocket
through four hydrogen bonds to Gly-292, Ala-295, Tyr-305,
and the monomethyl -amino group. These interactions hinder
the dissociation of water 1 and the related rearrangement of the
monomethyl-lysine side chain required for a second methyl
transfer reaction, explaining why the WT enzyme cannot cata-
lyze di- and trimethylation. These findings concur with the
FIGURE 4. Catalytic models of the methyl transfer reactions catalyzed by
WT SET7/9 and the Y305F mutant. A, monomethylation of TAF10-K189 by
the WT enzyme. The reaction scheme depicts the modeled substrate ternary
complex (left) and the product complex (right) for the transfer of the methyl
group from AdoMet (green carbon atoms) to K189 in TAF10 (yellow carbons),
yielding AdoHcy and K189me1. The red arrow indicates the direction of the
nucleophilic attack of the deprotonated -amino group on the AdoMet
methyl group. The transferred methyl group is colored green, and the white
atoms represent the hydrogens of the -amino group. Hydrogen bonds
and residues in the enzyme active site are illustrated as in Fig. 2. The
reaction distance and angle are labeled in red. B and C, models of the
Y305F mutant for the first methyl transfer reaction with TAF10-K189 (B)
and second methyl transfer reaction with TAF10-K189me1 (C). Color
schemes are the same as in A.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31855
model for SET7/9 product specificity reported in previous
structural and functional studies (6, 11).
Similar reaction geometry is observed in the model for the
monomethyl transfer reaction catalyzed by SET7/9 Y305F.
Hydrogen bonds from the Tyr-245 hydroxyl group and water 1
align the lysine -amino group with the AdoMet methyl group
at a distance of 2.1 Å and an angle of 160°, equivalent to those
measured in the Michaelis complex of the WT enzyme (Fig. 4, A
and B). In the product complex, monomethyl-lysine adopts an
extended trans configuration analogous to that in the WT
enzyme. For dimethylation to occur, the monomethyl-lysine
must undergo a conformational change in which its methyl
group is rotated out of the methyl transfer path with AdoMet.
The structure of the Y305F mutant bound to the dimethylated
TAF10 peptide (Fig. 2D) implies that this rearrangement occurs
through the dissociation of water 1 due to the loss of the Tyr-
305 hydrogen bond in the solvent pocket. The dissociation of
water 1 would enable the monomethyl-lysine side chain to
adopt an alternative conformation through a rotation about its
C–N bond, projecting the methyl group into the solvent
pocket (Fig. 4C). This rotation reorients the methyl group out of
the methyl transfer path while realigning the monomethyl -
amino group for a second methylation reaction through a direct
hydrogen bond to the Tyr-245 hydroxyl group and a CH–O
hydrogen bond between its methyl group and Ala-295. The
modeled reaction geometry for monomethyl-lysine substrate
complex of the Y305F mutant (2.7 Å, 160°) is equivalent to that
of the first methyl transfer reaction in SET7/9 Y305F. These
geometries concur with our previous models for mono- and
dimethylation catalyzed by SET8 Y334F (8), illustrating that the
orientation of a methyl group into the solvent pocket is a con-
served feature of SET domain KMTs that catalyze multiple
methylation.
In addition, we modeled the methyl transfer reactions cata-
lyzed by SET7/9 Y245A (Fig. 5). In the model of the lysine sub-
strate complex, the -amino group is aligned for methyl transfer
by a hydrogen bond to the carbonyl oxygen of Gly-264, result-
ing in a short reaction distance (2.3 Å) and a suboptimal reac-
tion angle (141°) with the methyl group of AdoMet (Fig. 5A).
This misalignment appears to be a direct consequence of the
Y245A mutation that abolishes hydrogen bonding to the -
amino group, illustrating that the suboptimal orientation of the
-amine likely contributes to the diminished activity of this
mutant toward unmodified substrates (11). Conversely, in the
modeled monomethyl-lysine substrate complex for SET7/9
Y245A, Tyr-305 hydrogen bonds to the -amino group, aligning
it for methyl transfer (Fig. 5B). In addition, CH–O hydrogen
bonds to the monomethyl-lysine methyl group and the dissoci-
ation of water 2 from the active site also contribute to reposition-
ing the -amino group for dimethylation. Collectively, these inter-
actions orient the -amine in a reaction angle of 165° that is more
conducive to methyl transfer. However, the reaction distance for
dimethylation is 0.6 Å longer than that in the corresponding
Y305F model because Tyr-305 is positioned further from
AdoMet than Tyr-245 (Figs. 4C and 5B).
In the third methyl transfer reaction catalyzed by SET7/9
Y245A, the lone pair of electrons of the dimethyl-lysine -
amino group acts as the nucleophile and thus cannot engage in
hydrogen bonding. The structure of the trimethyl-lysine prod-
uct complex (Fig. 3D) implies that the dimethyl -amine is
aligned via CH–O hydrogen bonds to its methyl groups, as
shown in the model of the Michaelis complex for this reaction
(Fig. 5C). These CH–O hydrogen bonds restrain the orienta-
tion of the -amino group and position one of the methyl
groups into the solvent pocket, displacing water 1 as discussed
earlier (Fig. 3, D and E). These interactions cumulatively align
the -amino group and AdoMet methyl group with a reaction
distance of 3.0 Å and angle of 162° (Fig. 5C). Taken together, the
models of the substrate complexes for SET7/9 Y245A suggest
that CH–O hydrogen bonds play an increasingly important role
in aligning the methylated -amino group in successive rounds
of methyl transfer.
DISCUSSION
The structural and functional characterization of the
SET7/9 Y245A and Y305F mutants presented here yields
new insights into the mechanism underlying the product
specificity of SET domain KMTs. Importantly, it resolves a
general paradox concerning this specificity. How does the
active site constrain the motion of the lysine -amino group
to align it for methyl transfer with AdoMet, while providing
adequate volume to accommodate the mono-, di-, and tri-
methylated lysine side chain generated during multiple
methyl transfer reactions? The structures of the Y305F and
Y245A mutants resolve this paradox, illustrating that alter-
ations in the positions or occupancies of water molecules
within their active sites generate the space required to
FIGURE 5. Models for the methyl transfer reactions catalyzed by the
SET7/9 Y245A mutant. Models of the Y245A mutant for the first methyl
transfer reaction with TAF10-K189 (A), the second methyl transfer reaction
with TAF10-K189me1 (B), and the third methyl transfer reaction with TAF10-
K189me2 (C). Residues and hydrogen bonds are depicted as in Fig. 3.
Lysine Methylation by SET7/9 Mutants
31856
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
accommodate the multiply methylated -amine produced
during successive catalytic cycles. Minor perturbations in
the side chains of certain active site residues, such as Tyr-
305, are also observed in alignments of the WT enzyme and
the Y245A and Y305F complexes, although these changes are
modest compared with the displacement or dissociation of
the water molecules in the active site. These findings suggest
that the waters function as transient place holders that facil-
itate the SN2 methyl transfer reaction. During monomethy-
lation, they function to constrain the movement of the lysine
-amino group by mediating hydrogen bonds between the
substrate and enzyme, thereby promoting the linear align-
ment with the methyl group and sulfonium cation of
AdoMet (Fig. 4, A and B). During di- and trimethylation, the
water molecules either relocate within the lysine binding
channel or dissociate from the enzyme, yielding the space
required to rotate the methyl group away from the methyl
transfer pore and to realign the -amine in productive geom-
etry for the next methyl transfer reaction (Figs. 4C and 5, B
and C). These findings agree with our prior analysis of the
SET8 Phe/Tyr switch mutant in which we demonstrated that
the Y334F substitution attenuates hydrogen bonding to the
water molecule bound in the solvent pocket, promoting its
dissociation and the conformational changes necessary for
lysine dimethylation (8). Indeed, there is a nearly identical
alignment of the dimethyl-lysine side chains in the structures of
SET7/9Y305FandSET8Y334Fcomplexes,despitethedifferences
in the orientations of the Phe-305 and Phe-334 side chains in each
structure (Fig. 2F). Finally, both the SET8 Y334F (8) and SET7/9
Y305F mutants (Table 2) displayed diminished catalytic efficien-
cies for lysine dimethylation versus monomethylation. These dif-
ferences may reflect the kinetics of the reorganization within the
active site, including the dissociation of the water molecule from
the solvent pocket and the concomitant realignment of the
monomethyl-lysine into a productive geometry for dimethylation.
In addition to their place-holding role, the active site waters
may also facilitate the deprotonation of the lysine -amino
group between methyl transfer reactions. For methylation to
occur, the -amino group must be deprotonated to function as
the nucleophile in the SN2 methyl transfer reaction with
AdoMet (Figs. 4 and 5). Although the pKa value of the lysine
-amine in solution is 10.5, molecular dynamics simulations
by Zhang and Bruice (25, 26) indicate that this value diminishes
to 8.2 upon formation of the SET7/9 Michaelis complex due to
the proximity of the AdoMet sulfonium cation and the low
dielectric constant of the active site. Furthermore, their simu-
lations show that a chain of water molecules facilitates the dep-
rotonation of the -amino group prior to methyl transfer, trans-
ferring the proton to bulk solvent. Although these water
molecule chains are not evident in our crystal structures, the
Y305F and Y245A complexes suggest another potential mech-
anism for deprotonation. In the dimethyl-lysine complexes of
the Y305F and Y245A mutants, the dissociation of water 1 and
2, respectively, from the lysine binding channel requires that
the solvent-mediated hydrogen bond to the -amino group is
broken (Figs. 2D and 3C). It is conceivable that these waters
dissociate from the active site as hydronium ions, promoting
the realignment and deprotonation of the methyl -amino
group for the next methyl transfer reaction.
A comparison of the SET7/9 Y305F and SET8 Y334F com-
plexes yields insights into the mechanism by which the Phe/Tyr
switch influences water binding within the solvent pocket. The
phenylalanine substitution in the Phe/Tyr switch results in the
loss of a single hydrogen bond to the water molecule (water 1) in
the solvent pocket compared with the four hydrogen bonds that
coordinate the solvent molecule in WT SET7/9 (Fig. 2, A and B)
and SET8 (7, 8). Although this attenuation in hydrogen bonding
may appear insignificant, this difference is nonetheless impor-
tant for at least two reasons. First, theoretical calculations indi-
cate that, on average, water molecules form 3.5 hydrogen
bonds in solutions (40, 41). This value is greater than the num-
ber of hydrogen bonds coordinating water 1 in the solvent
pocket in SET7/9 Y305F (Fig. 2, B and C) as well as in SET8
Y334F and other di- and trimethyltransferases that possess a
hydrophobic residue in the Phe/Tyr switch site (8). From the
perspective of the water molecule, the greater hydrogen bond-
ing potential in solution would tend to thermodynamically
favor its dissociation from the solvent pocket in SET domain
KMTs that lack a tyrosine in the Phe/Tyr switch position. Sec-
ond, the ordered binding of water molecules observed in the
active sites of SET domain ternary complexes represents an
unfavorable entropy compared with their diffusion in bulk sol-
vent. In WT SET7/9 (Fig. 2A) and SET8 (7, 8), this entropic
penalty can be partially offset through the favorable enthalpy of
binding associated with the four hydrogen bonds that coordi-
nate the water within the solvent pocket. It is conceivable that
the loss of the tyrosine-mediated hydrogen bond in the Phe/Tyr
switch shifts the equilibrium in favor of dissociation of the
water molecule from the solvent pocket, thereby facilitating
dimethylation in SET7/9 Y305F, SET8 Y334F, and other di- and
trimethyltransferases.
The structures of the SET7/9 Y245A and Y305F complexes
illustrate the interactions that align the lysine -amino group
during the methyl transfer reactions in each enzyme. In the WT
enzyme and the Y305F mutant, hydrogen bonding to the
hydroxyl group of Tyr-245 appears to be critical in properly
aligning the -amine for methyl transfer (Fig. 4). Tyr-245 is
conserved in the sequences of many SET domain KMTs (8, 42),
and substitutions of this residue generally impair or abolish
activity, indicating its importance in catalysis (8, 43). However,
SET7/9 appears to be an exception to this rule, as the Y245A
mutant is not only active but is capable of catalyzing lysine
trimethylation. In this mutant, Tyr-305 appears to assume the
role of Tyr-245 by hydrogen bonding to the monomethylated
-amino group to align it for methyl transfer with AdoMet, as
illustrated in the modeled substrate complex for the dimethy-
lation reaction (Fig. 5B). Conversely, in the model for trimethy-
lation, the Tyr-305 hydroxyl group does not hydrogen bond to
the -amine but instead participates in a CH–O hydrogen bond
with one of the methyl groups to assist in aligning the dimethy-
lated -amine for the methyl transfer reaction (Fig. 5C). Addi-
tional structural and functional studies of the SET domain tri-
methyltransferases will aid in further illuminating the roles
of CH–O hydrogen bonds in facilitating lysine multiple
methylation.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31857
Acknowledgments—We acknowledge S. Schiebold for assistance in
protein expression, purification, and crystallization and S. Anderson
and R. Sanishvili for their assistance with x-ray data collection. We
also thank S. Bulfer and S. Horowitz for reading the manuscript and
providing useful comments. This work utilized the Protein Structure
Facility of the Michigan Diabetes Research and Training Center, Uni-
versity of Michigan, supported by National Institutes of Health Grant
DK020572, NIDDK. Use of the Advanced Photon Source was sup-
ported by the United States Department of Energy, Basic Energy Sci-
ences, Office of Science, under Contract DE-AC02-06CH11357.
GM/CA CAT has been funded in whole or in part by National
Institutes of Health NCI Grant Y1-CO-1020 and NIGMS Grant
Y1-GM-1104. Use of the LS-CAT Sector 21 was supported by Michi-
gan Economic Development Corporation and the Michigan Technol-
ogy Tri-Corridor Grant 085P1000817 for the support of this research
program.
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Lysine Methylation by SET7/9 Mutants
31858
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
|
3M56
|
SET7/9 Y305F in complex with TAF10-K189me2 peptide and AdoHcy
|
SET7/9 Catalytic Mutants Reveal the Role of Active Site Water
Molecules in Lysine Multiple Methylation*□
S
Received for publication,February 15, 2010, and in revised form, July 14, 2010 Published, JBC Papers in Press,August 1, 2010, DOI 10.1074/jbc.M110.114587
Paul A. Del Rizzo‡, Jean-Franc¸ois Couture‡§1, Lynnette M. A. Dirk¶, Bethany S. Strunk‡, Marijo S. Roiko‡,
Joseph S. Brunzelle, Robert L. Houtz¶, and Raymond C. Trievel‡2
From the ‡Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109, the §Ottawa Institute of
Systems Biology, Department of Biochemistry, Microbiology, and Immunology, University of Ottawa, Ottawa, Ontario K1H 8M5,
Canada, the ¶Department of Horticulture, Plant Physiology/Biochemistry/Molecular Biology Program, University of Kentucky,
Lexington, Kentucky 40546, and the Department of Molecular Pharmacology and Biological Chemistry, Northwestern University
Center for Synchrotron Research, Life Sciences Collaborative Access Team, Argonne, Illinois 60439
SET domain lysine methyltransferases (KMTs) methylate
specific lysine residues in histone and non-histone substrates.
These enzymes also display product specificity by catalyzing dis-
tinct degrees of methylation of the lysine -amino group. To
elucidate the molecular mechanism underlying this specificity,
we have characterized the Y245A and Y305F mutants of the
human KMT SET7/9 (also known as KMT7) that alter its prod-
uct specificity from a monomethyltransferase to a di- and a tri-
methyltransferase, respectively. Crystal structures of these
mutants in complex with peptides bearing unmodified, mono-,
di-, and trimethylated lysines illustrate the roles of active site
water molecules in aligning the lysine -amino group for methyl
transfer with S-adenosylmethionine. Displacement or dissocia-
tion of these solvent molecules enlarges the diameter of the
active site, accommodating the increasing size of the methylated
-amino group during successive methyl transfer reactions.
Together, these results furnish new insights into the roles of
active site water molecules in modulating lysine multiple meth-
ylation by SET domain KMTs and provide the first molecular
snapshots of the mono-, di-, and trimethyl transfer reactions
catalyzed by these enzymes.
SET domain enzymes represent a family of S-adenosylmethi-
onine (AdoMet)3-dependent methyltransferases that catalyze
the site-specific methylation of protein lysyl residues in a host
of proteins, including histones, transcription factors, chroma-
tin-modifying enzymes, ribosomal subunits, and other sub-
strates (1–3). In many instances, these modifications serve to
recruit effector proteins that recognize methyl-lysyl residues in
a sequence-dependent fashion (4). In addition, SET domain
KMTs exhibit product specificity, defined as their ability to cat-
alyze mono-, di-, or trimethylation of the lysine -amino group.
This specificity is biologically relevant because many methyl-
lysine-binding proteins can discriminate among different
degrees of lysine methylation (4). Thus, both the site and degree
of lysine methylation are critical to recognition by effector
proteins.
Structural and functional studies have identified a Phe/Tyr
switch in the active site of SET domain KMTs that governs their
respective product specificities (5, 6). According to this model,
KMTs that possess a tyrosine in the Phe/Tyr switch site are
limited to catalyzing lysine monomethylation, whereas en-
zymes that possess a phenylalanine or another hydrophobic
residue in this position display di- or trimethyltransferase activ-
ity. Mutational analysis of various SET domain KMTs, includ-
ing DIM-5 (KMT1) (6), SET7/9 (6), G9A (KMT1C) (5), and
SET8 (also known as PR-SET7 and KMT5A) (7, 8), has demon-
strated that substitutions in the Phe/Tyr switch result in pre-
dictable changes in product specificity. Several models have
been proposed to explain the mechanism by which the Phe/Tyr
switch site governs this specificity, including variations in the
diameter of the active site due to the size of Phe/Tyr switch
residue and steric hindrance by the tyrosine hydroxyl group (6,
9–11). However, our recent studies of the Phe/Tyr switch
mutant Y334F in the human histone H4 Lys-20 (H4K20) meth-
yltransferase SET8 indicate that the Phe/Tyr switch regulates
product specificity via a more subtle mechanism (8). Specifi-
cally, the switch modulates the binding of an active site water
molecule that in turn regulates the transition from mono-
methylation to multiple methylation.
Among the KMTs that have been structurally characterized,
SET7/9 has emerged as an archetypal model for studying the
catalytic mechanism and product specificity of the SET domain
family. Although initially isolated as a histone H3 Lys-4
(H3K4)-specific methyltransferase, this KMT has been shown
to regulate the functions of numerous non-histone substrates
through site-specific methylation (12–21). Early structural and
functional studies of SET7/9 identified two active site mutants,
* Thisworkwassupported,inwholeorinpart,byNationalInstitutesofHealth
Grant R01 GM073839 (to R. C. T.) and National Institutes of Health Admin-
istrative Supplement GM073839-04S1 (to R. C. T.) funded through the
American Recovery and Reinvestment Act. This work was also supported
by Department of Energy Grant DE-FG02-92ER20075 (to R. L. H.).
□
S The on-line version of this article (available at http://www.jbc.org) contains
supplemental Table 1.
The atomic coordinates and structure factors (codes 3M53, 3M54, 3M55, 3M56,
3M57,3M58,3M59,and3M5A)havebeendepositedintheProteinDataBank,
Research Collaboratory for Structural Bioinformatics, Rutgers University, New
Brunswick, NJ (http://www.rcsb.org/).
1 Supported by a Canadian Institutes of Health Research postdoctoral
fellowship.
2 To whom correspondence should be addressed: Dept. of Biological Chem-
istry, University of Michigan Medical School, 1150 West Medical Center Dr.,
5301 MSRB III, Ann Arbor, MI 48109. Tel.: 734-647-0889; Fax: 734-763-4581;
E-mail: rtrievel@umich.edu.
3 The abbreviations used are: AdoMet, S-adenosylmethionine; AdoHcy, S-ad-
enosylhomocysteine; KMT, lysine methyltransferase; ITC, isothermal titra-
tion calorimetry; CH–O, carbon-oxygen hydrogen bond; BisTris, 2-[bis(2-
hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 41, pp. 31849–31858, October 8, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31849
Y245A and Y305F, which change its product specificity. The
Phe/Tyr switch mutant Y305F alters SET7/9 product specificity
from a mono- to dimethyltransferase (6), whereas the Y245A
substitution converts the enzyme into a trimethyltransferase
with weak monomethyltransferase activity (11). These mutants
have been the subjects of numerous molecular modeling simu-
lations that have led to various models to explain their distinct
product specificities (22–26). However, the lack of structural
data for the SET7/9 Y245A and Y305F mutants in complex with
cognate methylated peptides has hindered our understanding
of the mechanisms that define the respective product specific-
ities of these mutants. Moreover, these structures would yield a
framework for visualizing the mono-, di-, and trimethylation
reactions catalyzed by SET domain KMTs.
To gain insight into the molecular basis of their product
specificities, we have determined high resolution crystal
structures of the SET7/9 Y245A and Y305F mutants in com-
plex with peptides of the TATA box-binding protein-associ-
ated factor TAF10 bearing the Lys-189 methylation site in
unmodified (K189), monomethylated (K189me1), dimethyl-
ated (K189me2), and trimethylated (K189me3) states. The
structures and accompanying biochemical data support a
model whereby changes in the occupancy or position of water
molecules in the active site are critical in establishing the prod-
uct specificities of the SET7/9 Y245A and Y305F mutants.
Together, our results provide new insights into the mechanisms
that govern SET domain product specificity and provide step-
wise snapshots of the lysine mono-, di-, and trimethyl transfer
reactions catalyzed by KMTs.
EXPERIMENTAL PROCEDURES
Cloning, Expression, and Purification of the SET7/9 Mutants—
The Y245A and Y305F mutants were introduced into the pHIS2
SET7/9 expression vector encoding residues 110–366 (27) via
QuikChange site-directed mutagenesis (Stratagene) and were
verified by dideoxy DNA sequencing. The plasmids encoding
wild type (WT) SET7/9 and the Y245A and Y305F mutants
were transformed into Rosetta2 DE3 cells (Novagen) and were
expressed as described previously (27, 28). In the course of
characterizing WT SET7/9, we observed that the enzyme co-
purified with AdoMet or another contaminant that resulted in
technical difficulties in the isothermal titration calorimetry
(ITC) experiments and co-crystallization trials with the TAF10
peptides. To overcome this problem, a denaturation and refold-
ing step was inserted in the purification scheme. The denatur-
ation and refolding protocol involved adding 6 M guanidine
HCl, 50 mM sodium phosphate, pH 7.5, and 500 mM NaCl to the
protein while it was immobilized on a nickel-Sepharose column
(GE Healthcare). The column was washed with this buffer, fol-
lowed by a solution of 6 M urea, 50 mM sodium phosphate, pH
7.5, 150 mM NaCl, and 10 mM 2-mercaptoethanol to remove the
cofactor from the denatured enzyme. A reverse gradient from 6
to 0 M urea was then performed in the same buffer to refold the
protein, which was subsequently eluted from the column using
a linear gradient of 0–500 mM imidazole in 50 mM sodium
phosphate, pH 7.5, 500 NaCl, and 10 mM 2-mercaptoethanol.
The refolded protein was digested with tobacco etch virus pro-
tease (29) during dialysis against 20 mM Tris, pH 7.5, 150 mM
NaCl, and 5 mM 2-mercaptoethanol and then purified using a
Superdex 200 gel filtration column (GE Healthcare). Protein
concentration was determined by its absorbance at 280 nm.
Synthetic Peptides—The TAF10 peptides bearing K189,
K189me1,
K189me2,
and
K189me3
(sequence,
acetyl-
SKSK189DRKYTL-amide) and a biotinylated TAF10 peptide
(sequence,
acetyl-SKSK189DRKYTLT(K-EZLink-S-S-biotin)-
amide) were synthesized and purified by New England Peptide,
Inc. Peptide concentrations were measured using the absorb-
ance of their tyrosine residue at 274 nm.
Crystallization and Data Collection—Crystals were pro-
duced by hanging drop vapor diffusion by mixing the crystalli-
zation solution in a 1:1 ratio with 10–20 mg/ml SET7/9, 1 mM
S-adenosylhomocysteine (AdoHcy), and 1.0–3.0 mM unmodi-
fied or methylated TAF10-K189 peptide in 20 mM Tris, pH 8.0,
100 mM NaCl, and 2 mM tris(2-carboxyethyl)phosphine. Crys-
tals were obtained at 20 °C in either 1.8–2.0 M (NH4)2SO4, with
0.1 M BisTris, pH 6.2–6.6, or in 0.95–1.1 M sodium citrate with
100 mM imidazole pH 8.0–8.4. In both crystallization condi-
tions, the final pH values were between pH 8.0 and 9.0. Crystals
in the (NH4)2SO4 condition were typically flash-frozen in the
mother liquor containing 25–30% glycerol, and the crystals in
the citrate condition were frozen in 1.6 M sodium citrate. Data
were collected at the Advanced Photon Source beamlines
21-IDG (LS-CAT) and 23-IDD (GM/CA-CAT). Images were
indexed, integrated, and scaled using HKL2000 (30). Structures
of the mutants were solved by molecular replacement using
MOLREP (31) with the coordinates of a previously reported
SET7/9 ternary complex used as the search model (Protein
Data Bank code 2F69). Successive rounds of model building and
refinement were carried out using Coot (32) and REFMAC (33),
respectively. The geometry of the models were verified by Mol-
Probity (34). Simulated annealing omit maps were calculated
using CNS (35) with the peptide and cofactor removed to elim-
inate model bias in the active site. Structural figures were ren-
dered using PyMOL (Schro¨dinger, LLC.).
Fluorescent Methyltransferase Assay—A coupled fluorescent
methyltransferase assay was used to measure the kinetic
parameters of WT SET7/9 and the Y245A and Y305F mutants
as reported previously, with the exception that 50–150 nM
enzyme, 100 M AdoMet, and varying concentrations of TAF10
peptide substrate were used (27, 36). Assays were performed in
triplicate, and a homocysteine calibration curve was used to
calculate the initial velocities. Kinetic parameters were calcu-
lated by plotting the velocities versus peptide concentration and
by fitting the Michaelis-Menten equation to the data via non-
linear regression using Prism 5.0 (GraphPad). In cases where
the Km value was beyond the measurable range of the assay, the
kcat/Km value was determined as described previously (7).
Calorimetry Experiments—ITC was performed at 20 °C using
a MicroCal VP-ITC calorimeter (GE Healthcare) with 0.12 mM
protein and 4 mM AdoHcy in 20 mM sodium phosphate, pH 7,
and 100 mM NaCl with 1.5 mM peptide as the injectant. Data
were processed, and equilibrium dissociation constants (KD)
and curve fitting errors were calculated from the binding iso-
therms using Origin 7.0 (OriginLab Corp.). WT SET7/9 and the
Y245A and Y305F mutants displayed ligand:protein binding
stoichiometries (N values) between 0.8 and 1.0, demonstrating
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
that WT SET7/9 and its mutants were properly refolded due to
their ability to bind peptides in an 1:1 molar ratio.
TLC Product Analysis—Methyltransferase assays were per-
formed in triplicate at 37 °C with the biotinylated TAF10 pep-
tide (0.5 mM) for 30 min in the presence of WT SET7/9 (3
pmol), Y305F mutant (6 pmol), or the Y245A mutant (100
pmol). Assays contained 100 mM HEPES, pH 8.0, 50 mM NaCl,
1.0 mM [methyl-3H]AdoMet (diluted to a specific activity of 0.2
Ci/nmol with purified AdoMet (37)), 15 M Sulfolobus solfa-
taricus AdoHcy hydrolase (36), and 2 units of adenosine deami-
nase (Roche Applied Science) in a final volume of 20 l. The
reactions were terminated by addition of an equal volume of
200 mM MES, pH 5.1, prior to addition of a 2-fold molar excess
of immobilized avidin resin (UltraLink; Pierce). Biotinylated
peptides were allowed to bind at room temperature for 30 min,
and the resin was then collected by centrifugation (9000 g).
The resin was washed three times with 300 mM NaCl, and the
peptide was eluted overnight from the avidin resin by cleavage
of the disulfide bond in the linker of the peptide using 10 mM
tris(2-carboxyethyl)phosphine (Thermo Scientific). The resin
was incubated with additional 10 mM tris(2-carboxyethyl)phos-
phine the following day until the radiolabel was essentially
removed from the resin. The recovered peptides were hydro-
lyzed by incubation with 6 M HCl at 110 °C for 24 h. Subsequent
steps in measuring the radiolabel incorporated into the mono-,
di-, and trimethyl-lysine products were performed as reported
previously (8).
RESULTS
Functional Analysis of the SET7/9
Y305F Mutant—Prior studies of
SET7/9 by Zhang et al. (6) reported
that mutation of the Phe/Tyr switch
residue Tyr-305 to a phenylalanine
alters its product specificity from a
mono- to dimethyltransferase. We
verified these findings by demon-
strating that WT SET7/9 mono-
methylated the TAF10-K189 pep-
tide, whereas the Y305F mutant
mono- and dimethylated this sub-
strate, as demonstrated by mass
spectrometry (data not shown). We
next examined whether the Y305F
substitution altered the affinity of SET7/9 for the TAF10-K189
peptides using ITC (Fig. 1). A comparison of the KD values
revealed that SET7/9 Y305F bound the TAF10-K189 and
TAF10-K189me1 peptides 4- and 6-fold more tightly, respec-
tively, than the WT enzyme, whereas this mutant displayed a
substantially diminished affinity for the TAF10-K189me2 pep-
tide (Table 1). Although the WT enzyme and the Y305F mutant
exhibited discernable differences in their affinities for the
unmodified and monomethylated peptides, these variations are
modest and cannot account for their distinct product specific-
ities, suggesting that a kinetic effect during methylation may be
responsible.
To investigate this possibility, we characterized the kinetic
parameters of WT SET7/9 and the Y305F mutant using the
TAF10 peptides as substrates. Both enzymes methylated the
unmodified peptide with comparable kcat and Km values (Table
2). In analyzing the kinetic parameters for the methylation
of the monomethylated peptide by SET7/9 Y305F, we found
that this substrate displayed an elevated Km value that was
beyond the measurable range of the assay due to its limited
solubility. In this case, we measured the catalytic efficiency
(kcat/Km) for the methylation of this peptide and found that it
was methylated 15-fold less efficiently than the unmodified
peptide by SET7/9 Y305F. Given the fact that the Y305F mutant
exhibited a higher binding affinity for the TAF10-K189me1
peptide than the WT enzyme (Table 1), the kinetic data suggest
that a step in the reaction pathway following substrate binding
limits the catalytic efficiency of this mutant.
We next examined whether the Y305F mutant dimethylated
the TAF10-K189 peptide via a processive or a distributive
mechanism. In a processive mechanism, the methyl-lysine
substrate would remain bound to the enzyme during successive
methyl transfer reactions; thus, the concentration of an inter-
mediate, such as monomethyl-lysine, cannot exceed the en-
zyme concentration during the assay. In a distributive mecha-
nism, the intermediates are released into solution where they
accumulate prior to the next round of methylation, resulting in
an intermediate concentration that is greater than that of the
enzyme. Using a radiometric TLC assay and a biotinylated
TAF10 peptide, we quantified the amounts of monomethylated
products generated by the WT SET7/9 and the Y305F mutant
FIGURE 1. ITC analysis of WT SET7/9 and the Y245A and Y305F mutants. Representative titrations and
binding isotherms for the calorimetry experiments are shown for the unmodified TAF10-K189 peptide titrated
into WT SET7/9 (left plot), SET7/9 Y305F (middle plot), and SET7/9 Y245A (right plot). In each panel, the ITC
titration experiment (upper panel) is shown with the binding isotherms (lower panel) fit to a single binding site
model.
TABLE 1
Analysis of the binding affinity of WT SET7/9 and its catalytic
mutants for unmodified and methylated TAF10 peptides
SET7/9
TAF10 peptide
KD
a
M
WT
K189
4.9 0.20
WT
K189me1
4.0 0.36
Y305F
K189
1.3 0.10
Y305F
K189me1
0.62 0.065
Y305Fb
K189me2
70
Y245A
K189
4.0 0.25
Y245A
K189me1
3.3 0.10
Y245A
K189me2
5.8 0.22
Y245A
K189me3
11 0.28
a Curve fitting errors were calculated from the binding isotherms.
b An estimate of the affinity is reported due to weak peptide binding.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31851
(Table 3). The data demonstrate that comparable amounts of
monomethyl-lysine were generated when the quantity of
enzyme usedistakenintoaccount,inagreementwiththeirsimilar
turnover numbers for the TAF10-K189 peptide (Table 2). The
Y305F mutant also produced small but measurable quantities of
radiolabeled dimethyl-lysine product that were substantially
smaller than the amount of monomethyl-lysine generated.
Therefore, the TLC data are consistent with a distributive
mechanism for dimethylation by the Y305F mutant because the
amount of monomethyl-lysine produced exceeded the quantity
of enzyme used in the assay.
Structures of WT SET7/9 and the Y305F Mutant in Complex
with Unmodified and Methylated TAF10 Peptides—To deter-
mine the mechanism by which the Y305F substitution alters the
product specificity of SET7/9, we determined the crystal struc-
tures of this mutant bound to AdoHcy and TAF10-K189,
TAF10-K189me1, and TAF10-K189me2 peptides and com-
pared these to the structures of the WT SET7/9AdoHcy
TAF10-K189 complex (supplemental Table 1). The structures
of these complexes were determined to 1.85 Å or higher reso-
lution, permitting unambiguous modeling of the K189 side
chains in the active site of the enzyme based on simulated
annealing omit maps (Fig. 2). The ternary complexes of the WT
and the Y305F mutant superimpose with overall root mean
square differences of less than 0.3 Å for all aligned atoms, indi-
cating that neither the Y305F mutation nor the binding of the
various TAF10-K189 peptides results in substantial changes in
its overall structure.
An inspection of the active sites of the SET7/9 WT and
Y305F complexes illustrates the binding modes of the unmod-
ified and methylated forms of K189 in the TAF10 peptides (Fig.
2, A–D). The K189 side chain binds in an extended all trans
conformation in a deep pocket, termed the lysine binding chan-
nel, that is composed of Tyr-245, Thr-266, Leu-267, Ser-268,
Tyr-305, Tyr-335, and Tyr-337 (not shown for clarity) (Fig. 2A).
These residues interact with the aliphatic portion of the K189
side chain primarily through van der Waals contacts. The lysine
binding channel connects to the AdoMet-binding site on the
opposite face of the catalytic domain via an oxygen-lined
methyl transfer pore (38). During catalysis, the methyl group of
the cofactor is positioned within the methyl transfer pore
for the SN2 reaction with the -amino group of the lysine or
methyl-lysine substrate (see below).
To lower the activation barrier for this reaction, the lysine
-amine nucleophile is aligned for methyl transfer through a
hydrogen bond network within the active site. In the WT
enzyme, the K189 -amino group hydrogen bonds to the
hydroxyl group of Tyr-245 as well as to two water molecules
(Fig. 2A). One of the water molecules (termed water 1), is coor-
dinated in a solvent pocket, through hydrogen bonds to the
carbonyl oxygens of Gly-292 and Ala-295 and to the hydroxyl
group of the Phe/Tyr switch residue Tyr-305. This solvent
pocket is structurally conserved in SET domain KMTs and has
an important role in defining product specificity through the
adjacent Phe/Tyr switch residue, as shown in our prior studies
of the human H4K20 methyltransferase SET8 (8). The other
water molecule is bound within the methyl transfer pore
between the lysine substrate and the thioether sulfur atom of
AdoHcy through hydrogen bonds to Tyr-245, Asn-265, and
His-293 in SET7/9 and the TAF10-K189 -amino group. This
water is not observed in other structures of SET7/9 ternary
complexes and may represent the approximate position that
the AdoMet methyl group occupies in the methyl transfer pore
in the Michaelis complex.
In structures of the Y305F ternary complexes, the K189,
K189me1, and K189me2 side chains also adopt extended trans
side chain geometries within the lysine binding channel that are
stabilized via hydrogen bonding to Tyr-245 in the enzyme (Fig.
2, B–D). The orientations of the K189me1 and K189me2 side
chains are further maintained through carbon-oxygen (CH–O)
hydrogen bonding between the methyl groups and oxygen
atoms within the vicinity of the methyl transfer pore, as
reported previously in other SET domain KMT structures (8,
10, 38). A superimposition of the SET7/9 WT and Y305F com-
plexes underscores the similarity of the lysyl binding conforma-
tions (Fig. 2E). However, there are notable differences in the
hydrogen bond patterns and occupancy of water 1 within the
solvent pocket in the Y305F mutant compared with the WT
enzyme. Specifically, the Y305F substitution results in the loss
of one hydrogen bond to water 1 in the structures of the TAF10-
K189 and TAF10-K189me1 complexes (Fig. 2, B and C). In con-
trast, water 1 is absent in TAF10-K189me2 complex, and the
vacated solvent pocket is occupied by one of the methyl groups
TABLE 2
Kinetic parameters of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
TAF10 peptide substrate
Km
a
kcat
a
kcat/Km
a
M
min1
M1 min1 103
WT
K189
160 17
17 0.62
110 17
Y305F
K189
88 5.0
17 0.30
190 11
Y305Fb
K189me1
11 0.50
Y245A
K189
200 35
0.53 0.04
2.6 0.47
Y245A
K189me1
210 23
5.9 0.23
28 3.3
Y245A
K189me2
400 29
6.5 0.16
15 1.2
a Curve fitting errors were calculated from the hyperbolic fits for the Michaelis-Menten equation.
b Km and kcat were not determined because the Km value was beyond the measurable range; therefore, the kcat/Km value is reported.
TABLE 3
Product analysis of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
Quantity
of enzyme
Measured
product
Amount of
product formeda
nmol
nmol
WT
0.003
Kme1
0.65 0.07
Y305F
0.006
Kme1
1.5 0.49
Kme2
0.033 0.009
Y245A
0.100
Kme1
0.80 0.22
Kme2
0.39 0.021
Kme3
0.076 0.019
a Standard deviation was calculated from triplicate measurements.
Lysine Methylation by SET7/9 Mutants
31852
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
of the dimethyl -amine (Fig. 2D). This methyl group forms a
3.5-Å CH–O hydrogen bond to the carbonyl oxygen of Ala-295,
further stabilizing the binding of the dimethyl-lysine side chain.
A homologous dimethyl-lysine-binding mode and CH–O
hydrogen bond was observed in our prior structural studies of
the SET8 Y334F Phe/Tyr switch mutant that confers an analo-
gous change in product specificity from a mono- to a dimeth-
yltransferase (8). A structural alignment of the active sites of the
SET7/9 Y305F and SET8 Y334F mutants bound to cognate
dimethylated peptides illustrates that the coordinates of the
dimethyl-lysyl side chains are virtually superimposable, with
one methyl group oriented toward the methyltransfer pore and
the second positioned within the vacant solvent pocket (Fig.
2F). Taken together, the structures of the SET7/9 Y305F com-
plexes and the similarities in the dimethyl-lysine conforma-
tions in the SET7/9 Y305F and SET8 Y334F mutants imply that
the Phe/Tyr switch governs product specificity through a con-
served mechanism whereby it indirectly influences the binding
modes of the methyl-lysine side chain by modulating the affin-
ity of the water molecule (water 1) bound in the solvent pocket.
Biochemical Characterization of
the SET7/9 Y245A Mutant—Previ-
ous studies by Xiao et al. (11)
reported that the Y245A mutation
yields an unusual change in the
product
specificity
of
SET7/9,
converting the enzyme to a trimeth-
yltransferase with weak monometh-
yltransferase activity. We deter-
mined that the SET7/9 Y245A could
mono-, di-, and trimethylate the
TAF10-K189 peptide by mass spec-
trometry (data not shown) and TLC
(Table 3), confirming the earlier
studies of Xiao et al. (11). ITC
analysis revealed that the Y245A
mutant displayed comparable KD
values
for
the
unmodified
and
methylated TAF10-K189 peptides
(Fig. 1), although its affinity for the
trimethylated peptide was modestly
diminished in comparison with the
other peptides (Table 1). The ITC
data demonstrate that the Y245A
mutant
bound
the
unmodified,
mono-,
and
dimethylated
sub-
strates with equivalent affinities,
suggesting that a kinetic effect or
a structural alteration in the active
site may be responsible for its
diminished
activity
toward
un-
modified substrates.
To gain further insight into its
peculiar
product
specificity,
we
characterized the kinetic properties
of
the
SET7/9
Y245A
mutant.
Steady state analysis demonstrated
that this mutant displayed similar
Km values for the unmodified, mono- and dimethylated TAF10
peptides (Table 2). However, the turnover number for the
TAF10-K189 peptide was diminished over 10-fold versus the
methylated peptides and was reduced 30-fold versus the WT
enzyme, in agreement with the weak monomethyltransferase
activity reported by Xiao et al. (11). In addition, we investigated
whether this mutant catalyzes lysine trimethylation via a pro-
cessive or distributive mechanism as described for SET7/9
Y305F. The TLC data illustrate that the mono- and dimethyl-
lysine intermediates accumulated at quantities greater than
that of the enzyme used in the assay, indicating that SET7/9
Y245A obeys a distributive mechanism, analogous to the Y305F
mutant (Table 3).
Structures of SET7/9 Y245A Bound to Unmodified and Meth-
ylated TAF10 Peptides—To elucidate the mechanism underly-
ing its unusual product specificity, we determined the crystal
structures of SET7/9 Y245A in complex with AdoHcy and
unmodified, mono-, di-, and trimethylated TAF10 peptides
(supplemental Table 1). These complexes superimpose with
the structure of the WT SET7/9AdoHcyTAF10-K189 com-
FIGURE 2. Crystal structures of WT SET7/9 and the Y305F mutant in complex with AdoHcy and unmodi-
fied and methylated TAF10 peptides. A, active site of WT SET7/9 bound to AdoHcy and TAF10-K189 and the
active site of SET7/9 Y305F bound to AdoHcy and TAF10-K189 (B), TAF10-K189me1 (C), and TAF10-K189me2
peptides (D). A–D, the K189 residue in the TAF10 peptide is depicted with yellow carbon atoms with the
corresponding Fo – Fc simulated-annealing omit maps (contoured at 2.5 ) shown. AdoHcy is colored with
green carbon atoms, and SET7/9 carbon atoms are colored gray, with the exception of Tyr-305/Y305F
(magenta) and Tyr-245 (cyan). Gly-264 to Ser-268 are shown, and the main chain atoms of Gly-292 to Ala-295
are depicted for clarity. Red dashed lines indicate conventional hydrogen bonds, and blue dashed lines indicate
CH–O hydrogen bonds. Cyan dashed lines indicate weak CH–O hydrogen bonds between 3.5 and 3.7 Å in
length. The water molecule in the solvent binding pocket is numbered 1. E, overlay of the active sites of the WT
enzyme (light blue carbon atoms) and the Y305F mutant (green, yellow, and pink carbon atoms correspond to
K189, K189me1, and K189me2, respectively). Red dashed lines indicate coordination of the waters in the struc-
turesoftheY305Fmutant,andthelightbluedashedlineindicateshydrogenbondingthatonlyoccursintheWT
enzyme. F, superimposition of the active sites of the SET7/9 Y305FAdoHcyTAF10-K189me2 and SET8
Y334FAdoHcyH4K20me2 (Protein Data Bank code 3F9X) complexes shown with green and yellow carbon
atoms, respectively.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31853
plex with root mean squared differences of less than 0.4 Å for all
aligned atoms, indicating that the Y245A mutant does not per-
turb the overall structure of the enzyme. Simulated annealing
omit maps illustrate that K189 side chains are bound within the
lysine binding channel through hydrogen bonds and van der
Waals contacts (Fig. 3, A–D), although the interactions and
binding modes are distinct from those in the complexes of WT
SET7/9 and the Y305F mutant (Fig. 2, A–D). In the unmodified
TAF10 peptide complex, the K189 -amino group forms a weak
hydrogen bond with the carbonyl oxygen of Gly-264 (Fig. 3A),
whereas the -amino groups of K189me1 and K189me2 hydro-
gen bond to the hydroxyl group of Tyr-305 in the mono- and
dimethylated peptide complexes (Fig. 3, B and C). The confor-
mations of the K189me1 and K189me2 side chains are further
stabilized by water-mediated hydrogen bonding and through
CH–O hydrogen bonding to their methyl groups. In the
TAF10-K189me3 peptide complex, the trimethyl-lysine side
chain is coordinated exclusively through direct and water-me-
diated CH–O hydrogen bonds to its methyl groups because the
quaternary -ammonium cation cannot engage in hydrogen
bonding (Fig. 3D).
A structural alignment of the four SET7/9 Y245A complexes
illustrates distinct binding modes for the unmodified versus the
methylated K189 side chains, highlighting the selectivity of this
mutant for methylated substrates. The side chains of K189me1,
K189me2, and K189me3 roughly overlay with their respective
-amino groups superimposed and adopt slightly kinked con-
formations (Fig. 3E), as opposed to the extended trans geome-
try of the unmodified and methylated lysines in the complexes
of the WT enzyme and the Y305F mutant (Fig. 2, A–D). Con-
versely, the unmodified K189 side chain does not superimpose
with its methylated counterparts and is oriented in an alterna-
tive configuration due to its hydrogen bonding to Gly-264 (Fig.
3, A and E). An overlay of the structures of the WT enzyme and
Y245A mutant bound to the unmodified TAF10 peptide illus-
trates that the side chains of K189 do not superimpose and that
the K189 -amino group appears to be misaligned with AdoHcy
in the Y245A complex (Fig. 3F). This suboptimal alignment
may explain the diminished kcat value of SET7/9 Y245A mutant
toward substrates with unmodified lysines (Table 2).
A comparison of the structures of the SET7/9 Y245A and
Y305F complexes yields a molecular explanation for the differ-
ent product specificities of these two mutants. In the SET7/9
Y305F complexes, Tyr-245 aligns the K189 -amino group for
methyl transfer through hydrogen bonding to its hydroxyl
group (Fig. 2, B–D). Conversely, in the Y245A mutant, the
K189me1 and K189me2 -amino groups are oriented through
hydrogen bonding to Tyr-305 (Fig. 3, B and C). These distinct
hydrogen bond patterns impart differences in the conforma-
tions of the lysyl side chains due to the relative orientations of
Tyr-245 and Tyr-305 in the lysine binding channel. Specifically,
the kinked conformation adopted by the K189me1 and
K189me2 side chains in the Y245A complexes (Fig. 3, B and C)
may contribute to the differences in the turnover numbers of
this mutant versus those of the WT enzyme and the Y305F
mutant (Table 2). In addition, the dimethyl -amino group of
the K189me2 side chain binds in distinct orientations in the
Y245A and Y305F mutants due to their hydrogen bonding to
FIGURE 3. Crystal structures of the SET7/9 Y245A mutant in complex with
AdoHcy and unmodified and methylated TAF10 peptides. Active site of
SET7/9 Y245A bound to AdoHcy and TAF10-K189 (A), TAF10-K189me1 (B),
TAF10-K189me2 (C), and TAF10-K189me3 peptides (D). A–D, Fo – Fc simulated-
annealing omit maps (contoured at 2.5 ) for the unmodified and methylated
K189 side chains are illustrated. The residues and hydrogen bonds in each com-
plexarecoloredasinFig.2.Thewatermoleculesinthelysinebindingchannelsof
the Y245A complexes are numbered 1–4, as described in the text. E, superimpo-
sition of the active sites of the Y245A complexes bound to the four methylated
statesofTAF10-K189(unmodified,mono-,di-,andtrimethylatedshowningreen,
yellow, pink, and blue, respectively). F, overlay of WT SET7/9 (blue carbons) and
SET7/9 Y245A (green carbons) in complex with TAF10-K189. Waters correspond-
ing to the WT and Y245A structures are colored cyan and green, respectively.
G, alignment of the active sites of the Y245A (yellow carbons) and Y305F (green
carbons) mutants in complex with the TAF10-K189me2 peptide. Hydrogen
bonds from the Y305F structure are shown as green dashed lines, and waters and
hydrogen bonds in the Y245A structure are shown in yellow and orange,
respectively.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
Tyr-305 and Tyr-245, respectively (Fig. 3G). In the Y305F
mutant, hydrogen bonds to the dimethyl -amino group cou-
pled with steric constraints in the lysine binding channel pre-
vent the K189me2 side chain from undergoing a conforma-
tional change that is conducive to trimethylation (Fig. 2D),
consistent with its dimethyltransferase activity. However, in the
Y245A mutant, the alanine substitution enlarges the diameter
of the lysine binding channel, accommodating trimethyl-lysine
(Fig. 3D). In addition, the larger diameter would permit the
dimethyl-lysine substrate to undergo the conformational reor-
ganization necessary to align the -amino group in a productive
geometry for trimethylation.
A major difference in the active site of the Y245A mutant
versus the other SET7/9 structures is the presence of several
water molecules bound in the cavity generated by the Y245A
mutation. In the structure of the Y245A mutant bound to
TAF10-K189, three water molecules (waters 2–4) occupy this
cavity and are arranged in a triangular geometry (Fig. 3A). In
addition, water 1 shifts 1.6 Å from its position in the solvent
pocket toward water 2 to which it forms a hydrogen bond (Fig.
3, A and E). The shift in water 1 was unexpected given its con-
served orientation in the solvent pocket of the SET7/9 WT and
Y305F complexes (Fig. 2, A–C) as well as in the structures of
other SET domain KMTs (8). This displacement is presumably
related to the alternative conformation of the K189 side chain
whose -amino group is too distant (4.3 Å) to form a productive
hydrogen bond to water 1. Conversely, in the Y245A complexes
bound to TAF10-K189me1 and TAF10-K189me2, water 1
remains tightly bound in the solvent pocket through hydrogen
bonds to Tyr-305 hydroxyl group, the carbonyl oxygens of Gly-
292 and Ala-295, and the K189 -amino group (Fig. 3, B, C, and
E), analogous to its binding in the WT enzyme (Fig. 2A). How-
ever, in the TAF10-K189me3 complex, one of the methyl
groups of the trimethyl -ammonium cation is oriented into the
solvent pocket (Fig. 3D), similar to the dimethyl-lysine binding
mode observed in the Y305F mutant (Fig. 2D). The binding of
the methyl group in the solvent pocket displaces water 1 by 3.2
Å relative to its position in the TAF10-K189me1 complex (Fig.
3E), thereby avoiding a steric clash with the trimethylated
-ammonium group. Variations in the occupancy of water 2 are
also seen in the different Y245A structures. Water 2 is bound in
similar orientations in the active site of the unmodified and
monomethylated peptide complexes but is absent in the di- and
trimethylated peptide complexes due to the binding of a methyl
group in this position (Fig. 3, A–E). In summary, the changes in
the positions or occupancies of waters 1 and 2 correlate with the
binding modes of the unmodified and methylated K189 within
the active site of the Y245A mutant.
Catalytic Models of Lysine Multiple Methylation by SET7/9
Y245A, and Y305F—The structures of the SET7/9 complexes
reported here offer a prime opportunity to generate stepwise
models for lysine mono-, di-, and trimethylation by a SET
domain KMT. We modeled the AdoMet-bound Michaelis
complexes by superimposing the SET7/9 product complexes
with the previously reported structure of the SET7/9-AdoMet
binary complex (Fig. 4) (39). The conformations of the mono-
and dimethyl -amino groups in the Michaelis complexes were
inferred from the coordinates of the corresponding dimethyl-
and trimethyl-lysine products, respectively. In addition, we
modeled the -amino group in a deprotonated state with its
hydrogen atoms oriented toward the hydrogen bond acceptors
that align the lysyl side chain for methylation. As a basis for this
comparison, we first modeled the monomethylation reaction
catalyzed by WT SET7/9 (Fig. 4A). In the substrate ternary
complex, the lysine -amine is aligned with the methyl group
and sulfonium cation of AdoMet through a hydrogen bond to
the Tyr-245 hydroxyl group and water 1 in the solvent pocket.
The values of the reaction distance and angle are 2.8 Å and 153°,
respectively, in approximate agreement with the linear geome-
try of a SN2 methyl transfer reaction calculated in other mod-
eled substrate complexes (8, 10). In the product complex, the
monomethyl-lysine side chain is bound in an extended confor-
mation with its methyl group oriented within the methyl trans-
fer pore, thereby obstructing AdoMet binding. Furthermore,
water 1 remains tightly coordinated in the solvent pocket
through four hydrogen bonds to Gly-292, Ala-295, Tyr-305,
and the monomethyl -amino group. These interactions hinder
the dissociation of water 1 and the related rearrangement of the
monomethyl-lysine side chain required for a second methyl
transfer reaction, explaining why the WT enzyme cannot cata-
lyze di- and trimethylation. These findings concur with the
FIGURE 4. Catalytic models of the methyl transfer reactions catalyzed by
WT SET7/9 and the Y305F mutant. A, monomethylation of TAF10-K189 by
the WT enzyme. The reaction scheme depicts the modeled substrate ternary
complex (left) and the product complex (right) for the transfer of the methyl
group from AdoMet (green carbon atoms) to K189 in TAF10 (yellow carbons),
yielding AdoHcy and K189me1. The red arrow indicates the direction of the
nucleophilic attack of the deprotonated -amino group on the AdoMet
methyl group. The transferred methyl group is colored green, and the white
atoms represent the hydrogens of the -amino group. Hydrogen bonds
and residues in the enzyme active site are illustrated as in Fig. 2. The
reaction distance and angle are labeled in red. B and C, models of the
Y305F mutant for the first methyl transfer reaction with TAF10-K189 (B)
and second methyl transfer reaction with TAF10-K189me1 (C). Color
schemes are the same as in A.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31855
model for SET7/9 product specificity reported in previous
structural and functional studies (6, 11).
Similar reaction geometry is observed in the model for the
monomethyl transfer reaction catalyzed by SET7/9 Y305F.
Hydrogen bonds from the Tyr-245 hydroxyl group and water 1
align the lysine -amino group with the AdoMet methyl group
at a distance of 2.1 Å and an angle of 160°, equivalent to those
measured in the Michaelis complex of the WT enzyme (Fig. 4, A
and B). In the product complex, monomethyl-lysine adopts an
extended trans configuration analogous to that in the WT
enzyme. For dimethylation to occur, the monomethyl-lysine
must undergo a conformational change in which its methyl
group is rotated out of the methyl transfer path with AdoMet.
The structure of the Y305F mutant bound to the dimethylated
TAF10 peptide (Fig. 2D) implies that this rearrangement occurs
through the dissociation of water 1 due to the loss of the Tyr-
305 hydrogen bond in the solvent pocket. The dissociation of
water 1 would enable the monomethyl-lysine side chain to
adopt an alternative conformation through a rotation about its
C–N bond, projecting the methyl group into the solvent
pocket (Fig. 4C). This rotation reorients the methyl group out of
the methyl transfer path while realigning the monomethyl -
amino group for a second methylation reaction through a direct
hydrogen bond to the Tyr-245 hydroxyl group and a CH–O
hydrogen bond between its methyl group and Ala-295. The
modeled reaction geometry for monomethyl-lysine substrate
complex of the Y305F mutant (2.7 Å, 160°) is equivalent to that
of the first methyl transfer reaction in SET7/9 Y305F. These
geometries concur with our previous models for mono- and
dimethylation catalyzed by SET8 Y334F (8), illustrating that the
orientation of a methyl group into the solvent pocket is a con-
served feature of SET domain KMTs that catalyze multiple
methylation.
In addition, we modeled the methyl transfer reactions cata-
lyzed by SET7/9 Y245A (Fig. 5). In the model of the lysine sub-
strate complex, the -amino group is aligned for methyl transfer
by a hydrogen bond to the carbonyl oxygen of Gly-264, result-
ing in a short reaction distance (2.3 Å) and a suboptimal reac-
tion angle (141°) with the methyl group of AdoMet (Fig. 5A).
This misalignment appears to be a direct consequence of the
Y245A mutation that abolishes hydrogen bonding to the -
amino group, illustrating that the suboptimal orientation of the
-amine likely contributes to the diminished activity of this
mutant toward unmodified substrates (11). Conversely, in the
modeled monomethyl-lysine substrate complex for SET7/9
Y245A, Tyr-305 hydrogen bonds to the -amino group, aligning
it for methyl transfer (Fig. 5B). In addition, CH–O hydrogen
bonds to the monomethyl-lysine methyl group and the dissoci-
ation of water 2 from the active site also contribute to reposition-
ing the -amino group for dimethylation. Collectively, these inter-
actions orient the -amine in a reaction angle of 165° that is more
conducive to methyl transfer. However, the reaction distance for
dimethylation is 0.6 Å longer than that in the corresponding
Y305F model because Tyr-305 is positioned further from
AdoMet than Tyr-245 (Figs. 4C and 5B).
In the third methyl transfer reaction catalyzed by SET7/9
Y245A, the lone pair of electrons of the dimethyl-lysine -
amino group acts as the nucleophile and thus cannot engage in
hydrogen bonding. The structure of the trimethyl-lysine prod-
uct complex (Fig. 3D) implies that the dimethyl -amine is
aligned via CH–O hydrogen bonds to its methyl groups, as
shown in the model of the Michaelis complex for this reaction
(Fig. 5C). These CH–O hydrogen bonds restrain the orienta-
tion of the -amino group and position one of the methyl
groups into the solvent pocket, displacing water 1 as discussed
earlier (Fig. 3, D and E). These interactions cumulatively align
the -amino group and AdoMet methyl group with a reaction
distance of 3.0 Å and angle of 162° (Fig. 5C). Taken together, the
models of the substrate complexes for SET7/9 Y245A suggest
that CH–O hydrogen bonds play an increasingly important role
in aligning the methylated -amino group in successive rounds
of methyl transfer.
DISCUSSION
The structural and functional characterization of the
SET7/9 Y245A and Y305F mutants presented here yields
new insights into the mechanism underlying the product
specificity of SET domain KMTs. Importantly, it resolves a
general paradox concerning this specificity. How does the
active site constrain the motion of the lysine -amino group
to align it for methyl transfer with AdoMet, while providing
adequate volume to accommodate the mono-, di-, and tri-
methylated lysine side chain generated during multiple
methyl transfer reactions? The structures of the Y305F and
Y245A mutants resolve this paradox, illustrating that alter-
ations in the positions or occupancies of water molecules
within their active sites generate the space required to
FIGURE 5. Models for the methyl transfer reactions catalyzed by the
SET7/9 Y245A mutant. Models of the Y245A mutant for the first methyl
transfer reaction with TAF10-K189 (A), the second methyl transfer reaction
with TAF10-K189me1 (B), and the third methyl transfer reaction with TAF10-
K189me2 (C). Residues and hydrogen bonds are depicted as in Fig. 3.
Lysine Methylation by SET7/9 Mutants
31856
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
accommodate the multiply methylated -amine produced
during successive catalytic cycles. Minor perturbations in
the side chains of certain active site residues, such as Tyr-
305, are also observed in alignments of the WT enzyme and
the Y245A and Y305F complexes, although these changes are
modest compared with the displacement or dissociation of
the water molecules in the active site. These findings suggest
that the waters function as transient place holders that facil-
itate the SN2 methyl transfer reaction. During monomethy-
lation, they function to constrain the movement of the lysine
-amino group by mediating hydrogen bonds between the
substrate and enzyme, thereby promoting the linear align-
ment with the methyl group and sulfonium cation of
AdoMet (Fig. 4, A and B). During di- and trimethylation, the
water molecules either relocate within the lysine binding
channel or dissociate from the enzyme, yielding the space
required to rotate the methyl group away from the methyl
transfer pore and to realign the -amine in productive geom-
etry for the next methyl transfer reaction (Figs. 4C and 5, B
and C). These findings agree with our prior analysis of the
SET8 Phe/Tyr switch mutant in which we demonstrated that
the Y334F substitution attenuates hydrogen bonding to the
water molecule bound in the solvent pocket, promoting its
dissociation and the conformational changes necessary for
lysine dimethylation (8). Indeed, there is a nearly identical
alignment of the dimethyl-lysine side chains in the structures of
SET7/9Y305FandSET8Y334Fcomplexes,despitethedifferences
in the orientations of the Phe-305 and Phe-334 side chains in each
structure (Fig. 2F). Finally, both the SET8 Y334F (8) and SET7/9
Y305F mutants (Table 2) displayed diminished catalytic efficien-
cies for lysine dimethylation versus monomethylation. These dif-
ferences may reflect the kinetics of the reorganization within the
active site, including the dissociation of the water molecule from
the solvent pocket and the concomitant realignment of the
monomethyl-lysine into a productive geometry for dimethylation.
In addition to their place-holding role, the active site waters
may also facilitate the deprotonation of the lysine -amino
group between methyl transfer reactions. For methylation to
occur, the -amino group must be deprotonated to function as
the nucleophile in the SN2 methyl transfer reaction with
AdoMet (Figs. 4 and 5). Although the pKa value of the lysine
-amine in solution is 10.5, molecular dynamics simulations
by Zhang and Bruice (25, 26) indicate that this value diminishes
to 8.2 upon formation of the SET7/9 Michaelis complex due to
the proximity of the AdoMet sulfonium cation and the low
dielectric constant of the active site. Furthermore, their simu-
lations show that a chain of water molecules facilitates the dep-
rotonation of the -amino group prior to methyl transfer, trans-
ferring the proton to bulk solvent. Although these water
molecule chains are not evident in our crystal structures, the
Y305F and Y245A complexes suggest another potential mech-
anism for deprotonation. In the dimethyl-lysine complexes of
the Y305F and Y245A mutants, the dissociation of water 1 and
2, respectively, from the lysine binding channel requires that
the solvent-mediated hydrogen bond to the -amino group is
broken (Figs. 2D and 3C). It is conceivable that these waters
dissociate from the active site as hydronium ions, promoting
the realignment and deprotonation of the methyl -amino
group for the next methyl transfer reaction.
A comparison of the SET7/9 Y305F and SET8 Y334F com-
plexes yields insights into the mechanism by which the Phe/Tyr
switch influences water binding within the solvent pocket. The
phenylalanine substitution in the Phe/Tyr switch results in the
loss of a single hydrogen bond to the water molecule (water 1) in
the solvent pocket compared with the four hydrogen bonds that
coordinate the solvent molecule in WT SET7/9 (Fig. 2, A and B)
and SET8 (7, 8). Although this attenuation in hydrogen bonding
may appear insignificant, this difference is nonetheless impor-
tant for at least two reasons. First, theoretical calculations indi-
cate that, on average, water molecules form 3.5 hydrogen
bonds in solutions (40, 41). This value is greater than the num-
ber of hydrogen bonds coordinating water 1 in the solvent
pocket in SET7/9 Y305F (Fig. 2, B and C) as well as in SET8
Y334F and other di- and trimethyltransferases that possess a
hydrophobic residue in the Phe/Tyr switch site (8). From the
perspective of the water molecule, the greater hydrogen bond-
ing potential in solution would tend to thermodynamically
favor its dissociation from the solvent pocket in SET domain
KMTs that lack a tyrosine in the Phe/Tyr switch position. Sec-
ond, the ordered binding of water molecules observed in the
active sites of SET domain ternary complexes represents an
unfavorable entropy compared with their diffusion in bulk sol-
vent. In WT SET7/9 (Fig. 2A) and SET8 (7, 8), this entropic
penalty can be partially offset through the favorable enthalpy of
binding associated with the four hydrogen bonds that coordi-
nate the water within the solvent pocket. It is conceivable that
the loss of the tyrosine-mediated hydrogen bond in the Phe/Tyr
switch shifts the equilibrium in favor of dissociation of the
water molecule from the solvent pocket, thereby facilitating
dimethylation in SET7/9 Y305F, SET8 Y334F, and other di- and
trimethyltransferases.
The structures of the SET7/9 Y245A and Y305F complexes
illustrate the interactions that align the lysine -amino group
during the methyl transfer reactions in each enzyme. In the WT
enzyme and the Y305F mutant, hydrogen bonding to the
hydroxyl group of Tyr-245 appears to be critical in properly
aligning the -amine for methyl transfer (Fig. 4). Tyr-245 is
conserved in the sequences of many SET domain KMTs (8, 42),
and substitutions of this residue generally impair or abolish
activity, indicating its importance in catalysis (8, 43). However,
SET7/9 appears to be an exception to this rule, as the Y245A
mutant is not only active but is capable of catalyzing lysine
trimethylation. In this mutant, Tyr-305 appears to assume the
role of Tyr-245 by hydrogen bonding to the monomethylated
-amino group to align it for methyl transfer with AdoMet, as
illustrated in the modeled substrate complex for the dimethy-
lation reaction (Fig. 5B). Conversely, in the model for trimethy-
lation, the Tyr-305 hydroxyl group does not hydrogen bond to
the -amine but instead participates in a CH–O hydrogen bond
with one of the methyl groups to assist in aligning the dimethy-
lated -amine for the methyl transfer reaction (Fig. 5C). Addi-
tional structural and functional studies of the SET domain tri-
methyltransferases will aid in further illuminating the roles
of CH–O hydrogen bonds in facilitating lysine multiple
methylation.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31857
Acknowledgments—We acknowledge S. Schiebold for assistance in
protein expression, purification, and crystallization and S. Anderson
and R. Sanishvili for their assistance with x-ray data collection. We
also thank S. Bulfer and S. Horowitz for reading the manuscript and
providing useful comments. This work utilized the Protein Structure
Facility of the Michigan Diabetes Research and Training Center, Uni-
versity of Michigan, supported by National Institutes of Health Grant
DK020572, NIDDK. Use of the Advanced Photon Source was sup-
ported by the United States Department of Energy, Basic Energy Sci-
ences, Office of Science, under Contract DE-AC02-06CH11357.
GM/CA CAT has been funded in whole or in part by National
Institutes of Health NCI Grant Y1-CO-1020 and NIGMS Grant
Y1-GM-1104. Use of the LS-CAT Sector 21 was supported by Michi-
gan Economic Development Corporation and the Michigan Technol-
ogy Tri-Corridor Grant 085P1000817 for the support of this research
program.
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Lysine Methylation by SET7/9 Mutants
31858
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VOLUME 285•NUMBER 41•OCTOBER 8, 2010
|
3M57
|
SET7/9 Y245A in complex with TAF10 peptide and AdoHcy
|
SET7/9 Catalytic Mutants Reveal the Role of Active Site Water
Molecules in Lysine Multiple Methylation*□
S
Received for publication,February 15, 2010, and in revised form, July 14, 2010 Published, JBC Papers in Press,August 1, 2010, DOI 10.1074/jbc.M110.114587
Paul A. Del Rizzo‡, Jean-Franc¸ois Couture‡§1, Lynnette M. A. Dirk¶, Bethany S. Strunk‡, Marijo S. Roiko‡,
Joseph S. Brunzelle, Robert L. Houtz¶, and Raymond C. Trievel‡2
From the ‡Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109, the §Ottawa Institute of
Systems Biology, Department of Biochemistry, Microbiology, and Immunology, University of Ottawa, Ottawa, Ontario K1H 8M5,
Canada, the ¶Department of Horticulture, Plant Physiology/Biochemistry/Molecular Biology Program, University of Kentucky,
Lexington, Kentucky 40546, and the Department of Molecular Pharmacology and Biological Chemistry, Northwestern University
Center for Synchrotron Research, Life Sciences Collaborative Access Team, Argonne, Illinois 60439
SET domain lysine methyltransferases (KMTs) methylate
specific lysine residues in histone and non-histone substrates.
These enzymes also display product specificity by catalyzing dis-
tinct degrees of methylation of the lysine -amino group. To
elucidate the molecular mechanism underlying this specificity,
we have characterized the Y245A and Y305F mutants of the
human KMT SET7/9 (also known as KMT7) that alter its prod-
uct specificity from a monomethyltransferase to a di- and a tri-
methyltransferase, respectively. Crystal structures of these
mutants in complex with peptides bearing unmodified, mono-,
di-, and trimethylated lysines illustrate the roles of active site
water molecules in aligning the lysine -amino group for methyl
transfer with S-adenosylmethionine. Displacement or dissocia-
tion of these solvent molecules enlarges the diameter of the
active site, accommodating the increasing size of the methylated
-amino group during successive methyl transfer reactions.
Together, these results furnish new insights into the roles of
active site water molecules in modulating lysine multiple meth-
ylation by SET domain KMTs and provide the first molecular
snapshots of the mono-, di-, and trimethyl transfer reactions
catalyzed by these enzymes.
SET domain enzymes represent a family of S-adenosylmethi-
onine (AdoMet)3-dependent methyltransferases that catalyze
the site-specific methylation of protein lysyl residues in a host
of proteins, including histones, transcription factors, chroma-
tin-modifying enzymes, ribosomal subunits, and other sub-
strates (1–3). In many instances, these modifications serve to
recruit effector proteins that recognize methyl-lysyl residues in
a sequence-dependent fashion (4). In addition, SET domain
KMTs exhibit product specificity, defined as their ability to cat-
alyze mono-, di-, or trimethylation of the lysine -amino group.
This specificity is biologically relevant because many methyl-
lysine-binding proteins can discriminate among different
degrees of lysine methylation (4). Thus, both the site and degree
of lysine methylation are critical to recognition by effector
proteins.
Structural and functional studies have identified a Phe/Tyr
switch in the active site of SET domain KMTs that governs their
respective product specificities (5, 6). According to this model,
KMTs that possess a tyrosine in the Phe/Tyr switch site are
limited to catalyzing lysine monomethylation, whereas en-
zymes that possess a phenylalanine or another hydrophobic
residue in this position display di- or trimethyltransferase activ-
ity. Mutational analysis of various SET domain KMTs, includ-
ing DIM-5 (KMT1) (6), SET7/9 (6), G9A (KMT1C) (5), and
SET8 (also known as PR-SET7 and KMT5A) (7, 8), has demon-
strated that substitutions in the Phe/Tyr switch result in pre-
dictable changes in product specificity. Several models have
been proposed to explain the mechanism by which the Phe/Tyr
switch site governs this specificity, including variations in the
diameter of the active site due to the size of Phe/Tyr switch
residue and steric hindrance by the tyrosine hydroxyl group (6,
9–11). However, our recent studies of the Phe/Tyr switch
mutant Y334F in the human histone H4 Lys-20 (H4K20) meth-
yltransferase SET8 indicate that the Phe/Tyr switch regulates
product specificity via a more subtle mechanism (8). Specifi-
cally, the switch modulates the binding of an active site water
molecule that in turn regulates the transition from mono-
methylation to multiple methylation.
Among the KMTs that have been structurally characterized,
SET7/9 has emerged as an archetypal model for studying the
catalytic mechanism and product specificity of the SET domain
family. Although initially isolated as a histone H3 Lys-4
(H3K4)-specific methyltransferase, this KMT has been shown
to regulate the functions of numerous non-histone substrates
through site-specific methylation (12–21). Early structural and
functional studies of SET7/9 identified two active site mutants,
* Thisworkwassupported,inwholeorinpart,byNationalInstitutesofHealth
Grant R01 GM073839 (to R. C. T.) and National Institutes of Health Admin-
istrative Supplement GM073839-04S1 (to R. C. T.) funded through the
American Recovery and Reinvestment Act. This work was also supported
by Department of Energy Grant DE-FG02-92ER20075 (to R. L. H.).
□
S The on-line version of this article (available at http://www.jbc.org) contains
supplemental Table 1.
The atomic coordinates and structure factors (codes 3M53, 3M54, 3M55, 3M56,
3M57,3M58,3M59,and3M5A)havebeendepositedintheProteinDataBank,
Research Collaboratory for Structural Bioinformatics, Rutgers University, New
Brunswick, NJ (http://www.rcsb.org/).
1 Supported by a Canadian Institutes of Health Research postdoctoral
fellowship.
2 To whom correspondence should be addressed: Dept. of Biological Chem-
istry, University of Michigan Medical School, 1150 West Medical Center Dr.,
5301 MSRB III, Ann Arbor, MI 48109. Tel.: 734-647-0889; Fax: 734-763-4581;
E-mail: rtrievel@umich.edu.
3 The abbreviations used are: AdoMet, S-adenosylmethionine; AdoHcy, S-ad-
enosylhomocysteine; KMT, lysine methyltransferase; ITC, isothermal titra-
tion calorimetry; CH–O, carbon-oxygen hydrogen bond; BisTris, 2-[bis(2-
hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 41, pp. 31849–31858, October 8, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31849
Y245A and Y305F, which change its product specificity. The
Phe/Tyr switch mutant Y305F alters SET7/9 product specificity
from a mono- to dimethyltransferase (6), whereas the Y245A
substitution converts the enzyme into a trimethyltransferase
with weak monomethyltransferase activity (11). These mutants
have been the subjects of numerous molecular modeling simu-
lations that have led to various models to explain their distinct
product specificities (22–26). However, the lack of structural
data for the SET7/9 Y245A and Y305F mutants in complex with
cognate methylated peptides has hindered our understanding
of the mechanisms that define the respective product specific-
ities of these mutants. Moreover, these structures would yield a
framework for visualizing the mono-, di-, and trimethylation
reactions catalyzed by SET domain KMTs.
To gain insight into the molecular basis of their product
specificities, we have determined high resolution crystal
structures of the SET7/9 Y245A and Y305F mutants in com-
plex with peptides of the TATA box-binding protein-associ-
ated factor TAF10 bearing the Lys-189 methylation site in
unmodified (K189), monomethylated (K189me1), dimethyl-
ated (K189me2), and trimethylated (K189me3) states. The
structures and accompanying biochemical data support a
model whereby changes in the occupancy or position of water
molecules in the active site are critical in establishing the prod-
uct specificities of the SET7/9 Y245A and Y305F mutants.
Together, our results provide new insights into the mechanisms
that govern SET domain product specificity and provide step-
wise snapshots of the lysine mono-, di-, and trimethyl transfer
reactions catalyzed by KMTs.
EXPERIMENTAL PROCEDURES
Cloning, Expression, and Purification of the SET7/9 Mutants—
The Y245A and Y305F mutants were introduced into the pHIS2
SET7/9 expression vector encoding residues 110–366 (27) via
QuikChange site-directed mutagenesis (Stratagene) and were
verified by dideoxy DNA sequencing. The plasmids encoding
wild type (WT) SET7/9 and the Y245A and Y305F mutants
were transformed into Rosetta2 DE3 cells (Novagen) and were
expressed as described previously (27, 28). In the course of
characterizing WT SET7/9, we observed that the enzyme co-
purified with AdoMet or another contaminant that resulted in
technical difficulties in the isothermal titration calorimetry
(ITC) experiments and co-crystallization trials with the TAF10
peptides. To overcome this problem, a denaturation and refold-
ing step was inserted in the purification scheme. The denatur-
ation and refolding protocol involved adding 6 M guanidine
HCl, 50 mM sodium phosphate, pH 7.5, and 500 mM NaCl to the
protein while it was immobilized on a nickel-Sepharose column
(GE Healthcare). The column was washed with this buffer, fol-
lowed by a solution of 6 M urea, 50 mM sodium phosphate, pH
7.5, 150 mM NaCl, and 10 mM 2-mercaptoethanol to remove the
cofactor from the denatured enzyme. A reverse gradient from 6
to 0 M urea was then performed in the same buffer to refold the
protein, which was subsequently eluted from the column using
a linear gradient of 0–500 mM imidazole in 50 mM sodium
phosphate, pH 7.5, 500 NaCl, and 10 mM 2-mercaptoethanol.
The refolded protein was digested with tobacco etch virus pro-
tease (29) during dialysis against 20 mM Tris, pH 7.5, 150 mM
NaCl, and 5 mM 2-mercaptoethanol and then purified using a
Superdex 200 gel filtration column (GE Healthcare). Protein
concentration was determined by its absorbance at 280 nm.
Synthetic Peptides—The TAF10 peptides bearing K189,
K189me1,
K189me2,
and
K189me3
(sequence,
acetyl-
SKSK189DRKYTL-amide) and a biotinylated TAF10 peptide
(sequence,
acetyl-SKSK189DRKYTLT(K-EZLink-S-S-biotin)-
amide) were synthesized and purified by New England Peptide,
Inc. Peptide concentrations were measured using the absorb-
ance of their tyrosine residue at 274 nm.
Crystallization and Data Collection—Crystals were pro-
duced by hanging drop vapor diffusion by mixing the crystalli-
zation solution in a 1:1 ratio with 10–20 mg/ml SET7/9, 1 mM
S-adenosylhomocysteine (AdoHcy), and 1.0–3.0 mM unmodi-
fied or methylated TAF10-K189 peptide in 20 mM Tris, pH 8.0,
100 mM NaCl, and 2 mM tris(2-carboxyethyl)phosphine. Crys-
tals were obtained at 20 °C in either 1.8–2.0 M (NH4)2SO4, with
0.1 M BisTris, pH 6.2–6.6, or in 0.95–1.1 M sodium citrate with
100 mM imidazole pH 8.0–8.4. In both crystallization condi-
tions, the final pH values were between pH 8.0 and 9.0. Crystals
in the (NH4)2SO4 condition were typically flash-frozen in the
mother liquor containing 25–30% glycerol, and the crystals in
the citrate condition were frozen in 1.6 M sodium citrate. Data
were collected at the Advanced Photon Source beamlines
21-IDG (LS-CAT) and 23-IDD (GM/CA-CAT). Images were
indexed, integrated, and scaled using HKL2000 (30). Structures
of the mutants were solved by molecular replacement using
MOLREP (31) with the coordinates of a previously reported
SET7/9 ternary complex used as the search model (Protein
Data Bank code 2F69). Successive rounds of model building and
refinement were carried out using Coot (32) and REFMAC (33),
respectively. The geometry of the models were verified by Mol-
Probity (34). Simulated annealing omit maps were calculated
using CNS (35) with the peptide and cofactor removed to elim-
inate model bias in the active site. Structural figures were ren-
dered using PyMOL (Schro¨dinger, LLC.).
Fluorescent Methyltransferase Assay—A coupled fluorescent
methyltransferase assay was used to measure the kinetic
parameters of WT SET7/9 and the Y245A and Y305F mutants
as reported previously, with the exception that 50–150 nM
enzyme, 100 M AdoMet, and varying concentrations of TAF10
peptide substrate were used (27, 36). Assays were performed in
triplicate, and a homocysteine calibration curve was used to
calculate the initial velocities. Kinetic parameters were calcu-
lated by plotting the velocities versus peptide concentration and
by fitting the Michaelis-Menten equation to the data via non-
linear regression using Prism 5.0 (GraphPad). In cases where
the Km value was beyond the measurable range of the assay, the
kcat/Km value was determined as described previously (7).
Calorimetry Experiments—ITC was performed at 20 °C using
a MicroCal VP-ITC calorimeter (GE Healthcare) with 0.12 mM
protein and 4 mM AdoHcy in 20 mM sodium phosphate, pH 7,
and 100 mM NaCl with 1.5 mM peptide as the injectant. Data
were processed, and equilibrium dissociation constants (KD)
and curve fitting errors were calculated from the binding iso-
therms using Origin 7.0 (OriginLab Corp.). WT SET7/9 and the
Y245A and Y305F mutants displayed ligand:protein binding
stoichiometries (N values) between 0.8 and 1.0, demonstrating
Lysine Methylation by SET7/9 Mutants
31850
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
that WT SET7/9 and its mutants were properly refolded due to
their ability to bind peptides in an 1:1 molar ratio.
TLC Product Analysis—Methyltransferase assays were per-
formed in triplicate at 37 °C with the biotinylated TAF10 pep-
tide (0.5 mM) for 30 min in the presence of WT SET7/9 (3
pmol), Y305F mutant (6 pmol), or the Y245A mutant (100
pmol). Assays contained 100 mM HEPES, pH 8.0, 50 mM NaCl,
1.0 mM [methyl-3H]AdoMet (diluted to a specific activity of 0.2
Ci/nmol with purified AdoMet (37)), 15 M Sulfolobus solfa-
taricus AdoHcy hydrolase (36), and 2 units of adenosine deami-
nase (Roche Applied Science) in a final volume of 20 l. The
reactions were terminated by addition of an equal volume of
200 mM MES, pH 5.1, prior to addition of a 2-fold molar excess
of immobilized avidin resin (UltraLink; Pierce). Biotinylated
peptides were allowed to bind at room temperature for 30 min,
and the resin was then collected by centrifugation (9000 g).
The resin was washed three times with 300 mM NaCl, and the
peptide was eluted overnight from the avidin resin by cleavage
of the disulfide bond in the linker of the peptide using 10 mM
tris(2-carboxyethyl)phosphine (Thermo Scientific). The resin
was incubated with additional 10 mM tris(2-carboxyethyl)phos-
phine the following day until the radiolabel was essentially
removed from the resin. The recovered peptides were hydro-
lyzed by incubation with 6 M HCl at 110 °C for 24 h. Subsequent
steps in measuring the radiolabel incorporated into the mono-,
di-, and trimethyl-lysine products were performed as reported
previously (8).
RESULTS
Functional Analysis of the SET7/9
Y305F Mutant—Prior studies of
SET7/9 by Zhang et al. (6) reported
that mutation of the Phe/Tyr switch
residue Tyr-305 to a phenylalanine
alters its product specificity from a
mono- to dimethyltransferase. We
verified these findings by demon-
strating that WT SET7/9 mono-
methylated the TAF10-K189 pep-
tide, whereas the Y305F mutant
mono- and dimethylated this sub-
strate, as demonstrated by mass
spectrometry (data not shown). We
next examined whether the Y305F
substitution altered the affinity of SET7/9 for the TAF10-K189
peptides using ITC (Fig. 1). A comparison of the KD values
revealed that SET7/9 Y305F bound the TAF10-K189 and
TAF10-K189me1 peptides 4- and 6-fold more tightly, respec-
tively, than the WT enzyme, whereas this mutant displayed a
substantially diminished affinity for the TAF10-K189me2 pep-
tide (Table 1). Although the WT enzyme and the Y305F mutant
exhibited discernable differences in their affinities for the
unmodified and monomethylated peptides, these variations are
modest and cannot account for their distinct product specific-
ities, suggesting that a kinetic effect during methylation may be
responsible.
To investigate this possibility, we characterized the kinetic
parameters of WT SET7/9 and the Y305F mutant using the
TAF10 peptides as substrates. Both enzymes methylated the
unmodified peptide with comparable kcat and Km values (Table
2). In analyzing the kinetic parameters for the methylation
of the monomethylated peptide by SET7/9 Y305F, we found
that this substrate displayed an elevated Km value that was
beyond the measurable range of the assay due to its limited
solubility. In this case, we measured the catalytic efficiency
(kcat/Km) for the methylation of this peptide and found that it
was methylated 15-fold less efficiently than the unmodified
peptide by SET7/9 Y305F. Given the fact that the Y305F mutant
exhibited a higher binding affinity for the TAF10-K189me1
peptide than the WT enzyme (Table 1), the kinetic data suggest
that a step in the reaction pathway following substrate binding
limits the catalytic efficiency of this mutant.
We next examined whether the Y305F mutant dimethylated
the TAF10-K189 peptide via a processive or a distributive
mechanism. In a processive mechanism, the methyl-lysine
substrate would remain bound to the enzyme during successive
methyl transfer reactions; thus, the concentration of an inter-
mediate, such as monomethyl-lysine, cannot exceed the en-
zyme concentration during the assay. In a distributive mecha-
nism, the intermediates are released into solution where they
accumulate prior to the next round of methylation, resulting in
an intermediate concentration that is greater than that of the
enzyme. Using a radiometric TLC assay and a biotinylated
TAF10 peptide, we quantified the amounts of monomethylated
products generated by the WT SET7/9 and the Y305F mutant
FIGURE 1. ITC analysis of WT SET7/9 and the Y245A and Y305F mutants. Representative titrations and
binding isotherms for the calorimetry experiments are shown for the unmodified TAF10-K189 peptide titrated
into WT SET7/9 (left plot), SET7/9 Y305F (middle plot), and SET7/9 Y245A (right plot). In each panel, the ITC
titration experiment (upper panel) is shown with the binding isotherms (lower panel) fit to a single binding site
model.
TABLE 1
Analysis of the binding affinity of WT SET7/9 and its catalytic
mutants for unmodified and methylated TAF10 peptides
SET7/9
TAF10 peptide
KD
a
M
WT
K189
4.9 0.20
WT
K189me1
4.0 0.36
Y305F
K189
1.3 0.10
Y305F
K189me1
0.62 0.065
Y305Fb
K189me2
70
Y245A
K189
4.0 0.25
Y245A
K189me1
3.3 0.10
Y245A
K189me2
5.8 0.22
Y245A
K189me3
11 0.28
a Curve fitting errors were calculated from the binding isotherms.
b An estimate of the affinity is reported due to weak peptide binding.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31851
(Table 3). The data demonstrate that comparable amounts of
monomethyl-lysine were generated when the quantity of
enzyme usedistakenintoaccount,inagreementwiththeirsimilar
turnover numbers for the TAF10-K189 peptide (Table 2). The
Y305F mutant also produced small but measurable quantities of
radiolabeled dimethyl-lysine product that were substantially
smaller than the amount of monomethyl-lysine generated.
Therefore, the TLC data are consistent with a distributive
mechanism for dimethylation by the Y305F mutant because the
amount of monomethyl-lysine produced exceeded the quantity
of enzyme used in the assay.
Structures of WT SET7/9 and the Y305F Mutant in Complex
with Unmodified and Methylated TAF10 Peptides—To deter-
mine the mechanism by which the Y305F substitution alters the
product specificity of SET7/9, we determined the crystal struc-
tures of this mutant bound to AdoHcy and TAF10-K189,
TAF10-K189me1, and TAF10-K189me2 peptides and com-
pared these to the structures of the WT SET7/9AdoHcy
TAF10-K189 complex (supplemental Table 1). The structures
of these complexes were determined to 1.85 Å or higher reso-
lution, permitting unambiguous modeling of the K189 side
chains in the active site of the enzyme based on simulated
annealing omit maps (Fig. 2). The ternary complexes of the WT
and the Y305F mutant superimpose with overall root mean
square differences of less than 0.3 Å for all aligned atoms, indi-
cating that neither the Y305F mutation nor the binding of the
various TAF10-K189 peptides results in substantial changes in
its overall structure.
An inspection of the active sites of the SET7/9 WT and
Y305F complexes illustrates the binding modes of the unmod-
ified and methylated forms of K189 in the TAF10 peptides (Fig.
2, A–D). The K189 side chain binds in an extended all trans
conformation in a deep pocket, termed the lysine binding chan-
nel, that is composed of Tyr-245, Thr-266, Leu-267, Ser-268,
Tyr-305, Tyr-335, and Tyr-337 (not shown for clarity) (Fig. 2A).
These residues interact with the aliphatic portion of the K189
side chain primarily through van der Waals contacts. The lysine
binding channel connects to the AdoMet-binding site on the
opposite face of the catalytic domain via an oxygen-lined
methyl transfer pore (38). During catalysis, the methyl group of
the cofactor is positioned within the methyl transfer pore
for the SN2 reaction with the -amino group of the lysine or
methyl-lysine substrate (see below).
To lower the activation barrier for this reaction, the lysine
-amine nucleophile is aligned for methyl transfer through a
hydrogen bond network within the active site. In the WT
enzyme, the K189 -amino group hydrogen bonds to the
hydroxyl group of Tyr-245 as well as to two water molecules
(Fig. 2A). One of the water molecules (termed water 1), is coor-
dinated in a solvent pocket, through hydrogen bonds to the
carbonyl oxygens of Gly-292 and Ala-295 and to the hydroxyl
group of the Phe/Tyr switch residue Tyr-305. This solvent
pocket is structurally conserved in SET domain KMTs and has
an important role in defining product specificity through the
adjacent Phe/Tyr switch residue, as shown in our prior studies
of the human H4K20 methyltransferase SET8 (8). The other
water molecule is bound within the methyl transfer pore
between the lysine substrate and the thioether sulfur atom of
AdoHcy through hydrogen bonds to Tyr-245, Asn-265, and
His-293 in SET7/9 and the TAF10-K189 -amino group. This
water is not observed in other structures of SET7/9 ternary
complexes and may represent the approximate position that
the AdoMet methyl group occupies in the methyl transfer pore
in the Michaelis complex.
In structures of the Y305F ternary complexes, the K189,
K189me1, and K189me2 side chains also adopt extended trans
side chain geometries within the lysine binding channel that are
stabilized via hydrogen bonding to Tyr-245 in the enzyme (Fig.
2, B–D). The orientations of the K189me1 and K189me2 side
chains are further maintained through carbon-oxygen (CH–O)
hydrogen bonding between the methyl groups and oxygen
atoms within the vicinity of the methyl transfer pore, as
reported previously in other SET domain KMT structures (8,
10, 38). A superimposition of the SET7/9 WT and Y305F com-
plexes underscores the similarity of the lysyl binding conforma-
tions (Fig. 2E). However, there are notable differences in the
hydrogen bond patterns and occupancy of water 1 within the
solvent pocket in the Y305F mutant compared with the WT
enzyme. Specifically, the Y305F substitution results in the loss
of one hydrogen bond to water 1 in the structures of the TAF10-
K189 and TAF10-K189me1 complexes (Fig. 2, B and C). In con-
trast, water 1 is absent in TAF10-K189me2 complex, and the
vacated solvent pocket is occupied by one of the methyl groups
TABLE 2
Kinetic parameters of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
TAF10 peptide substrate
Km
a
kcat
a
kcat/Km
a
M
min1
M1 min1 103
WT
K189
160 17
17 0.62
110 17
Y305F
K189
88 5.0
17 0.30
190 11
Y305Fb
K189me1
11 0.50
Y245A
K189
200 35
0.53 0.04
2.6 0.47
Y245A
K189me1
210 23
5.9 0.23
28 3.3
Y245A
K189me2
400 29
6.5 0.16
15 1.2
a Curve fitting errors were calculated from the hyperbolic fits for the Michaelis-Menten equation.
b Km and kcat were not determined because the Km value was beyond the measurable range; therefore, the kcat/Km value is reported.
TABLE 3
Product analysis of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
Quantity
of enzyme
Measured
product
Amount of
product formeda
nmol
nmol
WT
0.003
Kme1
0.65 0.07
Y305F
0.006
Kme1
1.5 0.49
Kme2
0.033 0.009
Y245A
0.100
Kme1
0.80 0.22
Kme2
0.39 0.021
Kme3
0.076 0.019
a Standard deviation was calculated from triplicate measurements.
Lysine Methylation by SET7/9 Mutants
31852
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
of the dimethyl -amine (Fig. 2D). This methyl group forms a
3.5-Å CH–O hydrogen bond to the carbonyl oxygen of Ala-295,
further stabilizing the binding of the dimethyl-lysine side chain.
A homologous dimethyl-lysine-binding mode and CH–O
hydrogen bond was observed in our prior structural studies of
the SET8 Y334F Phe/Tyr switch mutant that confers an analo-
gous change in product specificity from a mono- to a dimeth-
yltransferase (8). A structural alignment of the active sites of the
SET7/9 Y305F and SET8 Y334F mutants bound to cognate
dimethylated peptides illustrates that the coordinates of the
dimethyl-lysyl side chains are virtually superimposable, with
one methyl group oriented toward the methyltransfer pore and
the second positioned within the vacant solvent pocket (Fig.
2F). Taken together, the structures of the SET7/9 Y305F com-
plexes and the similarities in the dimethyl-lysine conforma-
tions in the SET7/9 Y305F and SET8 Y334F mutants imply that
the Phe/Tyr switch governs product specificity through a con-
served mechanism whereby it indirectly influences the binding
modes of the methyl-lysine side chain by modulating the affin-
ity of the water molecule (water 1) bound in the solvent pocket.
Biochemical Characterization of
the SET7/9 Y245A Mutant—Previ-
ous studies by Xiao et al. (11)
reported that the Y245A mutation
yields an unusual change in the
product
specificity
of
SET7/9,
converting the enzyme to a trimeth-
yltransferase with weak monometh-
yltransferase activity. We deter-
mined that the SET7/9 Y245A could
mono-, di-, and trimethylate the
TAF10-K189 peptide by mass spec-
trometry (data not shown) and TLC
(Table 3), confirming the earlier
studies of Xiao et al. (11). ITC
analysis revealed that the Y245A
mutant displayed comparable KD
values
for
the
unmodified
and
methylated TAF10-K189 peptides
(Fig. 1), although its affinity for the
trimethylated peptide was modestly
diminished in comparison with the
other peptides (Table 1). The ITC
data demonstrate that the Y245A
mutant
bound
the
unmodified,
mono-,
and
dimethylated
sub-
strates with equivalent affinities,
suggesting that a kinetic effect or
a structural alteration in the active
site may be responsible for its
diminished
activity
toward
un-
modified substrates.
To gain further insight into its
peculiar
product
specificity,
we
characterized the kinetic properties
of
the
SET7/9
Y245A
mutant.
Steady state analysis demonstrated
that this mutant displayed similar
Km values for the unmodified, mono- and dimethylated TAF10
peptides (Table 2). However, the turnover number for the
TAF10-K189 peptide was diminished over 10-fold versus the
methylated peptides and was reduced 30-fold versus the WT
enzyme, in agreement with the weak monomethyltransferase
activity reported by Xiao et al. (11). In addition, we investigated
whether this mutant catalyzes lysine trimethylation via a pro-
cessive or distributive mechanism as described for SET7/9
Y305F. The TLC data illustrate that the mono- and dimethyl-
lysine intermediates accumulated at quantities greater than
that of the enzyme used in the assay, indicating that SET7/9
Y245A obeys a distributive mechanism, analogous to the Y305F
mutant (Table 3).
Structures of SET7/9 Y245A Bound to Unmodified and Meth-
ylated TAF10 Peptides—To elucidate the mechanism underly-
ing its unusual product specificity, we determined the crystal
structures of SET7/9 Y245A in complex with AdoHcy and
unmodified, mono-, di-, and trimethylated TAF10 peptides
(supplemental Table 1). These complexes superimpose with
the structure of the WT SET7/9AdoHcyTAF10-K189 com-
FIGURE 2. Crystal structures of WT SET7/9 and the Y305F mutant in complex with AdoHcy and unmodi-
fied and methylated TAF10 peptides. A, active site of WT SET7/9 bound to AdoHcy and TAF10-K189 and the
active site of SET7/9 Y305F bound to AdoHcy and TAF10-K189 (B), TAF10-K189me1 (C), and TAF10-K189me2
peptides (D). A–D, the K189 residue in the TAF10 peptide is depicted with yellow carbon atoms with the
corresponding Fo – Fc simulated-annealing omit maps (contoured at 2.5 ) shown. AdoHcy is colored with
green carbon atoms, and SET7/9 carbon atoms are colored gray, with the exception of Tyr-305/Y305F
(magenta) and Tyr-245 (cyan). Gly-264 to Ser-268 are shown, and the main chain atoms of Gly-292 to Ala-295
are depicted for clarity. Red dashed lines indicate conventional hydrogen bonds, and blue dashed lines indicate
CH–O hydrogen bonds. Cyan dashed lines indicate weak CH–O hydrogen bonds between 3.5 and 3.7 Å in
length. The water molecule in the solvent binding pocket is numbered 1. E, overlay of the active sites of the WT
enzyme (light blue carbon atoms) and the Y305F mutant (green, yellow, and pink carbon atoms correspond to
K189, K189me1, and K189me2, respectively). Red dashed lines indicate coordination of the waters in the struc-
turesoftheY305Fmutant,andthelightbluedashedlineindicateshydrogenbondingthatonlyoccursintheWT
enzyme. F, superimposition of the active sites of the SET7/9 Y305FAdoHcyTAF10-K189me2 and SET8
Y334FAdoHcyH4K20me2 (Protein Data Bank code 3F9X) complexes shown with green and yellow carbon
atoms, respectively.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31853
plex with root mean squared differences of less than 0.4 Å for all
aligned atoms, indicating that the Y245A mutant does not per-
turb the overall structure of the enzyme. Simulated annealing
omit maps illustrate that K189 side chains are bound within the
lysine binding channel through hydrogen bonds and van der
Waals contacts (Fig. 3, A–D), although the interactions and
binding modes are distinct from those in the complexes of WT
SET7/9 and the Y305F mutant (Fig. 2, A–D). In the unmodified
TAF10 peptide complex, the K189 -amino group forms a weak
hydrogen bond with the carbonyl oxygen of Gly-264 (Fig. 3A),
whereas the -amino groups of K189me1 and K189me2 hydro-
gen bond to the hydroxyl group of Tyr-305 in the mono- and
dimethylated peptide complexes (Fig. 3, B and C). The confor-
mations of the K189me1 and K189me2 side chains are further
stabilized by water-mediated hydrogen bonding and through
CH–O hydrogen bonding to their methyl groups. In the
TAF10-K189me3 peptide complex, the trimethyl-lysine side
chain is coordinated exclusively through direct and water-me-
diated CH–O hydrogen bonds to its methyl groups because the
quaternary -ammonium cation cannot engage in hydrogen
bonding (Fig. 3D).
A structural alignment of the four SET7/9 Y245A complexes
illustrates distinct binding modes for the unmodified versus the
methylated K189 side chains, highlighting the selectivity of this
mutant for methylated substrates. The side chains of K189me1,
K189me2, and K189me3 roughly overlay with their respective
-amino groups superimposed and adopt slightly kinked con-
formations (Fig. 3E), as opposed to the extended trans geome-
try of the unmodified and methylated lysines in the complexes
of the WT enzyme and the Y305F mutant (Fig. 2, A–D). Con-
versely, the unmodified K189 side chain does not superimpose
with its methylated counterparts and is oriented in an alterna-
tive configuration due to its hydrogen bonding to Gly-264 (Fig.
3, A and E). An overlay of the structures of the WT enzyme and
Y245A mutant bound to the unmodified TAF10 peptide illus-
trates that the side chains of K189 do not superimpose and that
the K189 -amino group appears to be misaligned with AdoHcy
in the Y245A complex (Fig. 3F). This suboptimal alignment
may explain the diminished kcat value of SET7/9 Y245A mutant
toward substrates with unmodified lysines (Table 2).
A comparison of the structures of the SET7/9 Y245A and
Y305F complexes yields a molecular explanation for the differ-
ent product specificities of these two mutants. In the SET7/9
Y305F complexes, Tyr-245 aligns the K189 -amino group for
methyl transfer through hydrogen bonding to its hydroxyl
group (Fig. 2, B–D). Conversely, in the Y245A mutant, the
K189me1 and K189me2 -amino groups are oriented through
hydrogen bonding to Tyr-305 (Fig. 3, B and C). These distinct
hydrogen bond patterns impart differences in the conforma-
tions of the lysyl side chains due to the relative orientations of
Tyr-245 and Tyr-305 in the lysine binding channel. Specifically,
the kinked conformation adopted by the K189me1 and
K189me2 side chains in the Y245A complexes (Fig. 3, B and C)
may contribute to the differences in the turnover numbers of
this mutant versus those of the WT enzyme and the Y305F
mutant (Table 2). In addition, the dimethyl -amino group of
the K189me2 side chain binds in distinct orientations in the
Y245A and Y305F mutants due to their hydrogen bonding to
FIGURE 3. Crystal structures of the SET7/9 Y245A mutant in complex with
AdoHcy and unmodified and methylated TAF10 peptides. Active site of
SET7/9 Y245A bound to AdoHcy and TAF10-K189 (A), TAF10-K189me1 (B),
TAF10-K189me2 (C), and TAF10-K189me3 peptides (D). A–D, Fo – Fc simulated-
annealing omit maps (contoured at 2.5 ) for the unmodified and methylated
K189 side chains are illustrated. The residues and hydrogen bonds in each com-
plexarecoloredasinFig.2.Thewatermoleculesinthelysinebindingchannelsof
the Y245A complexes are numbered 1–4, as described in the text. E, superimpo-
sition of the active sites of the Y245A complexes bound to the four methylated
statesofTAF10-K189(unmodified,mono-,di-,andtrimethylatedshowningreen,
yellow, pink, and blue, respectively). F, overlay of WT SET7/9 (blue carbons) and
SET7/9 Y245A (green carbons) in complex with TAF10-K189. Waters correspond-
ing to the WT and Y245A structures are colored cyan and green, respectively.
G, alignment of the active sites of the Y245A (yellow carbons) and Y305F (green
carbons) mutants in complex with the TAF10-K189me2 peptide. Hydrogen
bonds from the Y305F structure are shown as green dashed lines, and waters and
hydrogen bonds in the Y245A structure are shown in yellow and orange,
respectively.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
Tyr-305 and Tyr-245, respectively (Fig. 3G). In the Y305F
mutant, hydrogen bonds to the dimethyl -amino group cou-
pled with steric constraints in the lysine binding channel pre-
vent the K189me2 side chain from undergoing a conforma-
tional change that is conducive to trimethylation (Fig. 2D),
consistent with its dimethyltransferase activity. However, in the
Y245A mutant, the alanine substitution enlarges the diameter
of the lysine binding channel, accommodating trimethyl-lysine
(Fig. 3D). In addition, the larger diameter would permit the
dimethyl-lysine substrate to undergo the conformational reor-
ganization necessary to align the -amino group in a productive
geometry for trimethylation.
A major difference in the active site of the Y245A mutant
versus the other SET7/9 structures is the presence of several
water molecules bound in the cavity generated by the Y245A
mutation. In the structure of the Y245A mutant bound to
TAF10-K189, three water molecules (waters 2–4) occupy this
cavity and are arranged in a triangular geometry (Fig. 3A). In
addition, water 1 shifts 1.6 Å from its position in the solvent
pocket toward water 2 to which it forms a hydrogen bond (Fig.
3, A and E). The shift in water 1 was unexpected given its con-
served orientation in the solvent pocket of the SET7/9 WT and
Y305F complexes (Fig. 2, A–C) as well as in the structures of
other SET domain KMTs (8). This displacement is presumably
related to the alternative conformation of the K189 side chain
whose -amino group is too distant (4.3 Å) to form a productive
hydrogen bond to water 1. Conversely, in the Y245A complexes
bound to TAF10-K189me1 and TAF10-K189me2, water 1
remains tightly bound in the solvent pocket through hydrogen
bonds to Tyr-305 hydroxyl group, the carbonyl oxygens of Gly-
292 and Ala-295, and the K189 -amino group (Fig. 3, B, C, and
E), analogous to its binding in the WT enzyme (Fig. 2A). How-
ever, in the TAF10-K189me3 complex, one of the methyl
groups of the trimethyl -ammonium cation is oriented into the
solvent pocket (Fig. 3D), similar to the dimethyl-lysine binding
mode observed in the Y305F mutant (Fig. 2D). The binding of
the methyl group in the solvent pocket displaces water 1 by 3.2
Å relative to its position in the TAF10-K189me1 complex (Fig.
3E), thereby avoiding a steric clash with the trimethylated
-ammonium group. Variations in the occupancy of water 2 are
also seen in the different Y245A structures. Water 2 is bound in
similar orientations in the active site of the unmodified and
monomethylated peptide complexes but is absent in the di- and
trimethylated peptide complexes due to the binding of a methyl
group in this position (Fig. 3, A–E). In summary, the changes in
the positions or occupancies of waters 1 and 2 correlate with the
binding modes of the unmodified and methylated K189 within
the active site of the Y245A mutant.
Catalytic Models of Lysine Multiple Methylation by SET7/9
Y245A, and Y305F—The structures of the SET7/9 complexes
reported here offer a prime opportunity to generate stepwise
models for lysine mono-, di-, and trimethylation by a SET
domain KMT. We modeled the AdoMet-bound Michaelis
complexes by superimposing the SET7/9 product complexes
with the previously reported structure of the SET7/9-AdoMet
binary complex (Fig. 4) (39). The conformations of the mono-
and dimethyl -amino groups in the Michaelis complexes were
inferred from the coordinates of the corresponding dimethyl-
and trimethyl-lysine products, respectively. In addition, we
modeled the -amino group in a deprotonated state with its
hydrogen atoms oriented toward the hydrogen bond acceptors
that align the lysyl side chain for methylation. As a basis for this
comparison, we first modeled the monomethylation reaction
catalyzed by WT SET7/9 (Fig. 4A). In the substrate ternary
complex, the lysine -amine is aligned with the methyl group
and sulfonium cation of AdoMet through a hydrogen bond to
the Tyr-245 hydroxyl group and water 1 in the solvent pocket.
The values of the reaction distance and angle are 2.8 Å and 153°,
respectively, in approximate agreement with the linear geome-
try of a SN2 methyl transfer reaction calculated in other mod-
eled substrate complexes (8, 10). In the product complex, the
monomethyl-lysine side chain is bound in an extended confor-
mation with its methyl group oriented within the methyl trans-
fer pore, thereby obstructing AdoMet binding. Furthermore,
water 1 remains tightly coordinated in the solvent pocket
through four hydrogen bonds to Gly-292, Ala-295, Tyr-305,
and the monomethyl -amino group. These interactions hinder
the dissociation of water 1 and the related rearrangement of the
monomethyl-lysine side chain required for a second methyl
transfer reaction, explaining why the WT enzyme cannot cata-
lyze di- and trimethylation. These findings concur with the
FIGURE 4. Catalytic models of the methyl transfer reactions catalyzed by
WT SET7/9 and the Y305F mutant. A, monomethylation of TAF10-K189 by
the WT enzyme. The reaction scheme depicts the modeled substrate ternary
complex (left) and the product complex (right) for the transfer of the methyl
group from AdoMet (green carbon atoms) to K189 in TAF10 (yellow carbons),
yielding AdoHcy and K189me1. The red arrow indicates the direction of the
nucleophilic attack of the deprotonated -amino group on the AdoMet
methyl group. The transferred methyl group is colored green, and the white
atoms represent the hydrogens of the -amino group. Hydrogen bonds
and residues in the enzyme active site are illustrated as in Fig. 2. The
reaction distance and angle are labeled in red. B and C, models of the
Y305F mutant for the first methyl transfer reaction with TAF10-K189 (B)
and second methyl transfer reaction with TAF10-K189me1 (C). Color
schemes are the same as in A.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31855
model for SET7/9 product specificity reported in previous
structural and functional studies (6, 11).
Similar reaction geometry is observed in the model for the
monomethyl transfer reaction catalyzed by SET7/9 Y305F.
Hydrogen bonds from the Tyr-245 hydroxyl group and water 1
align the lysine -amino group with the AdoMet methyl group
at a distance of 2.1 Å and an angle of 160°, equivalent to those
measured in the Michaelis complex of the WT enzyme (Fig. 4, A
and B). In the product complex, monomethyl-lysine adopts an
extended trans configuration analogous to that in the WT
enzyme. For dimethylation to occur, the monomethyl-lysine
must undergo a conformational change in which its methyl
group is rotated out of the methyl transfer path with AdoMet.
The structure of the Y305F mutant bound to the dimethylated
TAF10 peptide (Fig. 2D) implies that this rearrangement occurs
through the dissociation of water 1 due to the loss of the Tyr-
305 hydrogen bond in the solvent pocket. The dissociation of
water 1 would enable the monomethyl-lysine side chain to
adopt an alternative conformation through a rotation about its
C–N bond, projecting the methyl group into the solvent
pocket (Fig. 4C). This rotation reorients the methyl group out of
the methyl transfer path while realigning the monomethyl -
amino group for a second methylation reaction through a direct
hydrogen bond to the Tyr-245 hydroxyl group and a CH–O
hydrogen bond between its methyl group and Ala-295. The
modeled reaction geometry for monomethyl-lysine substrate
complex of the Y305F mutant (2.7 Å, 160°) is equivalent to that
of the first methyl transfer reaction in SET7/9 Y305F. These
geometries concur with our previous models for mono- and
dimethylation catalyzed by SET8 Y334F (8), illustrating that the
orientation of a methyl group into the solvent pocket is a con-
served feature of SET domain KMTs that catalyze multiple
methylation.
In addition, we modeled the methyl transfer reactions cata-
lyzed by SET7/9 Y245A (Fig. 5). In the model of the lysine sub-
strate complex, the -amino group is aligned for methyl transfer
by a hydrogen bond to the carbonyl oxygen of Gly-264, result-
ing in a short reaction distance (2.3 Å) and a suboptimal reac-
tion angle (141°) with the methyl group of AdoMet (Fig. 5A).
This misalignment appears to be a direct consequence of the
Y245A mutation that abolishes hydrogen bonding to the -
amino group, illustrating that the suboptimal orientation of the
-amine likely contributes to the diminished activity of this
mutant toward unmodified substrates (11). Conversely, in the
modeled monomethyl-lysine substrate complex for SET7/9
Y245A, Tyr-305 hydrogen bonds to the -amino group, aligning
it for methyl transfer (Fig. 5B). In addition, CH–O hydrogen
bonds to the monomethyl-lysine methyl group and the dissoci-
ation of water 2 from the active site also contribute to reposition-
ing the -amino group for dimethylation. Collectively, these inter-
actions orient the -amine in a reaction angle of 165° that is more
conducive to methyl transfer. However, the reaction distance for
dimethylation is 0.6 Å longer than that in the corresponding
Y305F model because Tyr-305 is positioned further from
AdoMet than Tyr-245 (Figs. 4C and 5B).
In the third methyl transfer reaction catalyzed by SET7/9
Y245A, the lone pair of electrons of the dimethyl-lysine -
amino group acts as the nucleophile and thus cannot engage in
hydrogen bonding. The structure of the trimethyl-lysine prod-
uct complex (Fig. 3D) implies that the dimethyl -amine is
aligned via CH–O hydrogen bonds to its methyl groups, as
shown in the model of the Michaelis complex for this reaction
(Fig. 5C). These CH–O hydrogen bonds restrain the orienta-
tion of the -amino group and position one of the methyl
groups into the solvent pocket, displacing water 1 as discussed
earlier (Fig. 3, D and E). These interactions cumulatively align
the -amino group and AdoMet methyl group with a reaction
distance of 3.0 Å and angle of 162° (Fig. 5C). Taken together, the
models of the substrate complexes for SET7/9 Y245A suggest
that CH–O hydrogen bonds play an increasingly important role
in aligning the methylated -amino group in successive rounds
of methyl transfer.
DISCUSSION
The structural and functional characterization of the
SET7/9 Y245A and Y305F mutants presented here yields
new insights into the mechanism underlying the product
specificity of SET domain KMTs. Importantly, it resolves a
general paradox concerning this specificity. How does the
active site constrain the motion of the lysine -amino group
to align it for methyl transfer with AdoMet, while providing
adequate volume to accommodate the mono-, di-, and tri-
methylated lysine side chain generated during multiple
methyl transfer reactions? The structures of the Y305F and
Y245A mutants resolve this paradox, illustrating that alter-
ations in the positions or occupancies of water molecules
within their active sites generate the space required to
FIGURE 5. Models for the methyl transfer reactions catalyzed by the
SET7/9 Y245A mutant. Models of the Y245A mutant for the first methyl
transfer reaction with TAF10-K189 (A), the second methyl transfer reaction
with TAF10-K189me1 (B), and the third methyl transfer reaction with TAF10-
K189me2 (C). Residues and hydrogen bonds are depicted as in Fig. 3.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
accommodate the multiply methylated -amine produced
during successive catalytic cycles. Minor perturbations in
the side chains of certain active site residues, such as Tyr-
305, are also observed in alignments of the WT enzyme and
the Y245A and Y305F complexes, although these changes are
modest compared with the displacement or dissociation of
the water molecules in the active site. These findings suggest
that the waters function as transient place holders that facil-
itate the SN2 methyl transfer reaction. During monomethy-
lation, they function to constrain the movement of the lysine
-amino group by mediating hydrogen bonds between the
substrate and enzyme, thereby promoting the linear align-
ment with the methyl group and sulfonium cation of
AdoMet (Fig. 4, A and B). During di- and trimethylation, the
water molecules either relocate within the lysine binding
channel or dissociate from the enzyme, yielding the space
required to rotate the methyl group away from the methyl
transfer pore and to realign the -amine in productive geom-
etry for the next methyl transfer reaction (Figs. 4C and 5, B
and C). These findings agree with our prior analysis of the
SET8 Phe/Tyr switch mutant in which we demonstrated that
the Y334F substitution attenuates hydrogen bonding to the
water molecule bound in the solvent pocket, promoting its
dissociation and the conformational changes necessary for
lysine dimethylation (8). Indeed, there is a nearly identical
alignment of the dimethyl-lysine side chains in the structures of
SET7/9Y305FandSET8Y334Fcomplexes,despitethedifferences
in the orientations of the Phe-305 and Phe-334 side chains in each
structure (Fig. 2F). Finally, both the SET8 Y334F (8) and SET7/9
Y305F mutants (Table 2) displayed diminished catalytic efficien-
cies for lysine dimethylation versus monomethylation. These dif-
ferences may reflect the kinetics of the reorganization within the
active site, including the dissociation of the water molecule from
the solvent pocket and the concomitant realignment of the
monomethyl-lysine into a productive geometry for dimethylation.
In addition to their place-holding role, the active site waters
may also facilitate the deprotonation of the lysine -amino
group between methyl transfer reactions. For methylation to
occur, the -amino group must be deprotonated to function as
the nucleophile in the SN2 methyl transfer reaction with
AdoMet (Figs. 4 and 5). Although the pKa value of the lysine
-amine in solution is 10.5, molecular dynamics simulations
by Zhang and Bruice (25, 26) indicate that this value diminishes
to 8.2 upon formation of the SET7/9 Michaelis complex due to
the proximity of the AdoMet sulfonium cation and the low
dielectric constant of the active site. Furthermore, their simu-
lations show that a chain of water molecules facilitates the dep-
rotonation of the -amino group prior to methyl transfer, trans-
ferring the proton to bulk solvent. Although these water
molecule chains are not evident in our crystal structures, the
Y305F and Y245A complexes suggest another potential mech-
anism for deprotonation. In the dimethyl-lysine complexes of
the Y305F and Y245A mutants, the dissociation of water 1 and
2, respectively, from the lysine binding channel requires that
the solvent-mediated hydrogen bond to the -amino group is
broken (Figs. 2D and 3C). It is conceivable that these waters
dissociate from the active site as hydronium ions, promoting
the realignment and deprotonation of the methyl -amino
group for the next methyl transfer reaction.
A comparison of the SET7/9 Y305F and SET8 Y334F com-
plexes yields insights into the mechanism by which the Phe/Tyr
switch influences water binding within the solvent pocket. The
phenylalanine substitution in the Phe/Tyr switch results in the
loss of a single hydrogen bond to the water molecule (water 1) in
the solvent pocket compared with the four hydrogen bonds that
coordinate the solvent molecule in WT SET7/9 (Fig. 2, A and B)
and SET8 (7, 8). Although this attenuation in hydrogen bonding
may appear insignificant, this difference is nonetheless impor-
tant for at least two reasons. First, theoretical calculations indi-
cate that, on average, water molecules form 3.5 hydrogen
bonds in solutions (40, 41). This value is greater than the num-
ber of hydrogen bonds coordinating water 1 in the solvent
pocket in SET7/9 Y305F (Fig. 2, B and C) as well as in SET8
Y334F and other di- and trimethyltransferases that possess a
hydrophobic residue in the Phe/Tyr switch site (8). From the
perspective of the water molecule, the greater hydrogen bond-
ing potential in solution would tend to thermodynamically
favor its dissociation from the solvent pocket in SET domain
KMTs that lack a tyrosine in the Phe/Tyr switch position. Sec-
ond, the ordered binding of water molecules observed in the
active sites of SET domain ternary complexes represents an
unfavorable entropy compared with their diffusion in bulk sol-
vent. In WT SET7/9 (Fig. 2A) and SET8 (7, 8), this entropic
penalty can be partially offset through the favorable enthalpy of
binding associated with the four hydrogen bonds that coordi-
nate the water within the solvent pocket. It is conceivable that
the loss of the tyrosine-mediated hydrogen bond in the Phe/Tyr
switch shifts the equilibrium in favor of dissociation of the
water molecule from the solvent pocket, thereby facilitating
dimethylation in SET7/9 Y305F, SET8 Y334F, and other di- and
trimethyltransferases.
The structures of the SET7/9 Y245A and Y305F complexes
illustrate the interactions that align the lysine -amino group
during the methyl transfer reactions in each enzyme. In the WT
enzyme and the Y305F mutant, hydrogen bonding to the
hydroxyl group of Tyr-245 appears to be critical in properly
aligning the -amine for methyl transfer (Fig. 4). Tyr-245 is
conserved in the sequences of many SET domain KMTs (8, 42),
and substitutions of this residue generally impair or abolish
activity, indicating its importance in catalysis (8, 43). However,
SET7/9 appears to be an exception to this rule, as the Y245A
mutant is not only active but is capable of catalyzing lysine
trimethylation. In this mutant, Tyr-305 appears to assume the
role of Tyr-245 by hydrogen bonding to the monomethylated
-amino group to align it for methyl transfer with AdoMet, as
illustrated in the modeled substrate complex for the dimethy-
lation reaction (Fig. 5B). Conversely, in the model for trimethy-
lation, the Tyr-305 hydroxyl group does not hydrogen bond to
the -amine but instead participates in a CH–O hydrogen bond
with one of the methyl groups to assist in aligning the dimethy-
lated -amine for the methyl transfer reaction (Fig. 5C). Addi-
tional structural and functional studies of the SET domain tri-
methyltransferases will aid in further illuminating the roles
of CH–O hydrogen bonds in facilitating lysine multiple
methylation.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31857
Acknowledgments—We acknowledge S. Schiebold for assistance in
protein expression, purification, and crystallization and S. Anderson
and R. Sanishvili for their assistance with x-ray data collection. We
also thank S. Bulfer and S. Horowitz for reading the manuscript and
providing useful comments. This work utilized the Protein Structure
Facility of the Michigan Diabetes Research and Training Center, Uni-
versity of Michigan, supported by National Institutes of Health Grant
DK020572, NIDDK. Use of the Advanced Photon Source was sup-
ported by the United States Department of Energy, Basic Energy Sci-
ences, Office of Science, under Contract DE-AC02-06CH11357.
GM/CA CAT has been funded in whole or in part by National
Institutes of Health NCI Grant Y1-CO-1020 and NIGMS Grant
Y1-GM-1104. Use of the LS-CAT Sector 21 was supported by Michi-
gan Economic Development Corporation and the Michigan Technol-
ogy Tri-Corridor Grant 085P1000817 for the support of this research
program.
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Lysine Methylation by SET7/9 Mutants
31858
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
|
3M58
|
SET7/9 Y245A in complex with TAF10-K189me1 peptide and AdoHcy
|
SET7/9 Catalytic Mutants Reveal the Role of Active Site Water
Molecules in Lysine Multiple Methylation*□
S
Received for publication,February 15, 2010, and in revised form, July 14, 2010 Published, JBC Papers in Press,August 1, 2010, DOI 10.1074/jbc.M110.114587
Paul A. Del Rizzo‡, Jean-Franc¸ois Couture‡§1, Lynnette M. A. Dirk¶, Bethany S. Strunk‡, Marijo S. Roiko‡,
Joseph S. Brunzelle, Robert L. Houtz¶, and Raymond C. Trievel‡2
From the ‡Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109, the §Ottawa Institute of
Systems Biology, Department of Biochemistry, Microbiology, and Immunology, University of Ottawa, Ottawa, Ontario K1H 8M5,
Canada, the ¶Department of Horticulture, Plant Physiology/Biochemistry/Molecular Biology Program, University of Kentucky,
Lexington, Kentucky 40546, and the Department of Molecular Pharmacology and Biological Chemistry, Northwestern University
Center for Synchrotron Research, Life Sciences Collaborative Access Team, Argonne, Illinois 60439
SET domain lysine methyltransferases (KMTs) methylate
specific lysine residues in histone and non-histone substrates.
These enzymes also display product specificity by catalyzing dis-
tinct degrees of methylation of the lysine -amino group. To
elucidate the molecular mechanism underlying this specificity,
we have characterized the Y245A and Y305F mutants of the
human KMT SET7/9 (also known as KMT7) that alter its prod-
uct specificity from a monomethyltransferase to a di- and a tri-
methyltransferase, respectively. Crystal structures of these
mutants in complex with peptides bearing unmodified, mono-,
di-, and trimethylated lysines illustrate the roles of active site
water molecules in aligning the lysine -amino group for methyl
transfer with S-adenosylmethionine. Displacement or dissocia-
tion of these solvent molecules enlarges the diameter of the
active site, accommodating the increasing size of the methylated
-amino group during successive methyl transfer reactions.
Together, these results furnish new insights into the roles of
active site water molecules in modulating lysine multiple meth-
ylation by SET domain KMTs and provide the first molecular
snapshots of the mono-, di-, and trimethyl transfer reactions
catalyzed by these enzymes.
SET domain enzymes represent a family of S-adenosylmethi-
onine (AdoMet)3-dependent methyltransferases that catalyze
the site-specific methylation of protein lysyl residues in a host
of proteins, including histones, transcription factors, chroma-
tin-modifying enzymes, ribosomal subunits, and other sub-
strates (1–3). In many instances, these modifications serve to
recruit effector proteins that recognize methyl-lysyl residues in
a sequence-dependent fashion (4). In addition, SET domain
KMTs exhibit product specificity, defined as their ability to cat-
alyze mono-, di-, or trimethylation of the lysine -amino group.
This specificity is biologically relevant because many methyl-
lysine-binding proteins can discriminate among different
degrees of lysine methylation (4). Thus, both the site and degree
of lysine methylation are critical to recognition by effector
proteins.
Structural and functional studies have identified a Phe/Tyr
switch in the active site of SET domain KMTs that governs their
respective product specificities (5, 6). According to this model,
KMTs that possess a tyrosine in the Phe/Tyr switch site are
limited to catalyzing lysine monomethylation, whereas en-
zymes that possess a phenylalanine or another hydrophobic
residue in this position display di- or trimethyltransferase activ-
ity. Mutational analysis of various SET domain KMTs, includ-
ing DIM-5 (KMT1) (6), SET7/9 (6), G9A (KMT1C) (5), and
SET8 (also known as PR-SET7 and KMT5A) (7, 8), has demon-
strated that substitutions in the Phe/Tyr switch result in pre-
dictable changes in product specificity. Several models have
been proposed to explain the mechanism by which the Phe/Tyr
switch site governs this specificity, including variations in the
diameter of the active site due to the size of Phe/Tyr switch
residue and steric hindrance by the tyrosine hydroxyl group (6,
9–11). However, our recent studies of the Phe/Tyr switch
mutant Y334F in the human histone H4 Lys-20 (H4K20) meth-
yltransferase SET8 indicate that the Phe/Tyr switch regulates
product specificity via a more subtle mechanism (8). Specifi-
cally, the switch modulates the binding of an active site water
molecule that in turn regulates the transition from mono-
methylation to multiple methylation.
Among the KMTs that have been structurally characterized,
SET7/9 has emerged as an archetypal model for studying the
catalytic mechanism and product specificity of the SET domain
family. Although initially isolated as a histone H3 Lys-4
(H3K4)-specific methyltransferase, this KMT has been shown
to regulate the functions of numerous non-histone substrates
through site-specific methylation (12–21). Early structural and
functional studies of SET7/9 identified two active site mutants,
* Thisworkwassupported,inwholeorinpart,byNationalInstitutesofHealth
Grant R01 GM073839 (to R. C. T.) and National Institutes of Health Admin-
istrative Supplement GM073839-04S1 (to R. C. T.) funded through the
American Recovery and Reinvestment Act. This work was also supported
by Department of Energy Grant DE-FG02-92ER20075 (to R. L. H.).
□
S The on-line version of this article (available at http://www.jbc.org) contains
supplemental Table 1.
The atomic coordinates and structure factors (codes 3M53, 3M54, 3M55, 3M56,
3M57,3M58,3M59,and3M5A)havebeendepositedintheProteinDataBank,
Research Collaboratory for Structural Bioinformatics, Rutgers University, New
Brunswick, NJ (http://www.rcsb.org/).
1 Supported by a Canadian Institutes of Health Research postdoctoral
fellowship.
2 To whom correspondence should be addressed: Dept. of Biological Chem-
istry, University of Michigan Medical School, 1150 West Medical Center Dr.,
5301 MSRB III, Ann Arbor, MI 48109. Tel.: 734-647-0889; Fax: 734-763-4581;
E-mail: rtrievel@umich.edu.
3 The abbreviations used are: AdoMet, S-adenosylmethionine; AdoHcy, S-ad-
enosylhomocysteine; KMT, lysine methyltransferase; ITC, isothermal titra-
tion calorimetry; CH–O, carbon-oxygen hydrogen bond; BisTris, 2-[bis(2-
hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 41, pp. 31849–31858, October 8, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31849
Y245A and Y305F, which change its product specificity. The
Phe/Tyr switch mutant Y305F alters SET7/9 product specificity
from a mono- to dimethyltransferase (6), whereas the Y245A
substitution converts the enzyme into a trimethyltransferase
with weak monomethyltransferase activity (11). These mutants
have been the subjects of numerous molecular modeling simu-
lations that have led to various models to explain their distinct
product specificities (22–26). However, the lack of structural
data for the SET7/9 Y245A and Y305F mutants in complex with
cognate methylated peptides has hindered our understanding
of the mechanisms that define the respective product specific-
ities of these mutants. Moreover, these structures would yield a
framework for visualizing the mono-, di-, and trimethylation
reactions catalyzed by SET domain KMTs.
To gain insight into the molecular basis of their product
specificities, we have determined high resolution crystal
structures of the SET7/9 Y245A and Y305F mutants in com-
plex with peptides of the TATA box-binding protein-associ-
ated factor TAF10 bearing the Lys-189 methylation site in
unmodified (K189), monomethylated (K189me1), dimethyl-
ated (K189me2), and trimethylated (K189me3) states. The
structures and accompanying biochemical data support a
model whereby changes in the occupancy or position of water
molecules in the active site are critical in establishing the prod-
uct specificities of the SET7/9 Y245A and Y305F mutants.
Together, our results provide new insights into the mechanisms
that govern SET domain product specificity and provide step-
wise snapshots of the lysine mono-, di-, and trimethyl transfer
reactions catalyzed by KMTs.
EXPERIMENTAL PROCEDURES
Cloning, Expression, and Purification of the SET7/9 Mutants—
The Y245A and Y305F mutants were introduced into the pHIS2
SET7/9 expression vector encoding residues 110–366 (27) via
QuikChange site-directed mutagenesis (Stratagene) and were
verified by dideoxy DNA sequencing. The plasmids encoding
wild type (WT) SET7/9 and the Y245A and Y305F mutants
were transformed into Rosetta2 DE3 cells (Novagen) and were
expressed as described previously (27, 28). In the course of
characterizing WT SET7/9, we observed that the enzyme co-
purified with AdoMet or another contaminant that resulted in
technical difficulties in the isothermal titration calorimetry
(ITC) experiments and co-crystallization trials with the TAF10
peptides. To overcome this problem, a denaturation and refold-
ing step was inserted in the purification scheme. The denatur-
ation and refolding protocol involved adding 6 M guanidine
HCl, 50 mM sodium phosphate, pH 7.5, and 500 mM NaCl to the
protein while it was immobilized on a nickel-Sepharose column
(GE Healthcare). The column was washed with this buffer, fol-
lowed by a solution of 6 M urea, 50 mM sodium phosphate, pH
7.5, 150 mM NaCl, and 10 mM 2-mercaptoethanol to remove the
cofactor from the denatured enzyme. A reverse gradient from 6
to 0 M urea was then performed in the same buffer to refold the
protein, which was subsequently eluted from the column using
a linear gradient of 0–500 mM imidazole in 50 mM sodium
phosphate, pH 7.5, 500 NaCl, and 10 mM 2-mercaptoethanol.
The refolded protein was digested with tobacco etch virus pro-
tease (29) during dialysis against 20 mM Tris, pH 7.5, 150 mM
NaCl, and 5 mM 2-mercaptoethanol and then purified using a
Superdex 200 gel filtration column (GE Healthcare). Protein
concentration was determined by its absorbance at 280 nm.
Synthetic Peptides—The TAF10 peptides bearing K189,
K189me1,
K189me2,
and
K189me3
(sequence,
acetyl-
SKSK189DRKYTL-amide) and a biotinylated TAF10 peptide
(sequence,
acetyl-SKSK189DRKYTLT(K-EZLink-S-S-biotin)-
amide) were synthesized and purified by New England Peptide,
Inc. Peptide concentrations were measured using the absorb-
ance of their tyrosine residue at 274 nm.
Crystallization and Data Collection—Crystals were pro-
duced by hanging drop vapor diffusion by mixing the crystalli-
zation solution in a 1:1 ratio with 10–20 mg/ml SET7/9, 1 mM
S-adenosylhomocysteine (AdoHcy), and 1.0–3.0 mM unmodi-
fied or methylated TAF10-K189 peptide in 20 mM Tris, pH 8.0,
100 mM NaCl, and 2 mM tris(2-carboxyethyl)phosphine. Crys-
tals were obtained at 20 °C in either 1.8–2.0 M (NH4)2SO4, with
0.1 M BisTris, pH 6.2–6.6, or in 0.95–1.1 M sodium citrate with
100 mM imidazole pH 8.0–8.4. In both crystallization condi-
tions, the final pH values were between pH 8.0 and 9.0. Crystals
in the (NH4)2SO4 condition were typically flash-frozen in the
mother liquor containing 25–30% glycerol, and the crystals in
the citrate condition were frozen in 1.6 M sodium citrate. Data
were collected at the Advanced Photon Source beamlines
21-IDG (LS-CAT) and 23-IDD (GM/CA-CAT). Images were
indexed, integrated, and scaled using HKL2000 (30). Structures
of the mutants were solved by molecular replacement using
MOLREP (31) with the coordinates of a previously reported
SET7/9 ternary complex used as the search model (Protein
Data Bank code 2F69). Successive rounds of model building and
refinement were carried out using Coot (32) and REFMAC (33),
respectively. The geometry of the models were verified by Mol-
Probity (34). Simulated annealing omit maps were calculated
using CNS (35) with the peptide and cofactor removed to elim-
inate model bias in the active site. Structural figures were ren-
dered using PyMOL (Schro¨dinger, LLC.).
Fluorescent Methyltransferase Assay—A coupled fluorescent
methyltransferase assay was used to measure the kinetic
parameters of WT SET7/9 and the Y245A and Y305F mutants
as reported previously, with the exception that 50–150 nM
enzyme, 100 M AdoMet, and varying concentrations of TAF10
peptide substrate were used (27, 36). Assays were performed in
triplicate, and a homocysteine calibration curve was used to
calculate the initial velocities. Kinetic parameters were calcu-
lated by plotting the velocities versus peptide concentration and
by fitting the Michaelis-Menten equation to the data via non-
linear regression using Prism 5.0 (GraphPad). In cases where
the Km value was beyond the measurable range of the assay, the
kcat/Km value was determined as described previously (7).
Calorimetry Experiments—ITC was performed at 20 °C using
a MicroCal VP-ITC calorimeter (GE Healthcare) with 0.12 mM
protein and 4 mM AdoHcy in 20 mM sodium phosphate, pH 7,
and 100 mM NaCl with 1.5 mM peptide as the injectant. Data
were processed, and equilibrium dissociation constants (KD)
and curve fitting errors were calculated from the binding iso-
therms using Origin 7.0 (OriginLab Corp.). WT SET7/9 and the
Y245A and Y305F mutants displayed ligand:protein binding
stoichiometries (N values) between 0.8 and 1.0, demonstrating
Lysine Methylation by SET7/9 Mutants
31850
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
that WT SET7/9 and its mutants were properly refolded due to
their ability to bind peptides in an 1:1 molar ratio.
TLC Product Analysis—Methyltransferase assays were per-
formed in triplicate at 37 °C with the biotinylated TAF10 pep-
tide (0.5 mM) for 30 min in the presence of WT SET7/9 (3
pmol), Y305F mutant (6 pmol), or the Y245A mutant (100
pmol). Assays contained 100 mM HEPES, pH 8.0, 50 mM NaCl,
1.0 mM [methyl-3H]AdoMet (diluted to a specific activity of 0.2
Ci/nmol with purified AdoMet (37)), 15 M Sulfolobus solfa-
taricus AdoHcy hydrolase (36), and 2 units of adenosine deami-
nase (Roche Applied Science) in a final volume of 20 l. The
reactions were terminated by addition of an equal volume of
200 mM MES, pH 5.1, prior to addition of a 2-fold molar excess
of immobilized avidin resin (UltraLink; Pierce). Biotinylated
peptides were allowed to bind at room temperature for 30 min,
and the resin was then collected by centrifugation (9000 g).
The resin was washed three times with 300 mM NaCl, and the
peptide was eluted overnight from the avidin resin by cleavage
of the disulfide bond in the linker of the peptide using 10 mM
tris(2-carboxyethyl)phosphine (Thermo Scientific). The resin
was incubated with additional 10 mM tris(2-carboxyethyl)phos-
phine the following day until the radiolabel was essentially
removed from the resin. The recovered peptides were hydro-
lyzed by incubation with 6 M HCl at 110 °C for 24 h. Subsequent
steps in measuring the radiolabel incorporated into the mono-,
di-, and trimethyl-lysine products were performed as reported
previously (8).
RESULTS
Functional Analysis of the SET7/9
Y305F Mutant—Prior studies of
SET7/9 by Zhang et al. (6) reported
that mutation of the Phe/Tyr switch
residue Tyr-305 to a phenylalanine
alters its product specificity from a
mono- to dimethyltransferase. We
verified these findings by demon-
strating that WT SET7/9 mono-
methylated the TAF10-K189 pep-
tide, whereas the Y305F mutant
mono- and dimethylated this sub-
strate, as demonstrated by mass
spectrometry (data not shown). We
next examined whether the Y305F
substitution altered the affinity of SET7/9 for the TAF10-K189
peptides using ITC (Fig. 1). A comparison of the KD values
revealed that SET7/9 Y305F bound the TAF10-K189 and
TAF10-K189me1 peptides 4- and 6-fold more tightly, respec-
tively, than the WT enzyme, whereas this mutant displayed a
substantially diminished affinity for the TAF10-K189me2 pep-
tide (Table 1). Although the WT enzyme and the Y305F mutant
exhibited discernable differences in their affinities for the
unmodified and monomethylated peptides, these variations are
modest and cannot account for their distinct product specific-
ities, suggesting that a kinetic effect during methylation may be
responsible.
To investigate this possibility, we characterized the kinetic
parameters of WT SET7/9 and the Y305F mutant using the
TAF10 peptides as substrates. Both enzymes methylated the
unmodified peptide with comparable kcat and Km values (Table
2). In analyzing the kinetic parameters for the methylation
of the monomethylated peptide by SET7/9 Y305F, we found
that this substrate displayed an elevated Km value that was
beyond the measurable range of the assay due to its limited
solubility. In this case, we measured the catalytic efficiency
(kcat/Km) for the methylation of this peptide and found that it
was methylated 15-fold less efficiently than the unmodified
peptide by SET7/9 Y305F. Given the fact that the Y305F mutant
exhibited a higher binding affinity for the TAF10-K189me1
peptide than the WT enzyme (Table 1), the kinetic data suggest
that a step in the reaction pathway following substrate binding
limits the catalytic efficiency of this mutant.
We next examined whether the Y305F mutant dimethylated
the TAF10-K189 peptide via a processive or a distributive
mechanism. In a processive mechanism, the methyl-lysine
substrate would remain bound to the enzyme during successive
methyl transfer reactions; thus, the concentration of an inter-
mediate, such as monomethyl-lysine, cannot exceed the en-
zyme concentration during the assay. In a distributive mecha-
nism, the intermediates are released into solution where they
accumulate prior to the next round of methylation, resulting in
an intermediate concentration that is greater than that of the
enzyme. Using a radiometric TLC assay and a biotinylated
TAF10 peptide, we quantified the amounts of monomethylated
products generated by the WT SET7/9 and the Y305F mutant
FIGURE 1. ITC analysis of WT SET7/9 and the Y245A and Y305F mutants. Representative titrations and
binding isotherms for the calorimetry experiments are shown for the unmodified TAF10-K189 peptide titrated
into WT SET7/9 (left plot), SET7/9 Y305F (middle plot), and SET7/9 Y245A (right plot). In each panel, the ITC
titration experiment (upper panel) is shown with the binding isotherms (lower panel) fit to a single binding site
model.
TABLE 1
Analysis of the binding affinity of WT SET7/9 and its catalytic
mutants for unmodified and methylated TAF10 peptides
SET7/9
TAF10 peptide
KD
a
M
WT
K189
4.9 0.20
WT
K189me1
4.0 0.36
Y305F
K189
1.3 0.10
Y305F
K189me1
0.62 0.065
Y305Fb
K189me2
70
Y245A
K189
4.0 0.25
Y245A
K189me1
3.3 0.10
Y245A
K189me2
5.8 0.22
Y245A
K189me3
11 0.28
a Curve fitting errors were calculated from the binding isotherms.
b An estimate of the affinity is reported due to weak peptide binding.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31851
(Table 3). The data demonstrate that comparable amounts of
monomethyl-lysine were generated when the quantity of
enzyme usedistakenintoaccount,inagreementwiththeirsimilar
turnover numbers for the TAF10-K189 peptide (Table 2). The
Y305F mutant also produced small but measurable quantities of
radiolabeled dimethyl-lysine product that were substantially
smaller than the amount of monomethyl-lysine generated.
Therefore, the TLC data are consistent with a distributive
mechanism for dimethylation by the Y305F mutant because the
amount of monomethyl-lysine produced exceeded the quantity
of enzyme used in the assay.
Structures of WT SET7/9 and the Y305F Mutant in Complex
with Unmodified and Methylated TAF10 Peptides—To deter-
mine the mechanism by which the Y305F substitution alters the
product specificity of SET7/9, we determined the crystal struc-
tures of this mutant bound to AdoHcy and TAF10-K189,
TAF10-K189me1, and TAF10-K189me2 peptides and com-
pared these to the structures of the WT SET7/9AdoHcy
TAF10-K189 complex (supplemental Table 1). The structures
of these complexes were determined to 1.85 Å or higher reso-
lution, permitting unambiguous modeling of the K189 side
chains in the active site of the enzyme based on simulated
annealing omit maps (Fig. 2). The ternary complexes of the WT
and the Y305F mutant superimpose with overall root mean
square differences of less than 0.3 Å for all aligned atoms, indi-
cating that neither the Y305F mutation nor the binding of the
various TAF10-K189 peptides results in substantial changes in
its overall structure.
An inspection of the active sites of the SET7/9 WT and
Y305F complexes illustrates the binding modes of the unmod-
ified and methylated forms of K189 in the TAF10 peptides (Fig.
2, A–D). The K189 side chain binds in an extended all trans
conformation in a deep pocket, termed the lysine binding chan-
nel, that is composed of Tyr-245, Thr-266, Leu-267, Ser-268,
Tyr-305, Tyr-335, and Tyr-337 (not shown for clarity) (Fig. 2A).
These residues interact with the aliphatic portion of the K189
side chain primarily through van der Waals contacts. The lysine
binding channel connects to the AdoMet-binding site on the
opposite face of the catalytic domain via an oxygen-lined
methyl transfer pore (38). During catalysis, the methyl group of
the cofactor is positioned within the methyl transfer pore
for the SN2 reaction with the -amino group of the lysine or
methyl-lysine substrate (see below).
To lower the activation barrier for this reaction, the lysine
-amine nucleophile is aligned for methyl transfer through a
hydrogen bond network within the active site. In the WT
enzyme, the K189 -amino group hydrogen bonds to the
hydroxyl group of Tyr-245 as well as to two water molecules
(Fig. 2A). One of the water molecules (termed water 1), is coor-
dinated in a solvent pocket, through hydrogen bonds to the
carbonyl oxygens of Gly-292 and Ala-295 and to the hydroxyl
group of the Phe/Tyr switch residue Tyr-305. This solvent
pocket is structurally conserved in SET domain KMTs and has
an important role in defining product specificity through the
adjacent Phe/Tyr switch residue, as shown in our prior studies
of the human H4K20 methyltransferase SET8 (8). The other
water molecule is bound within the methyl transfer pore
between the lysine substrate and the thioether sulfur atom of
AdoHcy through hydrogen bonds to Tyr-245, Asn-265, and
His-293 in SET7/9 and the TAF10-K189 -amino group. This
water is not observed in other structures of SET7/9 ternary
complexes and may represent the approximate position that
the AdoMet methyl group occupies in the methyl transfer pore
in the Michaelis complex.
In structures of the Y305F ternary complexes, the K189,
K189me1, and K189me2 side chains also adopt extended trans
side chain geometries within the lysine binding channel that are
stabilized via hydrogen bonding to Tyr-245 in the enzyme (Fig.
2, B–D). The orientations of the K189me1 and K189me2 side
chains are further maintained through carbon-oxygen (CH–O)
hydrogen bonding between the methyl groups and oxygen
atoms within the vicinity of the methyl transfer pore, as
reported previously in other SET domain KMT structures (8,
10, 38). A superimposition of the SET7/9 WT and Y305F com-
plexes underscores the similarity of the lysyl binding conforma-
tions (Fig. 2E). However, there are notable differences in the
hydrogen bond patterns and occupancy of water 1 within the
solvent pocket in the Y305F mutant compared with the WT
enzyme. Specifically, the Y305F substitution results in the loss
of one hydrogen bond to water 1 in the structures of the TAF10-
K189 and TAF10-K189me1 complexes (Fig. 2, B and C). In con-
trast, water 1 is absent in TAF10-K189me2 complex, and the
vacated solvent pocket is occupied by one of the methyl groups
TABLE 2
Kinetic parameters of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
TAF10 peptide substrate
Km
a
kcat
a
kcat/Km
a
M
min1
M1 min1 103
WT
K189
160 17
17 0.62
110 17
Y305F
K189
88 5.0
17 0.30
190 11
Y305Fb
K189me1
11 0.50
Y245A
K189
200 35
0.53 0.04
2.6 0.47
Y245A
K189me1
210 23
5.9 0.23
28 3.3
Y245A
K189me2
400 29
6.5 0.16
15 1.2
a Curve fitting errors were calculated from the hyperbolic fits for the Michaelis-Menten equation.
b Km and kcat were not determined because the Km value was beyond the measurable range; therefore, the kcat/Km value is reported.
TABLE 3
Product analysis of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
Quantity
of enzyme
Measured
product
Amount of
product formeda
nmol
nmol
WT
0.003
Kme1
0.65 0.07
Y305F
0.006
Kme1
1.5 0.49
Kme2
0.033 0.009
Y245A
0.100
Kme1
0.80 0.22
Kme2
0.39 0.021
Kme3
0.076 0.019
a Standard deviation was calculated from triplicate measurements.
Lysine Methylation by SET7/9 Mutants
31852
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
of the dimethyl -amine (Fig. 2D). This methyl group forms a
3.5-Å CH–O hydrogen bond to the carbonyl oxygen of Ala-295,
further stabilizing the binding of the dimethyl-lysine side chain.
A homologous dimethyl-lysine-binding mode and CH–O
hydrogen bond was observed in our prior structural studies of
the SET8 Y334F Phe/Tyr switch mutant that confers an analo-
gous change in product specificity from a mono- to a dimeth-
yltransferase (8). A structural alignment of the active sites of the
SET7/9 Y305F and SET8 Y334F mutants bound to cognate
dimethylated peptides illustrates that the coordinates of the
dimethyl-lysyl side chains are virtually superimposable, with
one methyl group oriented toward the methyltransfer pore and
the second positioned within the vacant solvent pocket (Fig.
2F). Taken together, the structures of the SET7/9 Y305F com-
plexes and the similarities in the dimethyl-lysine conforma-
tions in the SET7/9 Y305F and SET8 Y334F mutants imply that
the Phe/Tyr switch governs product specificity through a con-
served mechanism whereby it indirectly influences the binding
modes of the methyl-lysine side chain by modulating the affin-
ity of the water molecule (water 1) bound in the solvent pocket.
Biochemical Characterization of
the SET7/9 Y245A Mutant—Previ-
ous studies by Xiao et al. (11)
reported that the Y245A mutation
yields an unusual change in the
product
specificity
of
SET7/9,
converting the enzyme to a trimeth-
yltransferase with weak monometh-
yltransferase activity. We deter-
mined that the SET7/9 Y245A could
mono-, di-, and trimethylate the
TAF10-K189 peptide by mass spec-
trometry (data not shown) and TLC
(Table 3), confirming the earlier
studies of Xiao et al. (11). ITC
analysis revealed that the Y245A
mutant displayed comparable KD
values
for
the
unmodified
and
methylated TAF10-K189 peptides
(Fig. 1), although its affinity for the
trimethylated peptide was modestly
diminished in comparison with the
other peptides (Table 1). The ITC
data demonstrate that the Y245A
mutant
bound
the
unmodified,
mono-,
and
dimethylated
sub-
strates with equivalent affinities,
suggesting that a kinetic effect or
a structural alteration in the active
site may be responsible for its
diminished
activity
toward
un-
modified substrates.
To gain further insight into its
peculiar
product
specificity,
we
characterized the kinetic properties
of
the
SET7/9
Y245A
mutant.
Steady state analysis demonstrated
that this mutant displayed similar
Km values for the unmodified, mono- and dimethylated TAF10
peptides (Table 2). However, the turnover number for the
TAF10-K189 peptide was diminished over 10-fold versus the
methylated peptides and was reduced 30-fold versus the WT
enzyme, in agreement with the weak monomethyltransferase
activity reported by Xiao et al. (11). In addition, we investigated
whether this mutant catalyzes lysine trimethylation via a pro-
cessive or distributive mechanism as described for SET7/9
Y305F. The TLC data illustrate that the mono- and dimethyl-
lysine intermediates accumulated at quantities greater than
that of the enzyme used in the assay, indicating that SET7/9
Y245A obeys a distributive mechanism, analogous to the Y305F
mutant (Table 3).
Structures of SET7/9 Y245A Bound to Unmodified and Meth-
ylated TAF10 Peptides—To elucidate the mechanism underly-
ing its unusual product specificity, we determined the crystal
structures of SET7/9 Y245A in complex with AdoHcy and
unmodified, mono-, di-, and trimethylated TAF10 peptides
(supplemental Table 1). These complexes superimpose with
the structure of the WT SET7/9AdoHcyTAF10-K189 com-
FIGURE 2. Crystal structures of WT SET7/9 and the Y305F mutant in complex with AdoHcy and unmodi-
fied and methylated TAF10 peptides. A, active site of WT SET7/9 bound to AdoHcy and TAF10-K189 and the
active site of SET7/9 Y305F bound to AdoHcy and TAF10-K189 (B), TAF10-K189me1 (C), and TAF10-K189me2
peptides (D). A–D, the K189 residue in the TAF10 peptide is depicted with yellow carbon atoms with the
corresponding Fo – Fc simulated-annealing omit maps (contoured at 2.5 ) shown. AdoHcy is colored with
green carbon atoms, and SET7/9 carbon atoms are colored gray, with the exception of Tyr-305/Y305F
(magenta) and Tyr-245 (cyan). Gly-264 to Ser-268 are shown, and the main chain atoms of Gly-292 to Ala-295
are depicted for clarity. Red dashed lines indicate conventional hydrogen bonds, and blue dashed lines indicate
CH–O hydrogen bonds. Cyan dashed lines indicate weak CH–O hydrogen bonds between 3.5 and 3.7 Å in
length. The water molecule in the solvent binding pocket is numbered 1. E, overlay of the active sites of the WT
enzyme (light blue carbon atoms) and the Y305F mutant (green, yellow, and pink carbon atoms correspond to
K189, K189me1, and K189me2, respectively). Red dashed lines indicate coordination of the waters in the struc-
turesoftheY305Fmutant,andthelightbluedashedlineindicateshydrogenbondingthatonlyoccursintheWT
enzyme. F, superimposition of the active sites of the SET7/9 Y305FAdoHcyTAF10-K189me2 and SET8
Y334FAdoHcyH4K20me2 (Protein Data Bank code 3F9X) complexes shown with green and yellow carbon
atoms, respectively.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31853
plex with root mean squared differences of less than 0.4 Å for all
aligned atoms, indicating that the Y245A mutant does not per-
turb the overall structure of the enzyme. Simulated annealing
omit maps illustrate that K189 side chains are bound within the
lysine binding channel through hydrogen bonds and van der
Waals contacts (Fig. 3, A–D), although the interactions and
binding modes are distinct from those in the complexes of WT
SET7/9 and the Y305F mutant (Fig. 2, A–D). In the unmodified
TAF10 peptide complex, the K189 -amino group forms a weak
hydrogen bond with the carbonyl oxygen of Gly-264 (Fig. 3A),
whereas the -amino groups of K189me1 and K189me2 hydro-
gen bond to the hydroxyl group of Tyr-305 in the mono- and
dimethylated peptide complexes (Fig. 3, B and C). The confor-
mations of the K189me1 and K189me2 side chains are further
stabilized by water-mediated hydrogen bonding and through
CH–O hydrogen bonding to their methyl groups. In the
TAF10-K189me3 peptide complex, the trimethyl-lysine side
chain is coordinated exclusively through direct and water-me-
diated CH–O hydrogen bonds to its methyl groups because the
quaternary -ammonium cation cannot engage in hydrogen
bonding (Fig. 3D).
A structural alignment of the four SET7/9 Y245A complexes
illustrates distinct binding modes for the unmodified versus the
methylated K189 side chains, highlighting the selectivity of this
mutant for methylated substrates. The side chains of K189me1,
K189me2, and K189me3 roughly overlay with their respective
-amino groups superimposed and adopt slightly kinked con-
formations (Fig. 3E), as opposed to the extended trans geome-
try of the unmodified and methylated lysines in the complexes
of the WT enzyme and the Y305F mutant (Fig. 2, A–D). Con-
versely, the unmodified K189 side chain does not superimpose
with its methylated counterparts and is oriented in an alterna-
tive configuration due to its hydrogen bonding to Gly-264 (Fig.
3, A and E). An overlay of the structures of the WT enzyme and
Y245A mutant bound to the unmodified TAF10 peptide illus-
trates that the side chains of K189 do not superimpose and that
the K189 -amino group appears to be misaligned with AdoHcy
in the Y245A complex (Fig. 3F). This suboptimal alignment
may explain the diminished kcat value of SET7/9 Y245A mutant
toward substrates with unmodified lysines (Table 2).
A comparison of the structures of the SET7/9 Y245A and
Y305F complexes yields a molecular explanation for the differ-
ent product specificities of these two mutants. In the SET7/9
Y305F complexes, Tyr-245 aligns the K189 -amino group for
methyl transfer through hydrogen bonding to its hydroxyl
group (Fig. 2, B–D). Conversely, in the Y245A mutant, the
K189me1 and K189me2 -amino groups are oriented through
hydrogen bonding to Tyr-305 (Fig. 3, B and C). These distinct
hydrogen bond patterns impart differences in the conforma-
tions of the lysyl side chains due to the relative orientations of
Tyr-245 and Tyr-305 in the lysine binding channel. Specifically,
the kinked conformation adopted by the K189me1 and
K189me2 side chains in the Y245A complexes (Fig. 3, B and C)
may contribute to the differences in the turnover numbers of
this mutant versus those of the WT enzyme and the Y305F
mutant (Table 2). In addition, the dimethyl -amino group of
the K189me2 side chain binds in distinct orientations in the
Y245A and Y305F mutants due to their hydrogen bonding to
FIGURE 3. Crystal structures of the SET7/9 Y245A mutant in complex with
AdoHcy and unmodified and methylated TAF10 peptides. Active site of
SET7/9 Y245A bound to AdoHcy and TAF10-K189 (A), TAF10-K189me1 (B),
TAF10-K189me2 (C), and TAF10-K189me3 peptides (D). A–D, Fo – Fc simulated-
annealing omit maps (contoured at 2.5 ) for the unmodified and methylated
K189 side chains are illustrated. The residues and hydrogen bonds in each com-
plexarecoloredasinFig.2.Thewatermoleculesinthelysinebindingchannelsof
the Y245A complexes are numbered 1–4, as described in the text. E, superimpo-
sition of the active sites of the Y245A complexes bound to the four methylated
statesofTAF10-K189(unmodified,mono-,di-,andtrimethylatedshowningreen,
yellow, pink, and blue, respectively). F, overlay of WT SET7/9 (blue carbons) and
SET7/9 Y245A (green carbons) in complex with TAF10-K189. Waters correspond-
ing to the WT and Y245A structures are colored cyan and green, respectively.
G, alignment of the active sites of the Y245A (yellow carbons) and Y305F (green
carbons) mutants in complex with the TAF10-K189me2 peptide. Hydrogen
bonds from the Y305F structure are shown as green dashed lines, and waters and
hydrogen bonds in the Y245A structure are shown in yellow and orange,
respectively.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
Tyr-305 and Tyr-245, respectively (Fig. 3G). In the Y305F
mutant, hydrogen bonds to the dimethyl -amino group cou-
pled with steric constraints in the lysine binding channel pre-
vent the K189me2 side chain from undergoing a conforma-
tional change that is conducive to trimethylation (Fig. 2D),
consistent with its dimethyltransferase activity. However, in the
Y245A mutant, the alanine substitution enlarges the diameter
of the lysine binding channel, accommodating trimethyl-lysine
(Fig. 3D). In addition, the larger diameter would permit the
dimethyl-lysine substrate to undergo the conformational reor-
ganization necessary to align the -amino group in a productive
geometry for trimethylation.
A major difference in the active site of the Y245A mutant
versus the other SET7/9 structures is the presence of several
water molecules bound in the cavity generated by the Y245A
mutation. In the structure of the Y245A mutant bound to
TAF10-K189, three water molecules (waters 2–4) occupy this
cavity and are arranged in a triangular geometry (Fig. 3A). In
addition, water 1 shifts 1.6 Å from its position in the solvent
pocket toward water 2 to which it forms a hydrogen bond (Fig.
3, A and E). The shift in water 1 was unexpected given its con-
served orientation in the solvent pocket of the SET7/9 WT and
Y305F complexes (Fig. 2, A–C) as well as in the structures of
other SET domain KMTs (8). This displacement is presumably
related to the alternative conformation of the K189 side chain
whose -amino group is too distant (4.3 Å) to form a productive
hydrogen bond to water 1. Conversely, in the Y245A complexes
bound to TAF10-K189me1 and TAF10-K189me2, water 1
remains tightly bound in the solvent pocket through hydrogen
bonds to Tyr-305 hydroxyl group, the carbonyl oxygens of Gly-
292 and Ala-295, and the K189 -amino group (Fig. 3, B, C, and
E), analogous to its binding in the WT enzyme (Fig. 2A). How-
ever, in the TAF10-K189me3 complex, one of the methyl
groups of the trimethyl -ammonium cation is oriented into the
solvent pocket (Fig. 3D), similar to the dimethyl-lysine binding
mode observed in the Y305F mutant (Fig. 2D). The binding of
the methyl group in the solvent pocket displaces water 1 by 3.2
Å relative to its position in the TAF10-K189me1 complex (Fig.
3E), thereby avoiding a steric clash with the trimethylated
-ammonium group. Variations in the occupancy of water 2 are
also seen in the different Y245A structures. Water 2 is bound in
similar orientations in the active site of the unmodified and
monomethylated peptide complexes but is absent in the di- and
trimethylated peptide complexes due to the binding of a methyl
group in this position (Fig. 3, A–E). In summary, the changes in
the positions or occupancies of waters 1 and 2 correlate with the
binding modes of the unmodified and methylated K189 within
the active site of the Y245A mutant.
Catalytic Models of Lysine Multiple Methylation by SET7/9
Y245A, and Y305F—The structures of the SET7/9 complexes
reported here offer a prime opportunity to generate stepwise
models for lysine mono-, di-, and trimethylation by a SET
domain KMT. We modeled the AdoMet-bound Michaelis
complexes by superimposing the SET7/9 product complexes
with the previously reported structure of the SET7/9-AdoMet
binary complex (Fig. 4) (39). The conformations of the mono-
and dimethyl -amino groups in the Michaelis complexes were
inferred from the coordinates of the corresponding dimethyl-
and trimethyl-lysine products, respectively. In addition, we
modeled the -amino group in a deprotonated state with its
hydrogen atoms oriented toward the hydrogen bond acceptors
that align the lysyl side chain for methylation. As a basis for this
comparison, we first modeled the monomethylation reaction
catalyzed by WT SET7/9 (Fig. 4A). In the substrate ternary
complex, the lysine -amine is aligned with the methyl group
and sulfonium cation of AdoMet through a hydrogen bond to
the Tyr-245 hydroxyl group and water 1 in the solvent pocket.
The values of the reaction distance and angle are 2.8 Å and 153°,
respectively, in approximate agreement with the linear geome-
try of a SN2 methyl transfer reaction calculated in other mod-
eled substrate complexes (8, 10). In the product complex, the
monomethyl-lysine side chain is bound in an extended confor-
mation with its methyl group oriented within the methyl trans-
fer pore, thereby obstructing AdoMet binding. Furthermore,
water 1 remains tightly coordinated in the solvent pocket
through four hydrogen bonds to Gly-292, Ala-295, Tyr-305,
and the monomethyl -amino group. These interactions hinder
the dissociation of water 1 and the related rearrangement of the
monomethyl-lysine side chain required for a second methyl
transfer reaction, explaining why the WT enzyme cannot cata-
lyze di- and trimethylation. These findings concur with the
FIGURE 4. Catalytic models of the methyl transfer reactions catalyzed by
WT SET7/9 and the Y305F mutant. A, monomethylation of TAF10-K189 by
the WT enzyme. The reaction scheme depicts the modeled substrate ternary
complex (left) and the product complex (right) for the transfer of the methyl
group from AdoMet (green carbon atoms) to K189 in TAF10 (yellow carbons),
yielding AdoHcy and K189me1. The red arrow indicates the direction of the
nucleophilic attack of the deprotonated -amino group on the AdoMet
methyl group. The transferred methyl group is colored green, and the white
atoms represent the hydrogens of the -amino group. Hydrogen bonds
and residues in the enzyme active site are illustrated as in Fig. 2. The
reaction distance and angle are labeled in red. B and C, models of the
Y305F mutant for the first methyl transfer reaction with TAF10-K189 (B)
and second methyl transfer reaction with TAF10-K189me1 (C). Color
schemes are the same as in A.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31855
model for SET7/9 product specificity reported in previous
structural and functional studies (6, 11).
Similar reaction geometry is observed in the model for the
monomethyl transfer reaction catalyzed by SET7/9 Y305F.
Hydrogen bonds from the Tyr-245 hydroxyl group and water 1
align the lysine -amino group with the AdoMet methyl group
at a distance of 2.1 Å and an angle of 160°, equivalent to those
measured in the Michaelis complex of the WT enzyme (Fig. 4, A
and B). In the product complex, monomethyl-lysine adopts an
extended trans configuration analogous to that in the WT
enzyme. For dimethylation to occur, the monomethyl-lysine
must undergo a conformational change in which its methyl
group is rotated out of the methyl transfer path with AdoMet.
The structure of the Y305F mutant bound to the dimethylated
TAF10 peptide (Fig. 2D) implies that this rearrangement occurs
through the dissociation of water 1 due to the loss of the Tyr-
305 hydrogen bond in the solvent pocket. The dissociation of
water 1 would enable the monomethyl-lysine side chain to
adopt an alternative conformation through a rotation about its
C–N bond, projecting the methyl group into the solvent
pocket (Fig. 4C). This rotation reorients the methyl group out of
the methyl transfer path while realigning the monomethyl -
amino group for a second methylation reaction through a direct
hydrogen bond to the Tyr-245 hydroxyl group and a CH–O
hydrogen bond between its methyl group and Ala-295. The
modeled reaction geometry for monomethyl-lysine substrate
complex of the Y305F mutant (2.7 Å, 160°) is equivalent to that
of the first methyl transfer reaction in SET7/9 Y305F. These
geometries concur with our previous models for mono- and
dimethylation catalyzed by SET8 Y334F (8), illustrating that the
orientation of a methyl group into the solvent pocket is a con-
served feature of SET domain KMTs that catalyze multiple
methylation.
In addition, we modeled the methyl transfer reactions cata-
lyzed by SET7/9 Y245A (Fig. 5). In the model of the lysine sub-
strate complex, the -amino group is aligned for methyl transfer
by a hydrogen bond to the carbonyl oxygen of Gly-264, result-
ing in a short reaction distance (2.3 Å) and a suboptimal reac-
tion angle (141°) with the methyl group of AdoMet (Fig. 5A).
This misalignment appears to be a direct consequence of the
Y245A mutation that abolishes hydrogen bonding to the -
amino group, illustrating that the suboptimal orientation of the
-amine likely contributes to the diminished activity of this
mutant toward unmodified substrates (11). Conversely, in the
modeled monomethyl-lysine substrate complex for SET7/9
Y245A, Tyr-305 hydrogen bonds to the -amino group, aligning
it for methyl transfer (Fig. 5B). In addition, CH–O hydrogen
bonds to the monomethyl-lysine methyl group and the dissoci-
ation of water 2 from the active site also contribute to reposition-
ing the -amino group for dimethylation. Collectively, these inter-
actions orient the -amine in a reaction angle of 165° that is more
conducive to methyl transfer. However, the reaction distance for
dimethylation is 0.6 Å longer than that in the corresponding
Y305F model because Tyr-305 is positioned further from
AdoMet than Tyr-245 (Figs. 4C and 5B).
In the third methyl transfer reaction catalyzed by SET7/9
Y245A, the lone pair of electrons of the dimethyl-lysine -
amino group acts as the nucleophile and thus cannot engage in
hydrogen bonding. The structure of the trimethyl-lysine prod-
uct complex (Fig. 3D) implies that the dimethyl -amine is
aligned via CH–O hydrogen bonds to its methyl groups, as
shown in the model of the Michaelis complex for this reaction
(Fig. 5C). These CH–O hydrogen bonds restrain the orienta-
tion of the -amino group and position one of the methyl
groups into the solvent pocket, displacing water 1 as discussed
earlier (Fig. 3, D and E). These interactions cumulatively align
the -amino group and AdoMet methyl group with a reaction
distance of 3.0 Å and angle of 162° (Fig. 5C). Taken together, the
models of the substrate complexes for SET7/9 Y245A suggest
that CH–O hydrogen bonds play an increasingly important role
in aligning the methylated -amino group in successive rounds
of methyl transfer.
DISCUSSION
The structural and functional characterization of the
SET7/9 Y245A and Y305F mutants presented here yields
new insights into the mechanism underlying the product
specificity of SET domain KMTs. Importantly, it resolves a
general paradox concerning this specificity. How does the
active site constrain the motion of the lysine -amino group
to align it for methyl transfer with AdoMet, while providing
adequate volume to accommodate the mono-, di-, and tri-
methylated lysine side chain generated during multiple
methyl transfer reactions? The structures of the Y305F and
Y245A mutants resolve this paradox, illustrating that alter-
ations in the positions or occupancies of water molecules
within their active sites generate the space required to
FIGURE 5. Models for the methyl transfer reactions catalyzed by the
SET7/9 Y245A mutant. Models of the Y245A mutant for the first methyl
transfer reaction with TAF10-K189 (A), the second methyl transfer reaction
with TAF10-K189me1 (B), and the third methyl transfer reaction with TAF10-
K189me2 (C). Residues and hydrogen bonds are depicted as in Fig. 3.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
accommodate the multiply methylated -amine produced
during successive catalytic cycles. Minor perturbations in
the side chains of certain active site residues, such as Tyr-
305, are also observed in alignments of the WT enzyme and
the Y245A and Y305F complexes, although these changes are
modest compared with the displacement or dissociation of
the water molecules in the active site. These findings suggest
that the waters function as transient place holders that facil-
itate the SN2 methyl transfer reaction. During monomethy-
lation, they function to constrain the movement of the lysine
-amino group by mediating hydrogen bonds between the
substrate and enzyme, thereby promoting the linear align-
ment with the methyl group and sulfonium cation of
AdoMet (Fig. 4, A and B). During di- and trimethylation, the
water molecules either relocate within the lysine binding
channel or dissociate from the enzyme, yielding the space
required to rotate the methyl group away from the methyl
transfer pore and to realign the -amine in productive geom-
etry for the next methyl transfer reaction (Figs. 4C and 5, B
and C). These findings agree with our prior analysis of the
SET8 Phe/Tyr switch mutant in which we demonstrated that
the Y334F substitution attenuates hydrogen bonding to the
water molecule bound in the solvent pocket, promoting its
dissociation and the conformational changes necessary for
lysine dimethylation (8). Indeed, there is a nearly identical
alignment of the dimethyl-lysine side chains in the structures of
SET7/9Y305FandSET8Y334Fcomplexes,despitethedifferences
in the orientations of the Phe-305 and Phe-334 side chains in each
structure (Fig. 2F). Finally, both the SET8 Y334F (8) and SET7/9
Y305F mutants (Table 2) displayed diminished catalytic efficien-
cies for lysine dimethylation versus monomethylation. These dif-
ferences may reflect the kinetics of the reorganization within the
active site, including the dissociation of the water molecule from
the solvent pocket and the concomitant realignment of the
monomethyl-lysine into a productive geometry for dimethylation.
In addition to their place-holding role, the active site waters
may also facilitate the deprotonation of the lysine -amino
group between methyl transfer reactions. For methylation to
occur, the -amino group must be deprotonated to function as
the nucleophile in the SN2 methyl transfer reaction with
AdoMet (Figs. 4 and 5). Although the pKa value of the lysine
-amine in solution is 10.5, molecular dynamics simulations
by Zhang and Bruice (25, 26) indicate that this value diminishes
to 8.2 upon formation of the SET7/9 Michaelis complex due to
the proximity of the AdoMet sulfonium cation and the low
dielectric constant of the active site. Furthermore, their simu-
lations show that a chain of water molecules facilitates the dep-
rotonation of the -amino group prior to methyl transfer, trans-
ferring the proton to bulk solvent. Although these water
molecule chains are not evident in our crystal structures, the
Y305F and Y245A complexes suggest another potential mech-
anism for deprotonation. In the dimethyl-lysine complexes of
the Y305F and Y245A mutants, the dissociation of water 1 and
2, respectively, from the lysine binding channel requires that
the solvent-mediated hydrogen bond to the -amino group is
broken (Figs. 2D and 3C). It is conceivable that these waters
dissociate from the active site as hydronium ions, promoting
the realignment and deprotonation of the methyl -amino
group for the next methyl transfer reaction.
A comparison of the SET7/9 Y305F and SET8 Y334F com-
plexes yields insights into the mechanism by which the Phe/Tyr
switch influences water binding within the solvent pocket. The
phenylalanine substitution in the Phe/Tyr switch results in the
loss of a single hydrogen bond to the water molecule (water 1) in
the solvent pocket compared with the four hydrogen bonds that
coordinate the solvent molecule in WT SET7/9 (Fig. 2, A and B)
and SET8 (7, 8). Although this attenuation in hydrogen bonding
may appear insignificant, this difference is nonetheless impor-
tant for at least two reasons. First, theoretical calculations indi-
cate that, on average, water molecules form 3.5 hydrogen
bonds in solutions (40, 41). This value is greater than the num-
ber of hydrogen bonds coordinating water 1 in the solvent
pocket in SET7/9 Y305F (Fig. 2, B and C) as well as in SET8
Y334F and other di- and trimethyltransferases that possess a
hydrophobic residue in the Phe/Tyr switch site (8). From the
perspective of the water molecule, the greater hydrogen bond-
ing potential in solution would tend to thermodynamically
favor its dissociation from the solvent pocket in SET domain
KMTs that lack a tyrosine in the Phe/Tyr switch position. Sec-
ond, the ordered binding of water molecules observed in the
active sites of SET domain ternary complexes represents an
unfavorable entropy compared with their diffusion in bulk sol-
vent. In WT SET7/9 (Fig. 2A) and SET8 (7, 8), this entropic
penalty can be partially offset through the favorable enthalpy of
binding associated with the four hydrogen bonds that coordi-
nate the water within the solvent pocket. It is conceivable that
the loss of the tyrosine-mediated hydrogen bond in the Phe/Tyr
switch shifts the equilibrium in favor of dissociation of the
water molecule from the solvent pocket, thereby facilitating
dimethylation in SET7/9 Y305F, SET8 Y334F, and other di- and
trimethyltransferases.
The structures of the SET7/9 Y245A and Y305F complexes
illustrate the interactions that align the lysine -amino group
during the methyl transfer reactions in each enzyme. In the WT
enzyme and the Y305F mutant, hydrogen bonding to the
hydroxyl group of Tyr-245 appears to be critical in properly
aligning the -amine for methyl transfer (Fig. 4). Tyr-245 is
conserved in the sequences of many SET domain KMTs (8, 42),
and substitutions of this residue generally impair or abolish
activity, indicating its importance in catalysis (8, 43). However,
SET7/9 appears to be an exception to this rule, as the Y245A
mutant is not only active but is capable of catalyzing lysine
trimethylation. In this mutant, Tyr-305 appears to assume the
role of Tyr-245 by hydrogen bonding to the monomethylated
-amino group to align it for methyl transfer with AdoMet, as
illustrated in the modeled substrate complex for the dimethy-
lation reaction (Fig. 5B). Conversely, in the model for trimethy-
lation, the Tyr-305 hydroxyl group does not hydrogen bond to
the -amine but instead participates in a CH–O hydrogen bond
with one of the methyl groups to assist in aligning the dimethy-
lated -amine for the methyl transfer reaction (Fig. 5C). Addi-
tional structural and functional studies of the SET domain tri-
methyltransferases will aid in further illuminating the roles
of CH–O hydrogen bonds in facilitating lysine multiple
methylation.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31857
Acknowledgments—We acknowledge S. Schiebold for assistance in
protein expression, purification, and crystallization and S. Anderson
and R. Sanishvili for their assistance with x-ray data collection. We
also thank S. Bulfer and S. Horowitz for reading the manuscript and
providing useful comments. This work utilized the Protein Structure
Facility of the Michigan Diabetes Research and Training Center, Uni-
versity of Michigan, supported by National Institutes of Health Grant
DK020572, NIDDK. Use of the Advanced Photon Source was sup-
ported by the United States Department of Energy, Basic Energy Sci-
ences, Office of Science, under Contract DE-AC02-06CH11357.
GM/CA CAT has been funded in whole or in part by National
Institutes of Health NCI Grant Y1-CO-1020 and NIGMS Grant
Y1-GM-1104. Use of the LS-CAT Sector 21 was supported by Michi-
gan Economic Development Corporation and the Michigan Technol-
ogy Tri-Corridor Grant 085P1000817 for the support of this research
program.
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Lysine Methylation by SET7/9 Mutants
31858
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
|
3M59
|
SET7/9 Y245A in complex with TAF10-K189me2 peptide and AdoHcy
|
SET7/9 Catalytic Mutants Reveal the Role of Active Site Water
Molecules in Lysine Multiple Methylation*□
S
Received for publication,February 15, 2010, and in revised form, July 14, 2010 Published, JBC Papers in Press,August 1, 2010, DOI 10.1074/jbc.M110.114587
Paul A. Del Rizzo‡, Jean-Franc¸ois Couture‡§1, Lynnette M. A. Dirk¶, Bethany S. Strunk‡, Marijo S. Roiko‡,
Joseph S. Brunzelle, Robert L. Houtz¶, and Raymond C. Trievel‡2
From the ‡Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109, the §Ottawa Institute of
Systems Biology, Department of Biochemistry, Microbiology, and Immunology, University of Ottawa, Ottawa, Ontario K1H 8M5,
Canada, the ¶Department of Horticulture, Plant Physiology/Biochemistry/Molecular Biology Program, University of Kentucky,
Lexington, Kentucky 40546, and the Department of Molecular Pharmacology and Biological Chemistry, Northwestern University
Center for Synchrotron Research, Life Sciences Collaborative Access Team, Argonne, Illinois 60439
SET domain lysine methyltransferases (KMTs) methylate
specific lysine residues in histone and non-histone substrates.
These enzymes also display product specificity by catalyzing dis-
tinct degrees of methylation of the lysine -amino group. To
elucidate the molecular mechanism underlying this specificity,
we have characterized the Y245A and Y305F mutants of the
human KMT SET7/9 (also known as KMT7) that alter its prod-
uct specificity from a monomethyltransferase to a di- and a tri-
methyltransferase, respectively. Crystal structures of these
mutants in complex with peptides bearing unmodified, mono-,
di-, and trimethylated lysines illustrate the roles of active site
water molecules in aligning the lysine -amino group for methyl
transfer with S-adenosylmethionine. Displacement or dissocia-
tion of these solvent molecules enlarges the diameter of the
active site, accommodating the increasing size of the methylated
-amino group during successive methyl transfer reactions.
Together, these results furnish new insights into the roles of
active site water molecules in modulating lysine multiple meth-
ylation by SET domain KMTs and provide the first molecular
snapshots of the mono-, di-, and trimethyl transfer reactions
catalyzed by these enzymes.
SET domain enzymes represent a family of S-adenosylmethi-
onine (AdoMet)3-dependent methyltransferases that catalyze
the site-specific methylation of protein lysyl residues in a host
of proteins, including histones, transcription factors, chroma-
tin-modifying enzymes, ribosomal subunits, and other sub-
strates (1–3). In many instances, these modifications serve to
recruit effector proteins that recognize methyl-lysyl residues in
a sequence-dependent fashion (4). In addition, SET domain
KMTs exhibit product specificity, defined as their ability to cat-
alyze mono-, di-, or trimethylation of the lysine -amino group.
This specificity is biologically relevant because many methyl-
lysine-binding proteins can discriminate among different
degrees of lysine methylation (4). Thus, both the site and degree
of lysine methylation are critical to recognition by effector
proteins.
Structural and functional studies have identified a Phe/Tyr
switch in the active site of SET domain KMTs that governs their
respective product specificities (5, 6). According to this model,
KMTs that possess a tyrosine in the Phe/Tyr switch site are
limited to catalyzing lysine monomethylation, whereas en-
zymes that possess a phenylalanine or another hydrophobic
residue in this position display di- or trimethyltransferase activ-
ity. Mutational analysis of various SET domain KMTs, includ-
ing DIM-5 (KMT1) (6), SET7/9 (6), G9A (KMT1C) (5), and
SET8 (also known as PR-SET7 and KMT5A) (7, 8), has demon-
strated that substitutions in the Phe/Tyr switch result in pre-
dictable changes in product specificity. Several models have
been proposed to explain the mechanism by which the Phe/Tyr
switch site governs this specificity, including variations in the
diameter of the active site due to the size of Phe/Tyr switch
residue and steric hindrance by the tyrosine hydroxyl group (6,
9–11). However, our recent studies of the Phe/Tyr switch
mutant Y334F in the human histone H4 Lys-20 (H4K20) meth-
yltransferase SET8 indicate that the Phe/Tyr switch regulates
product specificity via a more subtle mechanism (8). Specifi-
cally, the switch modulates the binding of an active site water
molecule that in turn regulates the transition from mono-
methylation to multiple methylation.
Among the KMTs that have been structurally characterized,
SET7/9 has emerged as an archetypal model for studying the
catalytic mechanism and product specificity of the SET domain
family. Although initially isolated as a histone H3 Lys-4
(H3K4)-specific methyltransferase, this KMT has been shown
to regulate the functions of numerous non-histone substrates
through site-specific methylation (12–21). Early structural and
functional studies of SET7/9 identified two active site mutants,
* Thisworkwassupported,inwholeorinpart,byNationalInstitutesofHealth
Grant R01 GM073839 (to R. C. T.) and National Institutes of Health Admin-
istrative Supplement GM073839-04S1 (to R. C. T.) funded through the
American Recovery and Reinvestment Act. This work was also supported
by Department of Energy Grant DE-FG02-92ER20075 (to R. L. H.).
□
S The on-line version of this article (available at http://www.jbc.org) contains
supplemental Table 1.
The atomic coordinates and structure factors (codes 3M53, 3M54, 3M55, 3M56,
3M57,3M58,3M59,and3M5A)havebeendepositedintheProteinDataBank,
Research Collaboratory for Structural Bioinformatics, Rutgers University, New
Brunswick, NJ (http://www.rcsb.org/).
1 Supported by a Canadian Institutes of Health Research postdoctoral
fellowship.
2 To whom correspondence should be addressed: Dept. of Biological Chem-
istry, University of Michigan Medical School, 1150 West Medical Center Dr.,
5301 MSRB III, Ann Arbor, MI 48109. Tel.: 734-647-0889; Fax: 734-763-4581;
E-mail: rtrievel@umich.edu.
3 The abbreviations used are: AdoMet, S-adenosylmethionine; AdoHcy, S-ad-
enosylhomocysteine; KMT, lysine methyltransferase; ITC, isothermal titra-
tion calorimetry; CH–O, carbon-oxygen hydrogen bond; BisTris, 2-[bis(2-
hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 41, pp. 31849–31858, October 8, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31849
Y245A and Y305F, which change its product specificity. The
Phe/Tyr switch mutant Y305F alters SET7/9 product specificity
from a mono- to dimethyltransferase (6), whereas the Y245A
substitution converts the enzyme into a trimethyltransferase
with weak monomethyltransferase activity (11). These mutants
have been the subjects of numerous molecular modeling simu-
lations that have led to various models to explain their distinct
product specificities (22–26). However, the lack of structural
data for the SET7/9 Y245A and Y305F mutants in complex with
cognate methylated peptides has hindered our understanding
of the mechanisms that define the respective product specific-
ities of these mutants. Moreover, these structures would yield a
framework for visualizing the mono-, di-, and trimethylation
reactions catalyzed by SET domain KMTs.
To gain insight into the molecular basis of their product
specificities, we have determined high resolution crystal
structures of the SET7/9 Y245A and Y305F mutants in com-
plex with peptides of the TATA box-binding protein-associ-
ated factor TAF10 bearing the Lys-189 methylation site in
unmodified (K189), monomethylated (K189me1), dimethyl-
ated (K189me2), and trimethylated (K189me3) states. The
structures and accompanying biochemical data support a
model whereby changes in the occupancy or position of water
molecules in the active site are critical in establishing the prod-
uct specificities of the SET7/9 Y245A and Y305F mutants.
Together, our results provide new insights into the mechanisms
that govern SET domain product specificity and provide step-
wise snapshots of the lysine mono-, di-, and trimethyl transfer
reactions catalyzed by KMTs.
EXPERIMENTAL PROCEDURES
Cloning, Expression, and Purification of the SET7/9 Mutants—
The Y245A and Y305F mutants were introduced into the pHIS2
SET7/9 expression vector encoding residues 110–366 (27) via
QuikChange site-directed mutagenesis (Stratagene) and were
verified by dideoxy DNA sequencing. The plasmids encoding
wild type (WT) SET7/9 and the Y245A and Y305F mutants
were transformed into Rosetta2 DE3 cells (Novagen) and were
expressed as described previously (27, 28). In the course of
characterizing WT SET7/9, we observed that the enzyme co-
purified with AdoMet or another contaminant that resulted in
technical difficulties in the isothermal titration calorimetry
(ITC) experiments and co-crystallization trials with the TAF10
peptides. To overcome this problem, a denaturation and refold-
ing step was inserted in the purification scheme. The denatur-
ation and refolding protocol involved adding 6 M guanidine
HCl, 50 mM sodium phosphate, pH 7.5, and 500 mM NaCl to the
protein while it was immobilized on a nickel-Sepharose column
(GE Healthcare). The column was washed with this buffer, fol-
lowed by a solution of 6 M urea, 50 mM sodium phosphate, pH
7.5, 150 mM NaCl, and 10 mM 2-mercaptoethanol to remove the
cofactor from the denatured enzyme. A reverse gradient from 6
to 0 M urea was then performed in the same buffer to refold the
protein, which was subsequently eluted from the column using
a linear gradient of 0–500 mM imidazole in 50 mM sodium
phosphate, pH 7.5, 500 NaCl, and 10 mM 2-mercaptoethanol.
The refolded protein was digested with tobacco etch virus pro-
tease (29) during dialysis against 20 mM Tris, pH 7.5, 150 mM
NaCl, and 5 mM 2-mercaptoethanol and then purified using a
Superdex 200 gel filtration column (GE Healthcare). Protein
concentration was determined by its absorbance at 280 nm.
Synthetic Peptides—The TAF10 peptides bearing K189,
K189me1,
K189me2,
and
K189me3
(sequence,
acetyl-
SKSK189DRKYTL-amide) and a biotinylated TAF10 peptide
(sequence,
acetyl-SKSK189DRKYTLT(K-EZLink-S-S-biotin)-
amide) were synthesized and purified by New England Peptide,
Inc. Peptide concentrations were measured using the absorb-
ance of their tyrosine residue at 274 nm.
Crystallization and Data Collection—Crystals were pro-
duced by hanging drop vapor diffusion by mixing the crystalli-
zation solution in a 1:1 ratio with 10–20 mg/ml SET7/9, 1 mM
S-adenosylhomocysteine (AdoHcy), and 1.0–3.0 mM unmodi-
fied or methylated TAF10-K189 peptide in 20 mM Tris, pH 8.0,
100 mM NaCl, and 2 mM tris(2-carboxyethyl)phosphine. Crys-
tals were obtained at 20 °C in either 1.8–2.0 M (NH4)2SO4, with
0.1 M BisTris, pH 6.2–6.6, or in 0.95–1.1 M sodium citrate with
100 mM imidazole pH 8.0–8.4. In both crystallization condi-
tions, the final pH values were between pH 8.0 and 9.0. Crystals
in the (NH4)2SO4 condition were typically flash-frozen in the
mother liquor containing 25–30% glycerol, and the crystals in
the citrate condition were frozen in 1.6 M sodium citrate. Data
were collected at the Advanced Photon Source beamlines
21-IDG (LS-CAT) and 23-IDD (GM/CA-CAT). Images were
indexed, integrated, and scaled using HKL2000 (30). Structures
of the mutants were solved by molecular replacement using
MOLREP (31) with the coordinates of a previously reported
SET7/9 ternary complex used as the search model (Protein
Data Bank code 2F69). Successive rounds of model building and
refinement were carried out using Coot (32) and REFMAC (33),
respectively. The geometry of the models were verified by Mol-
Probity (34). Simulated annealing omit maps were calculated
using CNS (35) with the peptide and cofactor removed to elim-
inate model bias in the active site. Structural figures were ren-
dered using PyMOL (Schro¨dinger, LLC.).
Fluorescent Methyltransferase Assay—A coupled fluorescent
methyltransferase assay was used to measure the kinetic
parameters of WT SET7/9 and the Y245A and Y305F mutants
as reported previously, with the exception that 50–150 nM
enzyme, 100 M AdoMet, and varying concentrations of TAF10
peptide substrate were used (27, 36). Assays were performed in
triplicate, and a homocysteine calibration curve was used to
calculate the initial velocities. Kinetic parameters were calcu-
lated by plotting the velocities versus peptide concentration and
by fitting the Michaelis-Menten equation to the data via non-
linear regression using Prism 5.0 (GraphPad). In cases where
the Km value was beyond the measurable range of the assay, the
kcat/Km value was determined as described previously (7).
Calorimetry Experiments—ITC was performed at 20 °C using
a MicroCal VP-ITC calorimeter (GE Healthcare) with 0.12 mM
protein and 4 mM AdoHcy in 20 mM sodium phosphate, pH 7,
and 100 mM NaCl with 1.5 mM peptide as the injectant. Data
were processed, and equilibrium dissociation constants (KD)
and curve fitting errors were calculated from the binding iso-
therms using Origin 7.0 (OriginLab Corp.). WT SET7/9 and the
Y245A and Y305F mutants displayed ligand:protein binding
stoichiometries (N values) between 0.8 and 1.0, demonstrating
Lysine Methylation by SET7/9 Mutants
31850
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
that WT SET7/9 and its mutants were properly refolded due to
their ability to bind peptides in an 1:1 molar ratio.
TLC Product Analysis—Methyltransferase assays were per-
formed in triplicate at 37 °C with the biotinylated TAF10 pep-
tide (0.5 mM) for 30 min in the presence of WT SET7/9 (3
pmol), Y305F mutant (6 pmol), or the Y245A mutant (100
pmol). Assays contained 100 mM HEPES, pH 8.0, 50 mM NaCl,
1.0 mM [methyl-3H]AdoMet (diluted to a specific activity of 0.2
Ci/nmol with purified AdoMet (37)), 15 M Sulfolobus solfa-
taricus AdoHcy hydrolase (36), and 2 units of adenosine deami-
nase (Roche Applied Science) in a final volume of 20 l. The
reactions were terminated by addition of an equal volume of
200 mM MES, pH 5.1, prior to addition of a 2-fold molar excess
of immobilized avidin resin (UltraLink; Pierce). Biotinylated
peptides were allowed to bind at room temperature for 30 min,
and the resin was then collected by centrifugation (9000 g).
The resin was washed three times with 300 mM NaCl, and the
peptide was eluted overnight from the avidin resin by cleavage
of the disulfide bond in the linker of the peptide using 10 mM
tris(2-carboxyethyl)phosphine (Thermo Scientific). The resin
was incubated with additional 10 mM tris(2-carboxyethyl)phos-
phine the following day until the radiolabel was essentially
removed from the resin. The recovered peptides were hydro-
lyzed by incubation with 6 M HCl at 110 °C for 24 h. Subsequent
steps in measuring the radiolabel incorporated into the mono-,
di-, and trimethyl-lysine products were performed as reported
previously (8).
RESULTS
Functional Analysis of the SET7/9
Y305F Mutant—Prior studies of
SET7/9 by Zhang et al. (6) reported
that mutation of the Phe/Tyr switch
residue Tyr-305 to a phenylalanine
alters its product specificity from a
mono- to dimethyltransferase. We
verified these findings by demon-
strating that WT SET7/9 mono-
methylated the TAF10-K189 pep-
tide, whereas the Y305F mutant
mono- and dimethylated this sub-
strate, as demonstrated by mass
spectrometry (data not shown). We
next examined whether the Y305F
substitution altered the affinity of SET7/9 for the TAF10-K189
peptides using ITC (Fig. 1). A comparison of the KD values
revealed that SET7/9 Y305F bound the TAF10-K189 and
TAF10-K189me1 peptides 4- and 6-fold more tightly, respec-
tively, than the WT enzyme, whereas this mutant displayed a
substantially diminished affinity for the TAF10-K189me2 pep-
tide (Table 1). Although the WT enzyme and the Y305F mutant
exhibited discernable differences in their affinities for the
unmodified and monomethylated peptides, these variations are
modest and cannot account for their distinct product specific-
ities, suggesting that a kinetic effect during methylation may be
responsible.
To investigate this possibility, we characterized the kinetic
parameters of WT SET7/9 and the Y305F mutant using the
TAF10 peptides as substrates. Both enzymes methylated the
unmodified peptide with comparable kcat and Km values (Table
2). In analyzing the kinetic parameters for the methylation
of the monomethylated peptide by SET7/9 Y305F, we found
that this substrate displayed an elevated Km value that was
beyond the measurable range of the assay due to its limited
solubility. In this case, we measured the catalytic efficiency
(kcat/Km) for the methylation of this peptide and found that it
was methylated 15-fold less efficiently than the unmodified
peptide by SET7/9 Y305F. Given the fact that the Y305F mutant
exhibited a higher binding affinity for the TAF10-K189me1
peptide than the WT enzyme (Table 1), the kinetic data suggest
that a step in the reaction pathway following substrate binding
limits the catalytic efficiency of this mutant.
We next examined whether the Y305F mutant dimethylated
the TAF10-K189 peptide via a processive or a distributive
mechanism. In a processive mechanism, the methyl-lysine
substrate would remain bound to the enzyme during successive
methyl transfer reactions; thus, the concentration of an inter-
mediate, such as monomethyl-lysine, cannot exceed the en-
zyme concentration during the assay. In a distributive mecha-
nism, the intermediates are released into solution where they
accumulate prior to the next round of methylation, resulting in
an intermediate concentration that is greater than that of the
enzyme. Using a radiometric TLC assay and a biotinylated
TAF10 peptide, we quantified the amounts of monomethylated
products generated by the WT SET7/9 and the Y305F mutant
FIGURE 1. ITC analysis of WT SET7/9 and the Y245A and Y305F mutants. Representative titrations and
binding isotherms for the calorimetry experiments are shown for the unmodified TAF10-K189 peptide titrated
into WT SET7/9 (left plot), SET7/9 Y305F (middle plot), and SET7/9 Y245A (right plot). In each panel, the ITC
titration experiment (upper panel) is shown with the binding isotherms (lower panel) fit to a single binding site
model.
TABLE 1
Analysis of the binding affinity of WT SET7/9 and its catalytic
mutants for unmodified and methylated TAF10 peptides
SET7/9
TAF10 peptide
KD
a
M
WT
K189
4.9 0.20
WT
K189me1
4.0 0.36
Y305F
K189
1.3 0.10
Y305F
K189me1
0.62 0.065
Y305Fb
K189me2
70
Y245A
K189
4.0 0.25
Y245A
K189me1
3.3 0.10
Y245A
K189me2
5.8 0.22
Y245A
K189me3
11 0.28
a Curve fitting errors were calculated from the binding isotherms.
b An estimate of the affinity is reported due to weak peptide binding.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31851
(Table 3). The data demonstrate that comparable amounts of
monomethyl-lysine were generated when the quantity of
enzyme usedistakenintoaccount,inagreementwiththeirsimilar
turnover numbers for the TAF10-K189 peptide (Table 2). The
Y305F mutant also produced small but measurable quantities of
radiolabeled dimethyl-lysine product that were substantially
smaller than the amount of monomethyl-lysine generated.
Therefore, the TLC data are consistent with a distributive
mechanism for dimethylation by the Y305F mutant because the
amount of monomethyl-lysine produced exceeded the quantity
of enzyme used in the assay.
Structures of WT SET7/9 and the Y305F Mutant in Complex
with Unmodified and Methylated TAF10 Peptides—To deter-
mine the mechanism by which the Y305F substitution alters the
product specificity of SET7/9, we determined the crystal struc-
tures of this mutant bound to AdoHcy and TAF10-K189,
TAF10-K189me1, and TAF10-K189me2 peptides and com-
pared these to the structures of the WT SET7/9AdoHcy
TAF10-K189 complex (supplemental Table 1). The structures
of these complexes were determined to 1.85 Å or higher reso-
lution, permitting unambiguous modeling of the K189 side
chains in the active site of the enzyme based on simulated
annealing omit maps (Fig. 2). The ternary complexes of the WT
and the Y305F mutant superimpose with overall root mean
square differences of less than 0.3 Å for all aligned atoms, indi-
cating that neither the Y305F mutation nor the binding of the
various TAF10-K189 peptides results in substantial changes in
its overall structure.
An inspection of the active sites of the SET7/9 WT and
Y305F complexes illustrates the binding modes of the unmod-
ified and methylated forms of K189 in the TAF10 peptides (Fig.
2, A–D). The K189 side chain binds in an extended all trans
conformation in a deep pocket, termed the lysine binding chan-
nel, that is composed of Tyr-245, Thr-266, Leu-267, Ser-268,
Tyr-305, Tyr-335, and Tyr-337 (not shown for clarity) (Fig. 2A).
These residues interact with the aliphatic portion of the K189
side chain primarily through van der Waals contacts. The lysine
binding channel connects to the AdoMet-binding site on the
opposite face of the catalytic domain via an oxygen-lined
methyl transfer pore (38). During catalysis, the methyl group of
the cofactor is positioned within the methyl transfer pore
for the SN2 reaction with the -amino group of the lysine or
methyl-lysine substrate (see below).
To lower the activation barrier for this reaction, the lysine
-amine nucleophile is aligned for methyl transfer through a
hydrogen bond network within the active site. In the WT
enzyme, the K189 -amino group hydrogen bonds to the
hydroxyl group of Tyr-245 as well as to two water molecules
(Fig. 2A). One of the water molecules (termed water 1), is coor-
dinated in a solvent pocket, through hydrogen bonds to the
carbonyl oxygens of Gly-292 and Ala-295 and to the hydroxyl
group of the Phe/Tyr switch residue Tyr-305. This solvent
pocket is structurally conserved in SET domain KMTs and has
an important role in defining product specificity through the
adjacent Phe/Tyr switch residue, as shown in our prior studies
of the human H4K20 methyltransferase SET8 (8). The other
water molecule is bound within the methyl transfer pore
between the lysine substrate and the thioether sulfur atom of
AdoHcy through hydrogen bonds to Tyr-245, Asn-265, and
His-293 in SET7/9 and the TAF10-K189 -amino group. This
water is not observed in other structures of SET7/9 ternary
complexes and may represent the approximate position that
the AdoMet methyl group occupies in the methyl transfer pore
in the Michaelis complex.
In structures of the Y305F ternary complexes, the K189,
K189me1, and K189me2 side chains also adopt extended trans
side chain geometries within the lysine binding channel that are
stabilized via hydrogen bonding to Tyr-245 in the enzyme (Fig.
2, B–D). The orientations of the K189me1 and K189me2 side
chains are further maintained through carbon-oxygen (CH–O)
hydrogen bonding between the methyl groups and oxygen
atoms within the vicinity of the methyl transfer pore, as
reported previously in other SET domain KMT structures (8,
10, 38). A superimposition of the SET7/9 WT and Y305F com-
plexes underscores the similarity of the lysyl binding conforma-
tions (Fig. 2E). However, there are notable differences in the
hydrogen bond patterns and occupancy of water 1 within the
solvent pocket in the Y305F mutant compared with the WT
enzyme. Specifically, the Y305F substitution results in the loss
of one hydrogen bond to water 1 in the structures of the TAF10-
K189 and TAF10-K189me1 complexes (Fig. 2, B and C). In con-
trast, water 1 is absent in TAF10-K189me2 complex, and the
vacated solvent pocket is occupied by one of the methyl groups
TABLE 2
Kinetic parameters of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
TAF10 peptide substrate
Km
a
kcat
a
kcat/Km
a
M
min1
M1 min1 103
WT
K189
160 17
17 0.62
110 17
Y305F
K189
88 5.0
17 0.30
190 11
Y305Fb
K189me1
11 0.50
Y245A
K189
200 35
0.53 0.04
2.6 0.47
Y245A
K189me1
210 23
5.9 0.23
28 3.3
Y245A
K189me2
400 29
6.5 0.16
15 1.2
a Curve fitting errors were calculated from the hyperbolic fits for the Michaelis-Menten equation.
b Km and kcat were not determined because the Km value was beyond the measurable range; therefore, the kcat/Km value is reported.
TABLE 3
Product analysis of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
Quantity
of enzyme
Measured
product
Amount of
product formeda
nmol
nmol
WT
0.003
Kme1
0.65 0.07
Y305F
0.006
Kme1
1.5 0.49
Kme2
0.033 0.009
Y245A
0.100
Kme1
0.80 0.22
Kme2
0.39 0.021
Kme3
0.076 0.019
a Standard deviation was calculated from triplicate measurements.
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of the dimethyl -amine (Fig. 2D). This methyl group forms a
3.5-Å CH–O hydrogen bond to the carbonyl oxygen of Ala-295,
further stabilizing the binding of the dimethyl-lysine side chain.
A homologous dimethyl-lysine-binding mode and CH–O
hydrogen bond was observed in our prior structural studies of
the SET8 Y334F Phe/Tyr switch mutant that confers an analo-
gous change in product specificity from a mono- to a dimeth-
yltransferase (8). A structural alignment of the active sites of the
SET7/9 Y305F and SET8 Y334F mutants bound to cognate
dimethylated peptides illustrates that the coordinates of the
dimethyl-lysyl side chains are virtually superimposable, with
one methyl group oriented toward the methyltransfer pore and
the second positioned within the vacant solvent pocket (Fig.
2F). Taken together, the structures of the SET7/9 Y305F com-
plexes and the similarities in the dimethyl-lysine conforma-
tions in the SET7/9 Y305F and SET8 Y334F mutants imply that
the Phe/Tyr switch governs product specificity through a con-
served mechanism whereby it indirectly influences the binding
modes of the methyl-lysine side chain by modulating the affin-
ity of the water molecule (water 1) bound in the solvent pocket.
Biochemical Characterization of
the SET7/9 Y245A Mutant—Previ-
ous studies by Xiao et al. (11)
reported that the Y245A mutation
yields an unusual change in the
product
specificity
of
SET7/9,
converting the enzyme to a trimeth-
yltransferase with weak monometh-
yltransferase activity. We deter-
mined that the SET7/9 Y245A could
mono-, di-, and trimethylate the
TAF10-K189 peptide by mass spec-
trometry (data not shown) and TLC
(Table 3), confirming the earlier
studies of Xiao et al. (11). ITC
analysis revealed that the Y245A
mutant displayed comparable KD
values
for
the
unmodified
and
methylated TAF10-K189 peptides
(Fig. 1), although its affinity for the
trimethylated peptide was modestly
diminished in comparison with the
other peptides (Table 1). The ITC
data demonstrate that the Y245A
mutant
bound
the
unmodified,
mono-,
and
dimethylated
sub-
strates with equivalent affinities,
suggesting that a kinetic effect or
a structural alteration in the active
site may be responsible for its
diminished
activity
toward
un-
modified substrates.
To gain further insight into its
peculiar
product
specificity,
we
characterized the kinetic properties
of
the
SET7/9
Y245A
mutant.
Steady state analysis demonstrated
that this mutant displayed similar
Km values for the unmodified, mono- and dimethylated TAF10
peptides (Table 2). However, the turnover number for the
TAF10-K189 peptide was diminished over 10-fold versus the
methylated peptides and was reduced 30-fold versus the WT
enzyme, in agreement with the weak monomethyltransferase
activity reported by Xiao et al. (11). In addition, we investigated
whether this mutant catalyzes lysine trimethylation via a pro-
cessive or distributive mechanism as described for SET7/9
Y305F. The TLC data illustrate that the mono- and dimethyl-
lysine intermediates accumulated at quantities greater than
that of the enzyme used in the assay, indicating that SET7/9
Y245A obeys a distributive mechanism, analogous to the Y305F
mutant (Table 3).
Structures of SET7/9 Y245A Bound to Unmodified and Meth-
ylated TAF10 Peptides—To elucidate the mechanism underly-
ing its unusual product specificity, we determined the crystal
structures of SET7/9 Y245A in complex with AdoHcy and
unmodified, mono-, di-, and trimethylated TAF10 peptides
(supplemental Table 1). These complexes superimpose with
the structure of the WT SET7/9AdoHcyTAF10-K189 com-
FIGURE 2. Crystal structures of WT SET7/9 and the Y305F mutant in complex with AdoHcy and unmodi-
fied and methylated TAF10 peptides. A, active site of WT SET7/9 bound to AdoHcy and TAF10-K189 and the
active site of SET7/9 Y305F bound to AdoHcy and TAF10-K189 (B), TAF10-K189me1 (C), and TAF10-K189me2
peptides (D). A–D, the K189 residue in the TAF10 peptide is depicted with yellow carbon atoms with the
corresponding Fo – Fc simulated-annealing omit maps (contoured at 2.5 ) shown. AdoHcy is colored with
green carbon atoms, and SET7/9 carbon atoms are colored gray, with the exception of Tyr-305/Y305F
(magenta) and Tyr-245 (cyan). Gly-264 to Ser-268 are shown, and the main chain atoms of Gly-292 to Ala-295
are depicted for clarity. Red dashed lines indicate conventional hydrogen bonds, and blue dashed lines indicate
CH–O hydrogen bonds. Cyan dashed lines indicate weak CH–O hydrogen bonds between 3.5 and 3.7 Å in
length. The water molecule in the solvent binding pocket is numbered 1. E, overlay of the active sites of the WT
enzyme (light blue carbon atoms) and the Y305F mutant (green, yellow, and pink carbon atoms correspond to
K189, K189me1, and K189me2, respectively). Red dashed lines indicate coordination of the waters in the struc-
turesoftheY305Fmutant,andthelightbluedashedlineindicateshydrogenbondingthatonlyoccursintheWT
enzyme. F, superimposition of the active sites of the SET7/9 Y305FAdoHcyTAF10-K189me2 and SET8
Y334FAdoHcyH4K20me2 (Protein Data Bank code 3F9X) complexes shown with green and yellow carbon
atoms, respectively.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31853
plex with root mean squared differences of less than 0.4 Å for all
aligned atoms, indicating that the Y245A mutant does not per-
turb the overall structure of the enzyme. Simulated annealing
omit maps illustrate that K189 side chains are bound within the
lysine binding channel through hydrogen bonds and van der
Waals contacts (Fig. 3, A–D), although the interactions and
binding modes are distinct from those in the complexes of WT
SET7/9 and the Y305F mutant (Fig. 2, A–D). In the unmodified
TAF10 peptide complex, the K189 -amino group forms a weak
hydrogen bond with the carbonyl oxygen of Gly-264 (Fig. 3A),
whereas the -amino groups of K189me1 and K189me2 hydro-
gen bond to the hydroxyl group of Tyr-305 in the mono- and
dimethylated peptide complexes (Fig. 3, B and C). The confor-
mations of the K189me1 and K189me2 side chains are further
stabilized by water-mediated hydrogen bonding and through
CH–O hydrogen bonding to their methyl groups. In the
TAF10-K189me3 peptide complex, the trimethyl-lysine side
chain is coordinated exclusively through direct and water-me-
diated CH–O hydrogen bonds to its methyl groups because the
quaternary -ammonium cation cannot engage in hydrogen
bonding (Fig. 3D).
A structural alignment of the four SET7/9 Y245A complexes
illustrates distinct binding modes for the unmodified versus the
methylated K189 side chains, highlighting the selectivity of this
mutant for methylated substrates. The side chains of K189me1,
K189me2, and K189me3 roughly overlay with their respective
-amino groups superimposed and adopt slightly kinked con-
formations (Fig. 3E), as opposed to the extended trans geome-
try of the unmodified and methylated lysines in the complexes
of the WT enzyme and the Y305F mutant (Fig. 2, A–D). Con-
versely, the unmodified K189 side chain does not superimpose
with its methylated counterparts and is oriented in an alterna-
tive configuration due to its hydrogen bonding to Gly-264 (Fig.
3, A and E). An overlay of the structures of the WT enzyme and
Y245A mutant bound to the unmodified TAF10 peptide illus-
trates that the side chains of K189 do not superimpose and that
the K189 -amino group appears to be misaligned with AdoHcy
in the Y245A complex (Fig. 3F). This suboptimal alignment
may explain the diminished kcat value of SET7/9 Y245A mutant
toward substrates with unmodified lysines (Table 2).
A comparison of the structures of the SET7/9 Y245A and
Y305F complexes yields a molecular explanation for the differ-
ent product specificities of these two mutants. In the SET7/9
Y305F complexes, Tyr-245 aligns the K189 -amino group for
methyl transfer through hydrogen bonding to its hydroxyl
group (Fig. 2, B–D). Conversely, in the Y245A mutant, the
K189me1 and K189me2 -amino groups are oriented through
hydrogen bonding to Tyr-305 (Fig. 3, B and C). These distinct
hydrogen bond patterns impart differences in the conforma-
tions of the lysyl side chains due to the relative orientations of
Tyr-245 and Tyr-305 in the lysine binding channel. Specifically,
the kinked conformation adopted by the K189me1 and
K189me2 side chains in the Y245A complexes (Fig. 3, B and C)
may contribute to the differences in the turnover numbers of
this mutant versus those of the WT enzyme and the Y305F
mutant (Table 2). In addition, the dimethyl -amino group of
the K189me2 side chain binds in distinct orientations in the
Y245A and Y305F mutants due to their hydrogen bonding to
FIGURE 3. Crystal structures of the SET7/9 Y245A mutant in complex with
AdoHcy and unmodified and methylated TAF10 peptides. Active site of
SET7/9 Y245A bound to AdoHcy and TAF10-K189 (A), TAF10-K189me1 (B),
TAF10-K189me2 (C), and TAF10-K189me3 peptides (D). A–D, Fo – Fc simulated-
annealing omit maps (contoured at 2.5 ) for the unmodified and methylated
K189 side chains are illustrated. The residues and hydrogen bonds in each com-
plexarecoloredasinFig.2.Thewatermoleculesinthelysinebindingchannelsof
the Y245A complexes are numbered 1–4, as described in the text. E, superimpo-
sition of the active sites of the Y245A complexes bound to the four methylated
statesofTAF10-K189(unmodified,mono-,di-,andtrimethylatedshowningreen,
yellow, pink, and blue, respectively). F, overlay of WT SET7/9 (blue carbons) and
SET7/9 Y245A (green carbons) in complex with TAF10-K189. Waters correspond-
ing to the WT and Y245A structures are colored cyan and green, respectively.
G, alignment of the active sites of the Y245A (yellow carbons) and Y305F (green
carbons) mutants in complex with the TAF10-K189me2 peptide. Hydrogen
bonds from the Y305F structure are shown as green dashed lines, and waters and
hydrogen bonds in the Y245A structure are shown in yellow and orange,
respectively.
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Tyr-305 and Tyr-245, respectively (Fig. 3G). In the Y305F
mutant, hydrogen bonds to the dimethyl -amino group cou-
pled with steric constraints in the lysine binding channel pre-
vent the K189me2 side chain from undergoing a conforma-
tional change that is conducive to trimethylation (Fig. 2D),
consistent with its dimethyltransferase activity. However, in the
Y245A mutant, the alanine substitution enlarges the diameter
of the lysine binding channel, accommodating trimethyl-lysine
(Fig. 3D). In addition, the larger diameter would permit the
dimethyl-lysine substrate to undergo the conformational reor-
ganization necessary to align the -amino group in a productive
geometry for trimethylation.
A major difference in the active site of the Y245A mutant
versus the other SET7/9 structures is the presence of several
water molecules bound in the cavity generated by the Y245A
mutation. In the structure of the Y245A mutant bound to
TAF10-K189, three water molecules (waters 2–4) occupy this
cavity and are arranged in a triangular geometry (Fig. 3A). In
addition, water 1 shifts 1.6 Å from its position in the solvent
pocket toward water 2 to which it forms a hydrogen bond (Fig.
3, A and E). The shift in water 1 was unexpected given its con-
served orientation in the solvent pocket of the SET7/9 WT and
Y305F complexes (Fig. 2, A–C) as well as in the structures of
other SET domain KMTs (8). This displacement is presumably
related to the alternative conformation of the K189 side chain
whose -amino group is too distant (4.3 Å) to form a productive
hydrogen bond to water 1. Conversely, in the Y245A complexes
bound to TAF10-K189me1 and TAF10-K189me2, water 1
remains tightly bound in the solvent pocket through hydrogen
bonds to Tyr-305 hydroxyl group, the carbonyl oxygens of Gly-
292 and Ala-295, and the K189 -amino group (Fig. 3, B, C, and
E), analogous to its binding in the WT enzyme (Fig. 2A). How-
ever, in the TAF10-K189me3 complex, one of the methyl
groups of the trimethyl -ammonium cation is oriented into the
solvent pocket (Fig. 3D), similar to the dimethyl-lysine binding
mode observed in the Y305F mutant (Fig. 2D). The binding of
the methyl group in the solvent pocket displaces water 1 by 3.2
Å relative to its position in the TAF10-K189me1 complex (Fig.
3E), thereby avoiding a steric clash with the trimethylated
-ammonium group. Variations in the occupancy of water 2 are
also seen in the different Y245A structures. Water 2 is bound in
similar orientations in the active site of the unmodified and
monomethylated peptide complexes but is absent in the di- and
trimethylated peptide complexes due to the binding of a methyl
group in this position (Fig. 3, A–E). In summary, the changes in
the positions or occupancies of waters 1 and 2 correlate with the
binding modes of the unmodified and methylated K189 within
the active site of the Y245A mutant.
Catalytic Models of Lysine Multiple Methylation by SET7/9
Y245A, and Y305F—The structures of the SET7/9 complexes
reported here offer a prime opportunity to generate stepwise
models for lysine mono-, di-, and trimethylation by a SET
domain KMT. We modeled the AdoMet-bound Michaelis
complexes by superimposing the SET7/9 product complexes
with the previously reported structure of the SET7/9-AdoMet
binary complex (Fig. 4) (39). The conformations of the mono-
and dimethyl -amino groups in the Michaelis complexes were
inferred from the coordinates of the corresponding dimethyl-
and trimethyl-lysine products, respectively. In addition, we
modeled the -amino group in a deprotonated state with its
hydrogen atoms oriented toward the hydrogen bond acceptors
that align the lysyl side chain for methylation. As a basis for this
comparison, we first modeled the monomethylation reaction
catalyzed by WT SET7/9 (Fig. 4A). In the substrate ternary
complex, the lysine -amine is aligned with the methyl group
and sulfonium cation of AdoMet through a hydrogen bond to
the Tyr-245 hydroxyl group and water 1 in the solvent pocket.
The values of the reaction distance and angle are 2.8 Å and 153°,
respectively, in approximate agreement with the linear geome-
try of a SN2 methyl transfer reaction calculated in other mod-
eled substrate complexes (8, 10). In the product complex, the
monomethyl-lysine side chain is bound in an extended confor-
mation with its methyl group oriented within the methyl trans-
fer pore, thereby obstructing AdoMet binding. Furthermore,
water 1 remains tightly coordinated in the solvent pocket
through four hydrogen bonds to Gly-292, Ala-295, Tyr-305,
and the monomethyl -amino group. These interactions hinder
the dissociation of water 1 and the related rearrangement of the
monomethyl-lysine side chain required for a second methyl
transfer reaction, explaining why the WT enzyme cannot cata-
lyze di- and trimethylation. These findings concur with the
FIGURE 4. Catalytic models of the methyl transfer reactions catalyzed by
WT SET7/9 and the Y305F mutant. A, monomethylation of TAF10-K189 by
the WT enzyme. The reaction scheme depicts the modeled substrate ternary
complex (left) and the product complex (right) for the transfer of the methyl
group from AdoMet (green carbon atoms) to K189 in TAF10 (yellow carbons),
yielding AdoHcy and K189me1. The red arrow indicates the direction of the
nucleophilic attack of the deprotonated -amino group on the AdoMet
methyl group. The transferred methyl group is colored green, and the white
atoms represent the hydrogens of the -amino group. Hydrogen bonds
and residues in the enzyme active site are illustrated as in Fig. 2. The
reaction distance and angle are labeled in red. B and C, models of the
Y305F mutant for the first methyl transfer reaction with TAF10-K189 (B)
and second methyl transfer reaction with TAF10-K189me1 (C). Color
schemes are the same as in A.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY 31855
model for SET7/9 product specificity reported in previous
structural and functional studies (6, 11).
Similar reaction geometry is observed in the model for the
monomethyl transfer reaction catalyzed by SET7/9 Y305F.
Hydrogen bonds from the Tyr-245 hydroxyl group and water 1
align the lysine -amino group with the AdoMet methyl group
at a distance of 2.1 Å and an angle of 160°, equivalent to those
measured in the Michaelis complex of the WT enzyme (Fig. 4, A
and B). In the product complex, monomethyl-lysine adopts an
extended trans configuration analogous to that in the WT
enzyme. For dimethylation to occur, the monomethyl-lysine
must undergo a conformational change in which its methyl
group is rotated out of the methyl transfer path with AdoMet.
The structure of the Y305F mutant bound to the dimethylated
TAF10 peptide (Fig. 2D) implies that this rearrangement occurs
through the dissociation of water 1 due to the loss of the Tyr-
305 hydrogen bond in the solvent pocket. The dissociation of
water 1 would enable the monomethyl-lysine side chain to
adopt an alternative conformation through a rotation about its
C–N bond, projecting the methyl group into the solvent
pocket (Fig. 4C). This rotation reorients the methyl group out of
the methyl transfer path while realigning the monomethyl -
amino group for a second methylation reaction through a direct
hydrogen bond to the Tyr-245 hydroxyl group and a CH–O
hydrogen bond between its methyl group and Ala-295. The
modeled reaction geometry for monomethyl-lysine substrate
complex of the Y305F mutant (2.7 Å, 160°) is equivalent to that
of the first methyl transfer reaction in SET7/9 Y305F. These
geometries concur with our previous models for mono- and
dimethylation catalyzed by SET8 Y334F (8), illustrating that the
orientation of a methyl group into the solvent pocket is a con-
served feature of SET domain KMTs that catalyze multiple
methylation.
In addition, we modeled the methyl transfer reactions cata-
lyzed by SET7/9 Y245A (Fig. 5). In the model of the lysine sub-
strate complex, the -amino group is aligned for methyl transfer
by a hydrogen bond to the carbonyl oxygen of Gly-264, result-
ing in a short reaction distance (2.3 Å) and a suboptimal reac-
tion angle (141°) with the methyl group of AdoMet (Fig. 5A).
This misalignment appears to be a direct consequence of the
Y245A mutation that abolishes hydrogen bonding to the -
amino group, illustrating that the suboptimal orientation of the
-amine likely contributes to the diminished activity of this
mutant toward unmodified substrates (11). Conversely, in the
modeled monomethyl-lysine substrate complex for SET7/9
Y245A, Tyr-305 hydrogen bonds to the -amino group, aligning
it for methyl transfer (Fig. 5B). In addition, CH–O hydrogen
bonds to the monomethyl-lysine methyl group and the dissoci-
ation of water 2 from the active site also contribute to reposition-
ing the -amino group for dimethylation. Collectively, these inter-
actions orient the -amine in a reaction angle of 165° that is more
conducive to methyl transfer. However, the reaction distance for
dimethylation is 0.6 Å longer than that in the corresponding
Y305F model because Tyr-305 is positioned further from
AdoMet than Tyr-245 (Figs. 4C and 5B).
In the third methyl transfer reaction catalyzed by SET7/9
Y245A, the lone pair of electrons of the dimethyl-lysine -
amino group acts as the nucleophile and thus cannot engage in
hydrogen bonding. The structure of the trimethyl-lysine prod-
uct complex (Fig. 3D) implies that the dimethyl -amine is
aligned via CH–O hydrogen bonds to its methyl groups, as
shown in the model of the Michaelis complex for this reaction
(Fig. 5C). These CH–O hydrogen bonds restrain the orienta-
tion of the -amino group and position one of the methyl
groups into the solvent pocket, displacing water 1 as discussed
earlier (Fig. 3, D and E). These interactions cumulatively align
the -amino group and AdoMet methyl group with a reaction
distance of 3.0 Å and angle of 162° (Fig. 5C). Taken together, the
models of the substrate complexes for SET7/9 Y245A suggest
that CH–O hydrogen bonds play an increasingly important role
in aligning the methylated -amino group in successive rounds
of methyl transfer.
DISCUSSION
The structural and functional characterization of the
SET7/9 Y245A and Y305F mutants presented here yields
new insights into the mechanism underlying the product
specificity of SET domain KMTs. Importantly, it resolves a
general paradox concerning this specificity. How does the
active site constrain the motion of the lysine -amino group
to align it for methyl transfer with AdoMet, while providing
adequate volume to accommodate the mono-, di-, and tri-
methylated lysine side chain generated during multiple
methyl transfer reactions? The structures of the Y305F and
Y245A mutants resolve this paradox, illustrating that alter-
ations in the positions or occupancies of water molecules
within their active sites generate the space required to
FIGURE 5. Models for the methyl transfer reactions catalyzed by the
SET7/9 Y245A mutant. Models of the Y245A mutant for the first methyl
transfer reaction with TAF10-K189 (A), the second methyl transfer reaction
with TAF10-K189me1 (B), and the third methyl transfer reaction with TAF10-
K189me2 (C). Residues and hydrogen bonds are depicted as in Fig. 3.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
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accommodate the multiply methylated -amine produced
during successive catalytic cycles. Minor perturbations in
the side chains of certain active site residues, such as Tyr-
305, are also observed in alignments of the WT enzyme and
the Y245A and Y305F complexes, although these changes are
modest compared with the displacement or dissociation of
the water molecules in the active site. These findings suggest
that the waters function as transient place holders that facil-
itate the SN2 methyl transfer reaction. During monomethy-
lation, they function to constrain the movement of the lysine
-amino group by mediating hydrogen bonds between the
substrate and enzyme, thereby promoting the linear align-
ment with the methyl group and sulfonium cation of
AdoMet (Fig. 4, A and B). During di- and trimethylation, the
water molecules either relocate within the lysine binding
channel or dissociate from the enzyme, yielding the space
required to rotate the methyl group away from the methyl
transfer pore and to realign the -amine in productive geom-
etry for the next methyl transfer reaction (Figs. 4C and 5, B
and C). These findings agree with our prior analysis of the
SET8 Phe/Tyr switch mutant in which we demonstrated that
the Y334F substitution attenuates hydrogen bonding to the
water molecule bound in the solvent pocket, promoting its
dissociation and the conformational changes necessary for
lysine dimethylation (8). Indeed, there is a nearly identical
alignment of the dimethyl-lysine side chains in the structures of
SET7/9Y305FandSET8Y334Fcomplexes,despitethedifferences
in the orientations of the Phe-305 and Phe-334 side chains in each
structure (Fig. 2F). Finally, both the SET8 Y334F (8) and SET7/9
Y305F mutants (Table 2) displayed diminished catalytic efficien-
cies for lysine dimethylation versus monomethylation. These dif-
ferences may reflect the kinetics of the reorganization within the
active site, including the dissociation of the water molecule from
the solvent pocket and the concomitant realignment of the
monomethyl-lysine into a productive geometry for dimethylation.
In addition to their place-holding role, the active site waters
may also facilitate the deprotonation of the lysine -amino
group between methyl transfer reactions. For methylation to
occur, the -amino group must be deprotonated to function as
the nucleophile in the SN2 methyl transfer reaction with
AdoMet (Figs. 4 and 5). Although the pKa value of the lysine
-amine in solution is 10.5, molecular dynamics simulations
by Zhang and Bruice (25, 26) indicate that this value diminishes
to 8.2 upon formation of the SET7/9 Michaelis complex due to
the proximity of the AdoMet sulfonium cation and the low
dielectric constant of the active site. Furthermore, their simu-
lations show that a chain of water molecules facilitates the dep-
rotonation of the -amino group prior to methyl transfer, trans-
ferring the proton to bulk solvent. Although these water
molecule chains are not evident in our crystal structures, the
Y305F and Y245A complexes suggest another potential mech-
anism for deprotonation. In the dimethyl-lysine complexes of
the Y305F and Y245A mutants, the dissociation of water 1 and
2, respectively, from the lysine binding channel requires that
the solvent-mediated hydrogen bond to the -amino group is
broken (Figs. 2D and 3C). It is conceivable that these waters
dissociate from the active site as hydronium ions, promoting
the realignment and deprotonation of the methyl -amino
group for the next methyl transfer reaction.
A comparison of the SET7/9 Y305F and SET8 Y334F com-
plexes yields insights into the mechanism by which the Phe/Tyr
switch influences water binding within the solvent pocket. The
phenylalanine substitution in the Phe/Tyr switch results in the
loss of a single hydrogen bond to the water molecule (water 1) in
the solvent pocket compared with the four hydrogen bonds that
coordinate the solvent molecule in WT SET7/9 (Fig. 2, A and B)
and SET8 (7, 8). Although this attenuation in hydrogen bonding
may appear insignificant, this difference is nonetheless impor-
tant for at least two reasons. First, theoretical calculations indi-
cate that, on average, water molecules form 3.5 hydrogen
bonds in solutions (40, 41). This value is greater than the num-
ber of hydrogen bonds coordinating water 1 in the solvent
pocket in SET7/9 Y305F (Fig. 2, B and C) as well as in SET8
Y334F and other di- and trimethyltransferases that possess a
hydrophobic residue in the Phe/Tyr switch site (8). From the
perspective of the water molecule, the greater hydrogen bond-
ing potential in solution would tend to thermodynamically
favor its dissociation from the solvent pocket in SET domain
KMTs that lack a tyrosine in the Phe/Tyr switch position. Sec-
ond, the ordered binding of water molecules observed in the
active sites of SET domain ternary complexes represents an
unfavorable entropy compared with their diffusion in bulk sol-
vent. In WT SET7/9 (Fig. 2A) and SET8 (7, 8), this entropic
penalty can be partially offset through the favorable enthalpy of
binding associated with the four hydrogen bonds that coordi-
nate the water within the solvent pocket. It is conceivable that
the loss of the tyrosine-mediated hydrogen bond in the Phe/Tyr
switch shifts the equilibrium in favor of dissociation of the
water molecule from the solvent pocket, thereby facilitating
dimethylation in SET7/9 Y305F, SET8 Y334F, and other di- and
trimethyltransferases.
The structures of the SET7/9 Y245A and Y305F complexes
illustrate the interactions that align the lysine -amino group
during the methyl transfer reactions in each enzyme. In the WT
enzyme and the Y305F mutant, hydrogen bonding to the
hydroxyl group of Tyr-245 appears to be critical in properly
aligning the -amine for methyl transfer (Fig. 4). Tyr-245 is
conserved in the sequences of many SET domain KMTs (8, 42),
and substitutions of this residue generally impair or abolish
activity, indicating its importance in catalysis (8, 43). However,
SET7/9 appears to be an exception to this rule, as the Y245A
mutant is not only active but is capable of catalyzing lysine
trimethylation. In this mutant, Tyr-305 appears to assume the
role of Tyr-245 by hydrogen bonding to the monomethylated
-amino group to align it for methyl transfer with AdoMet, as
illustrated in the modeled substrate complex for the dimethy-
lation reaction (Fig. 5B). Conversely, in the model for trimethy-
lation, the Tyr-305 hydroxyl group does not hydrogen bond to
the -amine but instead participates in a CH–O hydrogen bond
with one of the methyl groups to assist in aligning the dimethy-
lated -amine for the methyl transfer reaction (Fig. 5C). Addi-
tional structural and functional studies of the SET domain tri-
methyltransferases will aid in further illuminating the roles
of CH–O hydrogen bonds in facilitating lysine multiple
methylation.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31857
Acknowledgments—We acknowledge S. Schiebold for assistance in
protein expression, purification, and crystallization and S. Anderson
and R. Sanishvili for their assistance with x-ray data collection. We
also thank S. Bulfer and S. Horowitz for reading the manuscript and
providing useful comments. This work utilized the Protein Structure
Facility of the Michigan Diabetes Research and Training Center, Uni-
versity of Michigan, supported by National Institutes of Health Grant
DK020572, NIDDK. Use of the Advanced Photon Source was sup-
ported by the United States Department of Energy, Basic Energy Sci-
ences, Office of Science, under Contract DE-AC02-06CH11357.
GM/CA CAT has been funded in whole or in part by National
Institutes of Health NCI Grant Y1-CO-1020 and NIGMS Grant
Y1-GM-1104. Use of the LS-CAT Sector 21 was supported by Michi-
gan Economic Development Corporation and the Michigan Technol-
ogy Tri-Corridor Grant 085P1000817 for the support of this research
program.
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Lysine Methylation by SET7/9 Mutants
31858
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
|
3M5A
|
SET7/9 Y245A in complex with TAF10-K189me3 peptide and AdoHcy
|
SET7/9 Catalytic Mutants Reveal the Role of Active Site Water
Molecules in Lysine Multiple Methylation*□
S
Received for publication,February 15, 2010, and in revised form, July 14, 2010 Published, JBC Papers in Press,August 1, 2010, DOI 10.1074/jbc.M110.114587
Paul A. Del Rizzo‡, Jean-Franc¸ois Couture‡§1, Lynnette M. A. Dirk¶, Bethany S. Strunk‡, Marijo S. Roiko‡,
Joseph S. Brunzelle, Robert L. Houtz¶, and Raymond C. Trievel‡2
From the ‡Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109, the §Ottawa Institute of
Systems Biology, Department of Biochemistry, Microbiology, and Immunology, University of Ottawa, Ottawa, Ontario K1H 8M5,
Canada, the ¶Department of Horticulture, Plant Physiology/Biochemistry/Molecular Biology Program, University of Kentucky,
Lexington, Kentucky 40546, and the Department of Molecular Pharmacology and Biological Chemistry, Northwestern University
Center for Synchrotron Research, Life Sciences Collaborative Access Team, Argonne, Illinois 60439
SET domain lysine methyltransferases (KMTs) methylate
specific lysine residues in histone and non-histone substrates.
These enzymes also display product specificity by catalyzing dis-
tinct degrees of methylation of the lysine -amino group. To
elucidate the molecular mechanism underlying this specificity,
we have characterized the Y245A and Y305F mutants of the
human KMT SET7/9 (also known as KMT7) that alter its prod-
uct specificity from a monomethyltransferase to a di- and a tri-
methyltransferase, respectively. Crystal structures of these
mutants in complex with peptides bearing unmodified, mono-,
di-, and trimethylated lysines illustrate the roles of active site
water molecules in aligning the lysine -amino group for methyl
transfer with S-adenosylmethionine. Displacement or dissocia-
tion of these solvent molecules enlarges the diameter of the
active site, accommodating the increasing size of the methylated
-amino group during successive methyl transfer reactions.
Together, these results furnish new insights into the roles of
active site water molecules in modulating lysine multiple meth-
ylation by SET domain KMTs and provide the first molecular
snapshots of the mono-, di-, and trimethyl transfer reactions
catalyzed by these enzymes.
SET domain enzymes represent a family of S-adenosylmethi-
onine (AdoMet)3-dependent methyltransferases that catalyze
the site-specific methylation of protein lysyl residues in a host
of proteins, including histones, transcription factors, chroma-
tin-modifying enzymes, ribosomal subunits, and other sub-
strates (1–3). In many instances, these modifications serve to
recruit effector proteins that recognize methyl-lysyl residues in
a sequence-dependent fashion (4). In addition, SET domain
KMTs exhibit product specificity, defined as their ability to cat-
alyze mono-, di-, or trimethylation of the lysine -amino group.
This specificity is biologically relevant because many methyl-
lysine-binding proteins can discriminate among different
degrees of lysine methylation (4). Thus, both the site and degree
of lysine methylation are critical to recognition by effector
proteins.
Structural and functional studies have identified a Phe/Tyr
switch in the active site of SET domain KMTs that governs their
respective product specificities (5, 6). According to this model,
KMTs that possess a tyrosine in the Phe/Tyr switch site are
limited to catalyzing lysine monomethylation, whereas en-
zymes that possess a phenylalanine or another hydrophobic
residue in this position display di- or trimethyltransferase activ-
ity. Mutational analysis of various SET domain KMTs, includ-
ing DIM-5 (KMT1) (6), SET7/9 (6), G9A (KMT1C) (5), and
SET8 (also known as PR-SET7 and KMT5A) (7, 8), has demon-
strated that substitutions in the Phe/Tyr switch result in pre-
dictable changes in product specificity. Several models have
been proposed to explain the mechanism by which the Phe/Tyr
switch site governs this specificity, including variations in the
diameter of the active site due to the size of Phe/Tyr switch
residue and steric hindrance by the tyrosine hydroxyl group (6,
9–11). However, our recent studies of the Phe/Tyr switch
mutant Y334F in the human histone H4 Lys-20 (H4K20) meth-
yltransferase SET8 indicate that the Phe/Tyr switch regulates
product specificity via a more subtle mechanism (8). Specifi-
cally, the switch modulates the binding of an active site water
molecule that in turn regulates the transition from mono-
methylation to multiple methylation.
Among the KMTs that have been structurally characterized,
SET7/9 has emerged as an archetypal model for studying the
catalytic mechanism and product specificity of the SET domain
family. Although initially isolated as a histone H3 Lys-4
(H3K4)-specific methyltransferase, this KMT has been shown
to regulate the functions of numerous non-histone substrates
through site-specific methylation (12–21). Early structural and
functional studies of SET7/9 identified two active site mutants,
* Thisworkwassupported,inwholeorinpart,byNationalInstitutesofHealth
Grant R01 GM073839 (to R. C. T.) and National Institutes of Health Admin-
istrative Supplement GM073839-04S1 (to R. C. T.) funded through the
American Recovery and Reinvestment Act. This work was also supported
by Department of Energy Grant DE-FG02-92ER20075 (to R. L. H.).
□
S The on-line version of this article (available at http://www.jbc.org) contains
supplemental Table 1.
The atomic coordinates and structure factors (codes 3M53, 3M54, 3M55, 3M56,
3M57,3M58,3M59,and3M5A)havebeendepositedintheProteinDataBank,
Research Collaboratory for Structural Bioinformatics, Rutgers University, New
Brunswick, NJ (http://www.rcsb.org/).
1 Supported by a Canadian Institutes of Health Research postdoctoral
fellowship.
2 To whom correspondence should be addressed: Dept. of Biological Chem-
istry, University of Michigan Medical School, 1150 West Medical Center Dr.,
5301 MSRB III, Ann Arbor, MI 48109. Tel.: 734-647-0889; Fax: 734-763-4581;
E-mail: rtrievel@umich.edu.
3 The abbreviations used are: AdoMet, S-adenosylmethionine; AdoHcy, S-ad-
enosylhomocysteine; KMT, lysine methyltransferase; ITC, isothermal titra-
tion calorimetry; CH–O, carbon-oxygen hydrogen bond; BisTris, 2-[bis(2-
hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 41, pp. 31849–31858, October 8, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31849
Y245A and Y305F, which change its product specificity. The
Phe/Tyr switch mutant Y305F alters SET7/9 product specificity
from a mono- to dimethyltransferase (6), whereas the Y245A
substitution converts the enzyme into a trimethyltransferase
with weak monomethyltransferase activity (11). These mutants
have been the subjects of numerous molecular modeling simu-
lations that have led to various models to explain their distinct
product specificities (22–26). However, the lack of structural
data for the SET7/9 Y245A and Y305F mutants in complex with
cognate methylated peptides has hindered our understanding
of the mechanisms that define the respective product specific-
ities of these mutants. Moreover, these structures would yield a
framework for visualizing the mono-, di-, and trimethylation
reactions catalyzed by SET domain KMTs.
To gain insight into the molecular basis of their product
specificities, we have determined high resolution crystal
structures of the SET7/9 Y245A and Y305F mutants in com-
plex with peptides of the TATA box-binding protein-associ-
ated factor TAF10 bearing the Lys-189 methylation site in
unmodified (K189), monomethylated (K189me1), dimethyl-
ated (K189me2), and trimethylated (K189me3) states. The
structures and accompanying biochemical data support a
model whereby changes in the occupancy or position of water
molecules in the active site are critical in establishing the prod-
uct specificities of the SET7/9 Y245A and Y305F mutants.
Together, our results provide new insights into the mechanisms
that govern SET domain product specificity and provide step-
wise snapshots of the lysine mono-, di-, and trimethyl transfer
reactions catalyzed by KMTs.
EXPERIMENTAL PROCEDURES
Cloning, Expression, and Purification of the SET7/9 Mutants—
The Y245A and Y305F mutants were introduced into the pHIS2
SET7/9 expression vector encoding residues 110–366 (27) via
QuikChange site-directed mutagenesis (Stratagene) and were
verified by dideoxy DNA sequencing. The plasmids encoding
wild type (WT) SET7/9 and the Y245A and Y305F mutants
were transformed into Rosetta2 DE3 cells (Novagen) and were
expressed as described previously (27, 28). In the course of
characterizing WT SET7/9, we observed that the enzyme co-
purified with AdoMet or another contaminant that resulted in
technical difficulties in the isothermal titration calorimetry
(ITC) experiments and co-crystallization trials with the TAF10
peptides. To overcome this problem, a denaturation and refold-
ing step was inserted in the purification scheme. The denatur-
ation and refolding protocol involved adding 6 M guanidine
HCl, 50 mM sodium phosphate, pH 7.5, and 500 mM NaCl to the
protein while it was immobilized on a nickel-Sepharose column
(GE Healthcare). The column was washed with this buffer, fol-
lowed by a solution of 6 M urea, 50 mM sodium phosphate, pH
7.5, 150 mM NaCl, and 10 mM 2-mercaptoethanol to remove the
cofactor from the denatured enzyme. A reverse gradient from 6
to 0 M urea was then performed in the same buffer to refold the
protein, which was subsequently eluted from the column using
a linear gradient of 0–500 mM imidazole in 50 mM sodium
phosphate, pH 7.5, 500 NaCl, and 10 mM 2-mercaptoethanol.
The refolded protein was digested with tobacco etch virus pro-
tease (29) during dialysis against 20 mM Tris, pH 7.5, 150 mM
NaCl, and 5 mM 2-mercaptoethanol and then purified using a
Superdex 200 gel filtration column (GE Healthcare). Protein
concentration was determined by its absorbance at 280 nm.
Synthetic Peptides—The TAF10 peptides bearing K189,
K189me1,
K189me2,
and
K189me3
(sequence,
acetyl-
SKSK189DRKYTL-amide) and a biotinylated TAF10 peptide
(sequence,
acetyl-SKSK189DRKYTLT(K-EZLink-S-S-biotin)-
amide) were synthesized and purified by New England Peptide,
Inc. Peptide concentrations were measured using the absorb-
ance of their tyrosine residue at 274 nm.
Crystallization and Data Collection—Crystals were pro-
duced by hanging drop vapor diffusion by mixing the crystalli-
zation solution in a 1:1 ratio with 10–20 mg/ml SET7/9, 1 mM
S-adenosylhomocysteine (AdoHcy), and 1.0–3.0 mM unmodi-
fied or methylated TAF10-K189 peptide in 20 mM Tris, pH 8.0,
100 mM NaCl, and 2 mM tris(2-carboxyethyl)phosphine. Crys-
tals were obtained at 20 °C in either 1.8–2.0 M (NH4)2SO4, with
0.1 M BisTris, pH 6.2–6.6, or in 0.95–1.1 M sodium citrate with
100 mM imidazole pH 8.0–8.4. In both crystallization condi-
tions, the final pH values were between pH 8.0 and 9.0. Crystals
in the (NH4)2SO4 condition were typically flash-frozen in the
mother liquor containing 25–30% glycerol, and the crystals in
the citrate condition were frozen in 1.6 M sodium citrate. Data
were collected at the Advanced Photon Source beamlines
21-IDG (LS-CAT) and 23-IDD (GM/CA-CAT). Images were
indexed, integrated, and scaled using HKL2000 (30). Structures
of the mutants were solved by molecular replacement using
MOLREP (31) with the coordinates of a previously reported
SET7/9 ternary complex used as the search model (Protein
Data Bank code 2F69). Successive rounds of model building and
refinement were carried out using Coot (32) and REFMAC (33),
respectively. The geometry of the models were verified by Mol-
Probity (34). Simulated annealing omit maps were calculated
using CNS (35) with the peptide and cofactor removed to elim-
inate model bias in the active site. Structural figures were ren-
dered using PyMOL (Schro¨dinger, LLC.).
Fluorescent Methyltransferase Assay—A coupled fluorescent
methyltransferase assay was used to measure the kinetic
parameters of WT SET7/9 and the Y245A and Y305F mutants
as reported previously, with the exception that 50–150 nM
enzyme, 100 M AdoMet, and varying concentrations of TAF10
peptide substrate were used (27, 36). Assays were performed in
triplicate, and a homocysteine calibration curve was used to
calculate the initial velocities. Kinetic parameters were calcu-
lated by plotting the velocities versus peptide concentration and
by fitting the Michaelis-Menten equation to the data via non-
linear regression using Prism 5.0 (GraphPad). In cases where
the Km value was beyond the measurable range of the assay, the
kcat/Km value was determined as described previously (7).
Calorimetry Experiments—ITC was performed at 20 °C using
a MicroCal VP-ITC calorimeter (GE Healthcare) with 0.12 mM
protein and 4 mM AdoHcy in 20 mM sodium phosphate, pH 7,
and 100 mM NaCl with 1.5 mM peptide as the injectant. Data
were processed, and equilibrium dissociation constants (KD)
and curve fitting errors were calculated from the binding iso-
therms using Origin 7.0 (OriginLab Corp.). WT SET7/9 and the
Y245A and Y305F mutants displayed ligand:protein binding
stoichiometries (N values) between 0.8 and 1.0, demonstrating
Lysine Methylation by SET7/9 Mutants
31850
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
that WT SET7/9 and its mutants were properly refolded due to
their ability to bind peptides in an 1:1 molar ratio.
TLC Product Analysis—Methyltransferase assays were per-
formed in triplicate at 37 °C with the biotinylated TAF10 pep-
tide (0.5 mM) for 30 min in the presence of WT SET7/9 (3
pmol), Y305F mutant (6 pmol), or the Y245A mutant (100
pmol). Assays contained 100 mM HEPES, pH 8.0, 50 mM NaCl,
1.0 mM [methyl-3H]AdoMet (diluted to a specific activity of 0.2
Ci/nmol with purified AdoMet (37)), 15 M Sulfolobus solfa-
taricus AdoHcy hydrolase (36), and 2 units of adenosine deami-
nase (Roche Applied Science) in a final volume of 20 l. The
reactions were terminated by addition of an equal volume of
200 mM MES, pH 5.1, prior to addition of a 2-fold molar excess
of immobilized avidin resin (UltraLink; Pierce). Biotinylated
peptides were allowed to bind at room temperature for 30 min,
and the resin was then collected by centrifugation (9000 g).
The resin was washed three times with 300 mM NaCl, and the
peptide was eluted overnight from the avidin resin by cleavage
of the disulfide bond in the linker of the peptide using 10 mM
tris(2-carboxyethyl)phosphine (Thermo Scientific). The resin
was incubated with additional 10 mM tris(2-carboxyethyl)phos-
phine the following day until the radiolabel was essentially
removed from the resin. The recovered peptides were hydro-
lyzed by incubation with 6 M HCl at 110 °C for 24 h. Subsequent
steps in measuring the radiolabel incorporated into the mono-,
di-, and trimethyl-lysine products were performed as reported
previously (8).
RESULTS
Functional Analysis of the SET7/9
Y305F Mutant—Prior studies of
SET7/9 by Zhang et al. (6) reported
that mutation of the Phe/Tyr switch
residue Tyr-305 to a phenylalanine
alters its product specificity from a
mono- to dimethyltransferase. We
verified these findings by demon-
strating that WT SET7/9 mono-
methylated the TAF10-K189 pep-
tide, whereas the Y305F mutant
mono- and dimethylated this sub-
strate, as demonstrated by mass
spectrometry (data not shown). We
next examined whether the Y305F
substitution altered the affinity of SET7/9 for the TAF10-K189
peptides using ITC (Fig. 1). A comparison of the KD values
revealed that SET7/9 Y305F bound the TAF10-K189 and
TAF10-K189me1 peptides 4- and 6-fold more tightly, respec-
tively, than the WT enzyme, whereas this mutant displayed a
substantially diminished affinity for the TAF10-K189me2 pep-
tide (Table 1). Although the WT enzyme and the Y305F mutant
exhibited discernable differences in their affinities for the
unmodified and monomethylated peptides, these variations are
modest and cannot account for their distinct product specific-
ities, suggesting that a kinetic effect during methylation may be
responsible.
To investigate this possibility, we characterized the kinetic
parameters of WT SET7/9 and the Y305F mutant using the
TAF10 peptides as substrates. Both enzymes methylated the
unmodified peptide with comparable kcat and Km values (Table
2). In analyzing the kinetic parameters for the methylation
of the monomethylated peptide by SET7/9 Y305F, we found
that this substrate displayed an elevated Km value that was
beyond the measurable range of the assay due to its limited
solubility. In this case, we measured the catalytic efficiency
(kcat/Km) for the methylation of this peptide and found that it
was methylated 15-fold less efficiently than the unmodified
peptide by SET7/9 Y305F. Given the fact that the Y305F mutant
exhibited a higher binding affinity for the TAF10-K189me1
peptide than the WT enzyme (Table 1), the kinetic data suggest
that a step in the reaction pathway following substrate binding
limits the catalytic efficiency of this mutant.
We next examined whether the Y305F mutant dimethylated
the TAF10-K189 peptide via a processive or a distributive
mechanism. In a processive mechanism, the methyl-lysine
substrate would remain bound to the enzyme during successive
methyl transfer reactions; thus, the concentration of an inter-
mediate, such as monomethyl-lysine, cannot exceed the en-
zyme concentration during the assay. In a distributive mecha-
nism, the intermediates are released into solution where they
accumulate prior to the next round of methylation, resulting in
an intermediate concentration that is greater than that of the
enzyme. Using a radiometric TLC assay and a biotinylated
TAF10 peptide, we quantified the amounts of monomethylated
products generated by the WT SET7/9 and the Y305F mutant
FIGURE 1. ITC analysis of WT SET7/9 and the Y245A and Y305F mutants. Representative titrations and
binding isotherms for the calorimetry experiments are shown for the unmodified TAF10-K189 peptide titrated
into WT SET7/9 (left plot), SET7/9 Y305F (middle plot), and SET7/9 Y245A (right plot). In each panel, the ITC
titration experiment (upper panel) is shown with the binding isotherms (lower panel) fit to a single binding site
model.
TABLE 1
Analysis of the binding affinity of WT SET7/9 and its catalytic
mutants for unmodified and methylated TAF10 peptides
SET7/9
TAF10 peptide
KD
a
M
WT
K189
4.9 0.20
WT
K189me1
4.0 0.36
Y305F
K189
1.3 0.10
Y305F
K189me1
0.62 0.065
Y305Fb
K189me2
70
Y245A
K189
4.0 0.25
Y245A
K189me1
3.3 0.10
Y245A
K189me2
5.8 0.22
Y245A
K189me3
11 0.28
a Curve fitting errors were calculated from the binding isotherms.
b An estimate of the affinity is reported due to weak peptide binding.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31851
(Table 3). The data demonstrate that comparable amounts of
monomethyl-lysine were generated when the quantity of
enzyme usedistakenintoaccount,inagreementwiththeirsimilar
turnover numbers for the TAF10-K189 peptide (Table 2). The
Y305F mutant also produced small but measurable quantities of
radiolabeled dimethyl-lysine product that were substantially
smaller than the amount of monomethyl-lysine generated.
Therefore, the TLC data are consistent with a distributive
mechanism for dimethylation by the Y305F mutant because the
amount of monomethyl-lysine produced exceeded the quantity
of enzyme used in the assay.
Structures of WT SET7/9 and the Y305F Mutant in Complex
with Unmodified and Methylated TAF10 Peptides—To deter-
mine the mechanism by which the Y305F substitution alters the
product specificity of SET7/9, we determined the crystal struc-
tures of this mutant bound to AdoHcy and TAF10-K189,
TAF10-K189me1, and TAF10-K189me2 peptides and com-
pared these to the structures of the WT SET7/9AdoHcy
TAF10-K189 complex (supplemental Table 1). The structures
of these complexes were determined to 1.85 Å or higher reso-
lution, permitting unambiguous modeling of the K189 side
chains in the active site of the enzyme based on simulated
annealing omit maps (Fig. 2). The ternary complexes of the WT
and the Y305F mutant superimpose with overall root mean
square differences of less than 0.3 Å for all aligned atoms, indi-
cating that neither the Y305F mutation nor the binding of the
various TAF10-K189 peptides results in substantial changes in
its overall structure.
An inspection of the active sites of the SET7/9 WT and
Y305F complexes illustrates the binding modes of the unmod-
ified and methylated forms of K189 in the TAF10 peptides (Fig.
2, A–D). The K189 side chain binds in an extended all trans
conformation in a deep pocket, termed the lysine binding chan-
nel, that is composed of Tyr-245, Thr-266, Leu-267, Ser-268,
Tyr-305, Tyr-335, and Tyr-337 (not shown for clarity) (Fig. 2A).
These residues interact with the aliphatic portion of the K189
side chain primarily through van der Waals contacts. The lysine
binding channel connects to the AdoMet-binding site on the
opposite face of the catalytic domain via an oxygen-lined
methyl transfer pore (38). During catalysis, the methyl group of
the cofactor is positioned within the methyl transfer pore
for the SN2 reaction with the -amino group of the lysine or
methyl-lysine substrate (see below).
To lower the activation barrier for this reaction, the lysine
-amine nucleophile is aligned for methyl transfer through a
hydrogen bond network within the active site. In the WT
enzyme, the K189 -amino group hydrogen bonds to the
hydroxyl group of Tyr-245 as well as to two water molecules
(Fig. 2A). One of the water molecules (termed water 1), is coor-
dinated in a solvent pocket, through hydrogen bonds to the
carbonyl oxygens of Gly-292 and Ala-295 and to the hydroxyl
group of the Phe/Tyr switch residue Tyr-305. This solvent
pocket is structurally conserved in SET domain KMTs and has
an important role in defining product specificity through the
adjacent Phe/Tyr switch residue, as shown in our prior studies
of the human H4K20 methyltransferase SET8 (8). The other
water molecule is bound within the methyl transfer pore
between the lysine substrate and the thioether sulfur atom of
AdoHcy through hydrogen bonds to Tyr-245, Asn-265, and
His-293 in SET7/9 and the TAF10-K189 -amino group. This
water is not observed in other structures of SET7/9 ternary
complexes and may represent the approximate position that
the AdoMet methyl group occupies in the methyl transfer pore
in the Michaelis complex.
In structures of the Y305F ternary complexes, the K189,
K189me1, and K189me2 side chains also adopt extended trans
side chain geometries within the lysine binding channel that are
stabilized via hydrogen bonding to Tyr-245 in the enzyme (Fig.
2, B–D). The orientations of the K189me1 and K189me2 side
chains are further maintained through carbon-oxygen (CH–O)
hydrogen bonding between the methyl groups and oxygen
atoms within the vicinity of the methyl transfer pore, as
reported previously in other SET domain KMT structures (8,
10, 38). A superimposition of the SET7/9 WT and Y305F com-
plexes underscores the similarity of the lysyl binding conforma-
tions (Fig. 2E). However, there are notable differences in the
hydrogen bond patterns and occupancy of water 1 within the
solvent pocket in the Y305F mutant compared with the WT
enzyme. Specifically, the Y305F substitution results in the loss
of one hydrogen bond to water 1 in the structures of the TAF10-
K189 and TAF10-K189me1 complexes (Fig. 2, B and C). In con-
trast, water 1 is absent in TAF10-K189me2 complex, and the
vacated solvent pocket is occupied by one of the methyl groups
TABLE 2
Kinetic parameters of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
TAF10 peptide substrate
Km
a
kcat
a
kcat/Km
a
M
min1
M1 min1 103
WT
K189
160 17
17 0.62
110 17
Y305F
K189
88 5.0
17 0.30
190 11
Y305Fb
K189me1
11 0.50
Y245A
K189
200 35
0.53 0.04
2.6 0.47
Y245A
K189me1
210 23
5.9 0.23
28 3.3
Y245A
K189me2
400 29
6.5 0.16
15 1.2
a Curve fitting errors were calculated from the hyperbolic fits for the Michaelis-Menten equation.
b Km and kcat were not determined because the Km value was beyond the measurable range; therefore, the kcat/Km value is reported.
TABLE 3
Product analysis of WT SET7/9 and the Y305F and Y245A mutants
Enzyme
Quantity
of enzyme
Measured
product
Amount of
product formeda
nmol
nmol
WT
0.003
Kme1
0.65 0.07
Y305F
0.006
Kme1
1.5 0.49
Kme2
0.033 0.009
Y245A
0.100
Kme1
0.80 0.22
Kme2
0.39 0.021
Kme3
0.076 0.019
a Standard deviation was calculated from triplicate measurements.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
of the dimethyl -amine (Fig. 2D). This methyl group forms a
3.5-Å CH–O hydrogen bond to the carbonyl oxygen of Ala-295,
further stabilizing the binding of the dimethyl-lysine side chain.
A homologous dimethyl-lysine-binding mode and CH–O
hydrogen bond was observed in our prior structural studies of
the SET8 Y334F Phe/Tyr switch mutant that confers an analo-
gous change in product specificity from a mono- to a dimeth-
yltransferase (8). A structural alignment of the active sites of the
SET7/9 Y305F and SET8 Y334F mutants bound to cognate
dimethylated peptides illustrates that the coordinates of the
dimethyl-lysyl side chains are virtually superimposable, with
one methyl group oriented toward the methyltransfer pore and
the second positioned within the vacant solvent pocket (Fig.
2F). Taken together, the structures of the SET7/9 Y305F com-
plexes and the similarities in the dimethyl-lysine conforma-
tions in the SET7/9 Y305F and SET8 Y334F mutants imply that
the Phe/Tyr switch governs product specificity through a con-
served mechanism whereby it indirectly influences the binding
modes of the methyl-lysine side chain by modulating the affin-
ity of the water molecule (water 1) bound in the solvent pocket.
Biochemical Characterization of
the SET7/9 Y245A Mutant—Previ-
ous studies by Xiao et al. (11)
reported that the Y245A mutation
yields an unusual change in the
product
specificity
of
SET7/9,
converting the enzyme to a trimeth-
yltransferase with weak monometh-
yltransferase activity. We deter-
mined that the SET7/9 Y245A could
mono-, di-, and trimethylate the
TAF10-K189 peptide by mass spec-
trometry (data not shown) and TLC
(Table 3), confirming the earlier
studies of Xiao et al. (11). ITC
analysis revealed that the Y245A
mutant displayed comparable KD
values
for
the
unmodified
and
methylated TAF10-K189 peptides
(Fig. 1), although its affinity for the
trimethylated peptide was modestly
diminished in comparison with the
other peptides (Table 1). The ITC
data demonstrate that the Y245A
mutant
bound
the
unmodified,
mono-,
and
dimethylated
sub-
strates with equivalent affinities,
suggesting that a kinetic effect or
a structural alteration in the active
site may be responsible for its
diminished
activity
toward
un-
modified substrates.
To gain further insight into its
peculiar
product
specificity,
we
characterized the kinetic properties
of
the
SET7/9
Y245A
mutant.
Steady state analysis demonstrated
that this mutant displayed similar
Km values for the unmodified, mono- and dimethylated TAF10
peptides (Table 2). However, the turnover number for the
TAF10-K189 peptide was diminished over 10-fold versus the
methylated peptides and was reduced 30-fold versus the WT
enzyme, in agreement with the weak monomethyltransferase
activity reported by Xiao et al. (11). In addition, we investigated
whether this mutant catalyzes lysine trimethylation via a pro-
cessive or distributive mechanism as described for SET7/9
Y305F. The TLC data illustrate that the mono- and dimethyl-
lysine intermediates accumulated at quantities greater than
that of the enzyme used in the assay, indicating that SET7/9
Y245A obeys a distributive mechanism, analogous to the Y305F
mutant (Table 3).
Structures of SET7/9 Y245A Bound to Unmodified and Meth-
ylated TAF10 Peptides—To elucidate the mechanism underly-
ing its unusual product specificity, we determined the crystal
structures of SET7/9 Y245A in complex with AdoHcy and
unmodified, mono-, di-, and trimethylated TAF10 peptides
(supplemental Table 1). These complexes superimpose with
the structure of the WT SET7/9AdoHcyTAF10-K189 com-
FIGURE 2. Crystal structures of WT SET7/9 and the Y305F mutant in complex with AdoHcy and unmodi-
fied and methylated TAF10 peptides. A, active site of WT SET7/9 bound to AdoHcy and TAF10-K189 and the
active site of SET7/9 Y305F bound to AdoHcy and TAF10-K189 (B), TAF10-K189me1 (C), and TAF10-K189me2
peptides (D). A–D, the K189 residue in the TAF10 peptide is depicted with yellow carbon atoms with the
corresponding Fo – Fc simulated-annealing omit maps (contoured at 2.5 ) shown. AdoHcy is colored with
green carbon atoms, and SET7/9 carbon atoms are colored gray, with the exception of Tyr-305/Y305F
(magenta) and Tyr-245 (cyan). Gly-264 to Ser-268 are shown, and the main chain atoms of Gly-292 to Ala-295
are depicted for clarity. Red dashed lines indicate conventional hydrogen bonds, and blue dashed lines indicate
CH–O hydrogen bonds. Cyan dashed lines indicate weak CH–O hydrogen bonds between 3.5 and 3.7 Å in
length. The water molecule in the solvent binding pocket is numbered 1. E, overlay of the active sites of the WT
enzyme (light blue carbon atoms) and the Y305F mutant (green, yellow, and pink carbon atoms correspond to
K189, K189me1, and K189me2, respectively). Red dashed lines indicate coordination of the waters in the struc-
turesoftheY305Fmutant,andthelightbluedashedlineindicateshydrogenbondingthatonlyoccursintheWT
enzyme. F, superimposition of the active sites of the SET7/9 Y305FAdoHcyTAF10-K189me2 and SET8
Y334FAdoHcyH4K20me2 (Protein Data Bank code 3F9X) complexes shown with green and yellow carbon
atoms, respectively.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31853
plex with root mean squared differences of less than 0.4 Å for all
aligned atoms, indicating that the Y245A mutant does not per-
turb the overall structure of the enzyme. Simulated annealing
omit maps illustrate that K189 side chains are bound within the
lysine binding channel through hydrogen bonds and van der
Waals contacts (Fig. 3, A–D), although the interactions and
binding modes are distinct from those in the complexes of WT
SET7/9 and the Y305F mutant (Fig. 2, A–D). In the unmodified
TAF10 peptide complex, the K189 -amino group forms a weak
hydrogen bond with the carbonyl oxygen of Gly-264 (Fig. 3A),
whereas the -amino groups of K189me1 and K189me2 hydro-
gen bond to the hydroxyl group of Tyr-305 in the mono- and
dimethylated peptide complexes (Fig. 3, B and C). The confor-
mations of the K189me1 and K189me2 side chains are further
stabilized by water-mediated hydrogen bonding and through
CH–O hydrogen bonding to their methyl groups. In the
TAF10-K189me3 peptide complex, the trimethyl-lysine side
chain is coordinated exclusively through direct and water-me-
diated CH–O hydrogen bonds to its methyl groups because the
quaternary -ammonium cation cannot engage in hydrogen
bonding (Fig. 3D).
A structural alignment of the four SET7/9 Y245A complexes
illustrates distinct binding modes for the unmodified versus the
methylated K189 side chains, highlighting the selectivity of this
mutant for methylated substrates. The side chains of K189me1,
K189me2, and K189me3 roughly overlay with their respective
-amino groups superimposed and adopt slightly kinked con-
formations (Fig. 3E), as opposed to the extended trans geome-
try of the unmodified and methylated lysines in the complexes
of the WT enzyme and the Y305F mutant (Fig. 2, A–D). Con-
versely, the unmodified K189 side chain does not superimpose
with its methylated counterparts and is oriented in an alterna-
tive configuration due to its hydrogen bonding to Gly-264 (Fig.
3, A and E). An overlay of the structures of the WT enzyme and
Y245A mutant bound to the unmodified TAF10 peptide illus-
trates that the side chains of K189 do not superimpose and that
the K189 -amino group appears to be misaligned with AdoHcy
in the Y245A complex (Fig. 3F). This suboptimal alignment
may explain the diminished kcat value of SET7/9 Y245A mutant
toward substrates with unmodified lysines (Table 2).
A comparison of the structures of the SET7/9 Y245A and
Y305F complexes yields a molecular explanation for the differ-
ent product specificities of these two mutants. In the SET7/9
Y305F complexes, Tyr-245 aligns the K189 -amino group for
methyl transfer through hydrogen bonding to its hydroxyl
group (Fig. 2, B–D). Conversely, in the Y245A mutant, the
K189me1 and K189me2 -amino groups are oriented through
hydrogen bonding to Tyr-305 (Fig. 3, B and C). These distinct
hydrogen bond patterns impart differences in the conforma-
tions of the lysyl side chains due to the relative orientations of
Tyr-245 and Tyr-305 in the lysine binding channel. Specifically,
the kinked conformation adopted by the K189me1 and
K189me2 side chains in the Y245A complexes (Fig. 3, B and C)
may contribute to the differences in the turnover numbers of
this mutant versus those of the WT enzyme and the Y305F
mutant (Table 2). In addition, the dimethyl -amino group of
the K189me2 side chain binds in distinct orientations in the
Y245A and Y305F mutants due to their hydrogen bonding to
FIGURE 3. Crystal structures of the SET7/9 Y245A mutant in complex with
AdoHcy and unmodified and methylated TAF10 peptides. Active site of
SET7/9 Y245A bound to AdoHcy and TAF10-K189 (A), TAF10-K189me1 (B),
TAF10-K189me2 (C), and TAF10-K189me3 peptides (D). A–D, Fo – Fc simulated-
annealing omit maps (contoured at 2.5 ) for the unmodified and methylated
K189 side chains are illustrated. The residues and hydrogen bonds in each com-
plexarecoloredasinFig.2.Thewatermoleculesinthelysinebindingchannelsof
the Y245A complexes are numbered 1–4, as described in the text. E, superimpo-
sition of the active sites of the Y245A complexes bound to the four methylated
statesofTAF10-K189(unmodified,mono-,di-,andtrimethylatedshowningreen,
yellow, pink, and blue, respectively). F, overlay of WT SET7/9 (blue carbons) and
SET7/9 Y245A (green carbons) in complex with TAF10-K189. Waters correspond-
ing to the WT and Y245A structures are colored cyan and green, respectively.
G, alignment of the active sites of the Y245A (yellow carbons) and Y305F (green
carbons) mutants in complex with the TAF10-K189me2 peptide. Hydrogen
bonds from the Y305F structure are shown as green dashed lines, and waters and
hydrogen bonds in the Y245A structure are shown in yellow and orange,
respectively.
Lysine Methylation by SET7/9 Mutants
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
Tyr-305 and Tyr-245, respectively (Fig. 3G). In the Y305F
mutant, hydrogen bonds to the dimethyl -amino group cou-
pled with steric constraints in the lysine binding channel pre-
vent the K189me2 side chain from undergoing a conforma-
tional change that is conducive to trimethylation (Fig. 2D),
consistent with its dimethyltransferase activity. However, in the
Y245A mutant, the alanine substitution enlarges the diameter
of the lysine binding channel, accommodating trimethyl-lysine
(Fig. 3D). In addition, the larger diameter would permit the
dimethyl-lysine substrate to undergo the conformational reor-
ganization necessary to align the -amino group in a productive
geometry for trimethylation.
A major difference in the active site of the Y245A mutant
versus the other SET7/9 structures is the presence of several
water molecules bound in the cavity generated by the Y245A
mutation. In the structure of the Y245A mutant bound to
TAF10-K189, three water molecules (waters 2–4) occupy this
cavity and are arranged in a triangular geometry (Fig. 3A). In
addition, water 1 shifts 1.6 Å from its position in the solvent
pocket toward water 2 to which it forms a hydrogen bond (Fig.
3, A and E). The shift in water 1 was unexpected given its con-
served orientation in the solvent pocket of the SET7/9 WT and
Y305F complexes (Fig. 2, A–C) as well as in the structures of
other SET domain KMTs (8). This displacement is presumably
related to the alternative conformation of the K189 side chain
whose -amino group is too distant (4.3 Å) to form a productive
hydrogen bond to water 1. Conversely, in the Y245A complexes
bound to TAF10-K189me1 and TAF10-K189me2, water 1
remains tightly bound in the solvent pocket through hydrogen
bonds to Tyr-305 hydroxyl group, the carbonyl oxygens of Gly-
292 and Ala-295, and the K189 -amino group (Fig. 3, B, C, and
E), analogous to its binding in the WT enzyme (Fig. 2A). How-
ever, in the TAF10-K189me3 complex, one of the methyl
groups of the trimethyl -ammonium cation is oriented into the
solvent pocket (Fig. 3D), similar to the dimethyl-lysine binding
mode observed in the Y305F mutant (Fig. 2D). The binding of
the methyl group in the solvent pocket displaces water 1 by 3.2
Å relative to its position in the TAF10-K189me1 complex (Fig.
3E), thereby avoiding a steric clash with the trimethylated
-ammonium group. Variations in the occupancy of water 2 are
also seen in the different Y245A structures. Water 2 is bound in
similar orientations in the active site of the unmodified and
monomethylated peptide complexes but is absent in the di- and
trimethylated peptide complexes due to the binding of a methyl
group in this position (Fig. 3, A–E). In summary, the changes in
the positions or occupancies of waters 1 and 2 correlate with the
binding modes of the unmodified and methylated K189 within
the active site of the Y245A mutant.
Catalytic Models of Lysine Multiple Methylation by SET7/9
Y245A, and Y305F—The structures of the SET7/9 complexes
reported here offer a prime opportunity to generate stepwise
models for lysine mono-, di-, and trimethylation by a SET
domain KMT. We modeled the AdoMet-bound Michaelis
complexes by superimposing the SET7/9 product complexes
with the previously reported structure of the SET7/9-AdoMet
binary complex (Fig. 4) (39). The conformations of the mono-
and dimethyl -amino groups in the Michaelis complexes were
inferred from the coordinates of the corresponding dimethyl-
and trimethyl-lysine products, respectively. In addition, we
modeled the -amino group in a deprotonated state with its
hydrogen atoms oriented toward the hydrogen bond acceptors
that align the lysyl side chain for methylation. As a basis for this
comparison, we first modeled the monomethylation reaction
catalyzed by WT SET7/9 (Fig. 4A). In the substrate ternary
complex, the lysine -amine is aligned with the methyl group
and sulfonium cation of AdoMet through a hydrogen bond to
the Tyr-245 hydroxyl group and water 1 in the solvent pocket.
The values of the reaction distance and angle are 2.8 Å and 153°,
respectively, in approximate agreement with the linear geome-
try of a SN2 methyl transfer reaction calculated in other mod-
eled substrate complexes (8, 10). In the product complex, the
monomethyl-lysine side chain is bound in an extended confor-
mation with its methyl group oriented within the methyl trans-
fer pore, thereby obstructing AdoMet binding. Furthermore,
water 1 remains tightly coordinated in the solvent pocket
through four hydrogen bonds to Gly-292, Ala-295, Tyr-305,
and the monomethyl -amino group. These interactions hinder
the dissociation of water 1 and the related rearrangement of the
monomethyl-lysine side chain required for a second methyl
transfer reaction, explaining why the WT enzyme cannot cata-
lyze di- and trimethylation. These findings concur with the
FIGURE 4. Catalytic models of the methyl transfer reactions catalyzed by
WT SET7/9 and the Y305F mutant. A, monomethylation of TAF10-K189 by
the WT enzyme. The reaction scheme depicts the modeled substrate ternary
complex (left) and the product complex (right) for the transfer of the methyl
group from AdoMet (green carbon atoms) to K189 in TAF10 (yellow carbons),
yielding AdoHcy and K189me1. The red arrow indicates the direction of the
nucleophilic attack of the deprotonated -amino group on the AdoMet
methyl group. The transferred methyl group is colored green, and the white
atoms represent the hydrogens of the -amino group. Hydrogen bonds
and residues in the enzyme active site are illustrated as in Fig. 2. The
reaction distance and angle are labeled in red. B and C, models of the
Y305F mutant for the first methyl transfer reaction with TAF10-K189 (B)
and second methyl transfer reaction with TAF10-K189me1 (C). Color
schemes are the same as in A.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31855
model for SET7/9 product specificity reported in previous
structural and functional studies (6, 11).
Similar reaction geometry is observed in the model for the
monomethyl transfer reaction catalyzed by SET7/9 Y305F.
Hydrogen bonds from the Tyr-245 hydroxyl group and water 1
align the lysine -amino group with the AdoMet methyl group
at a distance of 2.1 Å and an angle of 160°, equivalent to those
measured in the Michaelis complex of the WT enzyme (Fig. 4, A
and B). In the product complex, monomethyl-lysine adopts an
extended trans configuration analogous to that in the WT
enzyme. For dimethylation to occur, the monomethyl-lysine
must undergo a conformational change in which its methyl
group is rotated out of the methyl transfer path with AdoMet.
The structure of the Y305F mutant bound to the dimethylated
TAF10 peptide (Fig. 2D) implies that this rearrangement occurs
through the dissociation of water 1 due to the loss of the Tyr-
305 hydrogen bond in the solvent pocket. The dissociation of
water 1 would enable the monomethyl-lysine side chain to
adopt an alternative conformation through a rotation about its
C–N bond, projecting the methyl group into the solvent
pocket (Fig. 4C). This rotation reorients the methyl group out of
the methyl transfer path while realigning the monomethyl -
amino group for a second methylation reaction through a direct
hydrogen bond to the Tyr-245 hydroxyl group and a CH–O
hydrogen bond between its methyl group and Ala-295. The
modeled reaction geometry for monomethyl-lysine substrate
complex of the Y305F mutant (2.7 Å, 160°) is equivalent to that
of the first methyl transfer reaction in SET7/9 Y305F. These
geometries concur with our previous models for mono- and
dimethylation catalyzed by SET8 Y334F (8), illustrating that the
orientation of a methyl group into the solvent pocket is a con-
served feature of SET domain KMTs that catalyze multiple
methylation.
In addition, we modeled the methyl transfer reactions cata-
lyzed by SET7/9 Y245A (Fig. 5). In the model of the lysine sub-
strate complex, the -amino group is aligned for methyl transfer
by a hydrogen bond to the carbonyl oxygen of Gly-264, result-
ing in a short reaction distance (2.3 Å) and a suboptimal reac-
tion angle (141°) with the methyl group of AdoMet (Fig. 5A).
This misalignment appears to be a direct consequence of the
Y245A mutation that abolishes hydrogen bonding to the -
amino group, illustrating that the suboptimal orientation of the
-amine likely contributes to the diminished activity of this
mutant toward unmodified substrates (11). Conversely, in the
modeled monomethyl-lysine substrate complex for SET7/9
Y245A, Tyr-305 hydrogen bonds to the -amino group, aligning
it for methyl transfer (Fig. 5B). In addition, CH–O hydrogen
bonds to the monomethyl-lysine methyl group and the dissoci-
ation of water 2 from the active site also contribute to reposition-
ing the -amino group for dimethylation. Collectively, these inter-
actions orient the -amine in a reaction angle of 165° that is more
conducive to methyl transfer. However, the reaction distance for
dimethylation is 0.6 Å longer than that in the corresponding
Y305F model because Tyr-305 is positioned further from
AdoMet than Tyr-245 (Figs. 4C and 5B).
In the third methyl transfer reaction catalyzed by SET7/9
Y245A, the lone pair of electrons of the dimethyl-lysine -
amino group acts as the nucleophile and thus cannot engage in
hydrogen bonding. The structure of the trimethyl-lysine prod-
uct complex (Fig. 3D) implies that the dimethyl -amine is
aligned via CH–O hydrogen bonds to its methyl groups, as
shown in the model of the Michaelis complex for this reaction
(Fig. 5C). These CH–O hydrogen bonds restrain the orienta-
tion of the -amino group and position one of the methyl
groups into the solvent pocket, displacing water 1 as discussed
earlier (Fig. 3, D and E). These interactions cumulatively align
the -amino group and AdoMet methyl group with a reaction
distance of 3.0 Å and angle of 162° (Fig. 5C). Taken together, the
models of the substrate complexes for SET7/9 Y245A suggest
that CH–O hydrogen bonds play an increasingly important role
in aligning the methylated -amino group in successive rounds
of methyl transfer.
DISCUSSION
The structural and functional characterization of the
SET7/9 Y245A and Y305F mutants presented here yields
new insights into the mechanism underlying the product
specificity of SET domain KMTs. Importantly, it resolves a
general paradox concerning this specificity. How does the
active site constrain the motion of the lysine -amino group
to align it for methyl transfer with AdoMet, while providing
adequate volume to accommodate the mono-, di-, and tri-
methylated lysine side chain generated during multiple
methyl transfer reactions? The structures of the Y305F and
Y245A mutants resolve this paradox, illustrating that alter-
ations in the positions or occupancies of water molecules
within their active sites generate the space required to
FIGURE 5. Models for the methyl transfer reactions catalyzed by the
SET7/9 Y245A mutant. Models of the Y245A mutant for the first methyl
transfer reaction with TAF10-K189 (A), the second methyl transfer reaction
with TAF10-K189me1 (B), and the third methyl transfer reaction with TAF10-
K189me2 (C). Residues and hydrogen bonds are depicted as in Fig. 3.
Lysine Methylation by SET7/9 Mutants
31856
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 41•OCTOBER 8, 2010
accommodate the multiply methylated -amine produced
during successive catalytic cycles. Minor perturbations in
the side chains of certain active site residues, such as Tyr-
305, are also observed in alignments of the WT enzyme and
the Y245A and Y305F complexes, although these changes are
modest compared with the displacement or dissociation of
the water molecules in the active site. These findings suggest
that the waters function as transient place holders that facil-
itate the SN2 methyl transfer reaction. During monomethy-
lation, they function to constrain the movement of the lysine
-amino group by mediating hydrogen bonds between the
substrate and enzyme, thereby promoting the linear align-
ment with the methyl group and sulfonium cation of
AdoMet (Fig. 4, A and B). During di- and trimethylation, the
water molecules either relocate within the lysine binding
channel or dissociate from the enzyme, yielding the space
required to rotate the methyl group away from the methyl
transfer pore and to realign the -amine in productive geom-
etry for the next methyl transfer reaction (Figs. 4C and 5, B
and C). These findings agree with our prior analysis of the
SET8 Phe/Tyr switch mutant in which we demonstrated that
the Y334F substitution attenuates hydrogen bonding to the
water molecule bound in the solvent pocket, promoting its
dissociation and the conformational changes necessary for
lysine dimethylation (8). Indeed, there is a nearly identical
alignment of the dimethyl-lysine side chains in the structures of
SET7/9Y305FandSET8Y334Fcomplexes,despitethedifferences
in the orientations of the Phe-305 and Phe-334 side chains in each
structure (Fig. 2F). Finally, both the SET8 Y334F (8) and SET7/9
Y305F mutants (Table 2) displayed diminished catalytic efficien-
cies for lysine dimethylation versus monomethylation. These dif-
ferences may reflect the kinetics of the reorganization within the
active site, including the dissociation of the water molecule from
the solvent pocket and the concomitant realignment of the
monomethyl-lysine into a productive geometry for dimethylation.
In addition to their place-holding role, the active site waters
may also facilitate the deprotonation of the lysine -amino
group between methyl transfer reactions. For methylation to
occur, the -amino group must be deprotonated to function as
the nucleophile in the SN2 methyl transfer reaction with
AdoMet (Figs. 4 and 5). Although the pKa value of the lysine
-amine in solution is 10.5, molecular dynamics simulations
by Zhang and Bruice (25, 26) indicate that this value diminishes
to 8.2 upon formation of the SET7/9 Michaelis complex due to
the proximity of the AdoMet sulfonium cation and the low
dielectric constant of the active site. Furthermore, their simu-
lations show that a chain of water molecules facilitates the dep-
rotonation of the -amino group prior to methyl transfer, trans-
ferring the proton to bulk solvent. Although these water
molecule chains are not evident in our crystal structures, the
Y305F and Y245A complexes suggest another potential mech-
anism for deprotonation. In the dimethyl-lysine complexes of
the Y305F and Y245A mutants, the dissociation of water 1 and
2, respectively, from the lysine binding channel requires that
the solvent-mediated hydrogen bond to the -amino group is
broken (Figs. 2D and 3C). It is conceivable that these waters
dissociate from the active site as hydronium ions, promoting
the realignment and deprotonation of the methyl -amino
group for the next methyl transfer reaction.
A comparison of the SET7/9 Y305F and SET8 Y334F com-
plexes yields insights into the mechanism by which the Phe/Tyr
switch influences water binding within the solvent pocket. The
phenylalanine substitution in the Phe/Tyr switch results in the
loss of a single hydrogen bond to the water molecule (water 1) in
the solvent pocket compared with the four hydrogen bonds that
coordinate the solvent molecule in WT SET7/9 (Fig. 2, A and B)
and SET8 (7, 8). Although this attenuation in hydrogen bonding
may appear insignificant, this difference is nonetheless impor-
tant for at least two reasons. First, theoretical calculations indi-
cate that, on average, water molecules form 3.5 hydrogen
bonds in solutions (40, 41). This value is greater than the num-
ber of hydrogen bonds coordinating water 1 in the solvent
pocket in SET7/9 Y305F (Fig. 2, B and C) as well as in SET8
Y334F and other di- and trimethyltransferases that possess a
hydrophobic residue in the Phe/Tyr switch site (8). From the
perspective of the water molecule, the greater hydrogen bond-
ing potential in solution would tend to thermodynamically
favor its dissociation from the solvent pocket in SET domain
KMTs that lack a tyrosine in the Phe/Tyr switch position. Sec-
ond, the ordered binding of water molecules observed in the
active sites of SET domain ternary complexes represents an
unfavorable entropy compared with their diffusion in bulk sol-
vent. In WT SET7/9 (Fig. 2A) and SET8 (7, 8), this entropic
penalty can be partially offset through the favorable enthalpy of
binding associated with the four hydrogen bonds that coordi-
nate the water within the solvent pocket. It is conceivable that
the loss of the tyrosine-mediated hydrogen bond in the Phe/Tyr
switch shifts the equilibrium in favor of dissociation of the
water molecule from the solvent pocket, thereby facilitating
dimethylation in SET7/9 Y305F, SET8 Y334F, and other di- and
trimethyltransferases.
The structures of the SET7/9 Y245A and Y305F complexes
illustrate the interactions that align the lysine -amino group
during the methyl transfer reactions in each enzyme. In the WT
enzyme and the Y305F mutant, hydrogen bonding to the
hydroxyl group of Tyr-245 appears to be critical in properly
aligning the -amine for methyl transfer (Fig. 4). Tyr-245 is
conserved in the sequences of many SET domain KMTs (8, 42),
and substitutions of this residue generally impair or abolish
activity, indicating its importance in catalysis (8, 43). However,
SET7/9 appears to be an exception to this rule, as the Y245A
mutant is not only active but is capable of catalyzing lysine
trimethylation. In this mutant, Tyr-305 appears to assume the
role of Tyr-245 by hydrogen bonding to the monomethylated
-amino group to align it for methyl transfer with AdoMet, as
illustrated in the modeled substrate complex for the dimethy-
lation reaction (Fig. 5B). Conversely, in the model for trimethy-
lation, the Tyr-305 hydroxyl group does not hydrogen bond to
the -amine but instead participates in a CH–O hydrogen bond
with one of the methyl groups to assist in aligning the dimethy-
lated -amine for the methyl transfer reaction (Fig. 5C). Addi-
tional structural and functional studies of the SET domain tri-
methyltransferases will aid in further illuminating the roles
of CH–O hydrogen bonds in facilitating lysine multiple
methylation.
Lysine Methylation by SET7/9 Mutants
OCTOBER 8, 2010•VOLUME 285•NUMBER 41
JOURNAL OF BIOLOGICAL CHEMISTRY 31857
Acknowledgments—We acknowledge S. Schiebold for assistance in
protein expression, purification, and crystallization and S. Anderson
and R. Sanishvili for their assistance with x-ray data collection. We
also thank S. Bulfer and S. Horowitz for reading the manuscript and
providing useful comments. This work utilized the Protein Structure
Facility of the Michigan Diabetes Research and Training Center, Uni-
versity of Michigan, supported by National Institutes of Health Grant
DK020572, NIDDK. Use of the Advanced Photon Source was sup-
ported by the United States Department of Energy, Basic Energy Sci-
ences, Office of Science, under Contract DE-AC02-06CH11357.
GM/CA CAT has been funded in whole or in part by National
Institutes of Health NCI Grant Y1-CO-1020 and NIGMS Grant
Y1-GM-1104. Use of the LS-CAT Sector 21 was supported by Michi-
gan Economic Development Corporation and the Michigan Technol-
ogy Tri-Corridor Grant 085P1000817 for the support of this research
program.
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Lysine Methylation by SET7/9 Mutants
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VOLUME 285•NUMBER 41•OCTOBER 8, 2010
|
3M5C
|
Crystal structure of N-acetyl-L-ornithine transcarbamylase K302E mutant complexed with PALAO
|
Reversible Post-Translational Carboxylation Modulates The
Enzymatic Activity Of N-Acetyl-L-Ornithine Transcarbamylase†
Yongdong Li1,2, Xiaolin Yu1, Jeremy Ho1, David Fushman3, Norma M. Allewell3, Mendel
Tuchman1, and Dashuang Shi1,‡
1Research Center for Genetic Medicine and Department of Integrative Systems Biology,
Children’s National Medical Center, The George Washington University, Washington, DC 20010,
USA.
2Key Laboratory of Organo-Pharmaceutical Chemistry, Jiangxi Province, Gannan Normal
University, Ganzhou 341000, China.
3Department of Chemistry and Biochemistry, College of Chemical and Life Sciences, University of
Maryland, College Park, MD 20742, USA.
Abstract
N-acetyl-L-ornithine transcarbamylase (AOTCase), rather than ornithine transcarbamylase
(OTCase), is the essential carbamylase enzyme in the arginine biosynthesis of several plant and
human pathogens. The specificity of this unique enzyme provides a potential target for controlling
the spread of these pathogens. Recently, several crystal structures of AOTCase from Xanthomonas
campestris (xc) have been determined. In these structures, an unexplained electron density at the
tip of Lys302 side-chain was observed. Using 13C NMR spectroscopy, we show herein that
Lys302 is post-translationally carboxylated. The structure of wild-type AOTCase complexed with
the bisubstrate analogue, Nδ-(phosphonoacetyl)-Nα-acetyl-L-ornithine (PALAO), indicates that the
carboxyl group on Lys302 forms a strong hydrogen bonding network with surrounding active site
residues, Lys252, Ser253, His293, and Glu92 from the adjacent subunit either directly or via a
water molecule. Furthermore, the carboxyl group is involved in binding N-acetyl-L-ornithine via a
water molecule. Activity assays with the wild-type enzyme and several mutants demonstrate that
the post translational modification of lysine 302 has an important role in catalysis.
Post-translational modification of the ε-amino group of lysine residues in proteins is a
common mechanism used by organisms to regulate protein functions including DNA-protein
interactions, subcellular localization, transcriptional activity, and protein stability and
activity (1). Lysine residues can be modified by the addition of functional groups to become
acetylated, methylated, carbamylated or carboxylated. The role of histone lysine acetylation
and methylation in affecting chromatin structure and gene expression has been well
established for more than a decade (2). However, the biological roles for lysine
carbamylation and carboxylation have rarely been investigated.
†This work was supported by Public Health Service grants DK-47870 (MT) and DK-067935 (DS) from the National Institute of
Diabetes, Digestive and Kidney Diseases. JH was supported by a Scholarship from the Doug and Lynn Parsons Family Foundation.
The Cornell High Energy Synchrotron Source (CHESS) is supported by the National Science Foundation under award DMR 0225180
and the Macromolecular Diffraction Facility at CHESS (MacCHESS) is supported by award RR-01646 from the National Institutes of
Health, through its National Center for Research Resources.
‡Corresponding author. dshi@cnmcresearch.org. Phone: 202-476-5817. Fax: 202-476-6014.
SUPPORTING INFORMATION AVAILABLE
Figure S1. Structure and hydrogen bonding network around residue 302 for previously determined AOTCase structures. This material
is available free of charge via the Internet at http://pubs.acs.org.
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 August 17.
Published in final edited form as:
Biochemistry. 2010 August 17; 49(32): 6887–6895. doi:10.1021/bi1007386.
NIH-PA Author Manuscript
NIH-PA Author Manuscript
NIH-PA Author Manuscript
In vivo, lysine acetylation and methylation are usually carried out by acetyltransferase and
methyltransferase enzymes, respectively (3). In addition, some proteins such as hemoglobin
and human serum albumin can be acetylated non-enzymatically by chemicals such as
aspirin, methyl acetyl phosphate, and other acetylating agents such as acetyl-CoA (4–8).
Lysine can also be methylated by small chemicals in vitro, and this has routinely been used
as a rescue method for protein crystallization (9). Lysine carbamylation and lysine
carboxylation have only been achieved by using chemicals and no enzyme has yet been
found to catalyze these modifications. Lysine carbamylation was one of the earliest post-
translational modification of proteins to be elucidated when it was identified as a product of
reversible denaturation-renaturation studies of proteins with urea (10,11). This
carbamylation, which produces homocitrulline, has also been detected in uremic patients
(12) and in patients with elevated plasma and/or urinary lysine levels (13). In contrast, lysine
carboxylation is not as commonly reported, but has been identified in a number of proteins
via crystal structure determinations. In most of these proteins, the carboxyl groups of
modified lysines are involved in bridging two metal ions that play a structural role in the
active site. In several other proteins, however, a direct role for a carboxylated lysine in the
catalytic mechanism has been reported (14–16).
N-acetyl-L-ornithine transcarbamylase (AOTCase, EC 2.1.3.9) was recently discovered to
be part of a novel arginine biosynthesis pathway in plant pathogens of the
Xanthomonadaceae family such as Xylella and Xanthomonas (17–19). These pathogens
attack a variety of economically important crops including citrus fruits, cotton, tomatoes,
and rice (20,21). Genome sequence analyses showed that an AOTCase-like gene is also
present in some human pathogens such as Stenotrophomonas maltophilia and members of
the genus Bacteroides (22). In the case of Bacteroides fragilis, this gene was later confirmed
to encode another novel transcarbamylase, N-succinyl-L-ornithine transcarbamylase
(SOTCase, EC 2.1.3.11) (23). Crystal structures of both AOTCase and SOTCase bound with
substrate or substrate analogues have recently been determined (17,18,23). An extended
density at the side-chain tip of Lys302 in AOTCase was observed suggesting a post-
translational modification. Since Lys302 is located within the active site of AOTCase and is
not present in SOTCase, it was proposed as one of three key signature residues to
distinguish the two carbamylases (22). Here, we demonstrate that Lys302 is post-
translationally modified by carboxylation and that this change affects the catalytic function
of the enzyme.
MATERIALS AND METHODS
Materials
All chemicals were purchased from Sigma Chemical Company unless otherwise specified.
ANOR was purchased from Indofine Chemical Co., Inc. N-acetyl-L-citrulline was custom
synthesized and purified by Chiral Quest Company. PALAO (>95% purity) was synthesized
by IMI TAMI Institute of Research and Development Ltd. (19). xcAOTCase was prepared
and purified as previously described (18). Mutants K302A (primer: 5’-
CTGCGTCGCAACGTCGCGGCTACTGATGCGGTG-3’), K302E (primer:5’-
CTGCGTCGCAACGTCGAGGCTACTGATGCGGTG-3’) and K302R (primer: 5’-
CTGCGTCGCAACGTCAGGGCTACTGATGCGGTG-3’) were generated by site-directed
mutagenesis using the “Quik Change” mutagenesis kit (Stratagene) according to the
manufacturer’s protocol. The correct mutants were confirmed by DNA sequencing.
Recombinant mutant proteins were expressed and purified in the same manner as the wild-
type enzyme.
Li et al.
Page 2
Biochemistry. Author manuscript; available in PMC 2011 August 17.
NIH-PA Author Manuscript
NIH-PA Author Manuscript
NIH-PA Author Manuscript
Activity assay
The modified colorimetric assay method, which detects the formation of the ureido group
during the transcarbamylation reaction (24), was used to measure enzyme activity. CP and
AORN concentration were kept constant as 4.0 mM. After an incubation of 5 minutes, the
reaction was stopped by the addition of 1 ml of color reagent, as described previously (19).
A set of tubes containing known amounts of N-acetyl-L-citrulline was included with each
rack of enzyme assays to produce a standard curve for calculation of the enzyme specific
activity.
Mass spectrometric analysis
In order to identify the post-translational modification, mass spectrometric analysis was
carried out on a 4700 ABI TOF/TOF mass spectrometer (Applier Biosystems) based on the
method described previously (25). In brief, 10 µg of native protein were digested overnight
at 310 K using trypsin in 50 mM ammonium bicarbonate pH 7.4. After desalting using a
C18 ZipTip micropipette tip, the resulting peptides were eluted in 10 µl of acetonitrile/0.1%
TFA [70:30(v:v)]. The sample was mixed with matrix solution and spotted on a MALDI
plate to be submitted to the mass spectrometric analysis.
Chemical rescue experiments
The assay in the presence of various selected chemical was conducted as described above.
The stock solutions of small chemicals were titrated to the pH of the assay with KOH or
HCl.
13C NMR experiments
The wild-type and K302A mutant protein of AOTCase (~10 mg) was precipitated by
degassed buffer (pH 4.5) containing 25 mM sodium acetate. After centrifugation, the
precipitate was re-dissolved by adding a buffer containing 20 mM NaH13CO3, 100 mM Tris
HCl (pH 8.0) and 50 mM NaCl. Before NMR experiments, 40 µl D2O was added to 500 µl
protein sample. The 13C NMR spectra were collected on a Bruker Avance 600 spectrometer
(operating at 14.1 T) equipped with a direct 13C-detection probe at 298 K. The experimental
settings and processing parameters for the wild-type protein and K302A variant were
identical. 512 transients were collected with 4K time domain points and a spectral width of
3019 Hz centered at 160 ppm. The spectra were processed using exponential multiplication
with the line broadening factor set to 3Hz. The similarity of protein concentration in both
samples was verified by 1H NMR (not shown).
Crystallization, data collection and processing
PALAO-bound wild-type and mutant AOTCase crystals were grown using the hanging-drop
vapor diffusion method, with conditions similar to those used to produce native and ligand-
complexed AOTCase crystals (18,23). 2.0 µl of ~10 mg/ml solution of AOTCase were
mixed with 1.6 µl of reservoir solution and 0.4 µl PALAO solution (~0.01 M). The reservoir
solution contained 20% (w/v) PEG 3350, 0.2 M lithium sulfate and 0.1 M bis-Tris, pH 6.0.
Diffraction data for the PALAO-bound crystal were collected at 100 K at the F1 beam line
of the Cornell High Energy Synchrotron Source. Data sets for the PALAO-bound mutant
AOTCase crystals were obtained using a Rigaku anode x-ray generator in the Molecular
Structure Section of the National Institute of Health. All data were processed using
HKL2000 package (26) and reduced using the program TRUNCATE in the CCP4 suite (27).
Data collection parameters are listed in Table 1.
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Structure solution and refinement
Molecular replacement was used for phase determination of the PALAO-bound wild-type
and mutant AOTCase structures. The coordinates of AOTCase (PDB code: 3KZO) after
removal of ligands or water molecules were used for phase determination. Upon rigid-body
refinement, electron density corresponding to the ligands could be clearly visualized. The
ligands were built into the map using the graphics program O (28). Refinements were
carried out using molecular annealing, energy minimization and restrained B factor
refinement with the program CNS1.1 (29). During refinements, 5% of the reflections at
various resolutions were randomly selected to set aside to calculate Rfree to monitor the
progress of refinement (30). After every cycle of refinement, the model was manually
adjusted using the program O (28). Water molecules were automatically assigned using the
program WATERPICK of CNS. Model quality was checked using the program
PROCHECK (31) to ensure good stereochemistry for all three models. The final refinement
statistics are listed in Table 1.
Figures 1 was drawn using the programs Pymol (http://www.pymol.org). Figure 3 was
drawn using ChemDraw 8.0. The coordinates have been deposited with the RCSB PDB as
entries 3M4J, 3M4N, 3M5C and 3M5D.
RESULTS
Lys302 in AOTCase is carboxylated
To investigate the nature of the modification of Lys302 and how it affects catalytic activity,
we revisited all AOTCase structures. In the PALAO-bound AOTCase structure, the electron
density map clearly indicates that Lys302 is post-translationally modified (Figure 1A). The
type of modification can include methylation, acetylation, carbamylation, and carboxylation.
The shape of the electron density can been used to distinguish methyl groups from larger
functional groups, but it is difficult to distinguish between acetyl, carbamyl, and carboxyl
groups, all of which have three non-hydrogen atoms in a plane. Given the hydrogen bonding
network with surrounding residues (Lys252, Ser253 and His293, Table 2), a carboxylated
modification is the most likely choice for the modification of Lys302 in AOTCase. To
exclude that the modification’s identity represents chemically stable moieties (methyl,
acetyl, carbamyl), we analyzed trypsin digested fragments of purified AOTCase by TOF-
TOF mass spectroscopy. As expected, only a peptide fragment with an unmodified Lys302
was observed, consistent with the lability of the carboxylic group in acidic solutions. At low
pH, the carboxyl group is spontaneously released as carbon dioxide (14, 32), in contrast to
other modified groups that are stably bound and can be observed by mass spectrometry
analysis after proteolysis (33).
The putative carboxyl group on the modified Lys302 forms direct hydrogen bonds with
main-chain or side-chain nitrogen atoms of Lys252, Ser253 and His293 (Figure 1A and
Table 2). Among these, Lys252 is involved in direct hydrogen bonding to the carboxyl
group of the AORN moiety of PALAO, and His293 forms a strong hydrogen bond with the
main-chain nitrogen atom of Leu295 in the conserved His293-Cys294-Leu295-Pro206
(HCLP) motif. The hydrogen bonding network between the carboxyl group of modified
Lys302, His293 and the main-chain nitrogen atom of Leu295 is reminiscent of the similar
hydrogen bonding network, Glu310-His302-Leu304 and Glu299-Leu272-Leu274, found in
human and E. coli OTCase, respectively (34, 35). These three residues are conserved in all
OTCase sequences, and the interactions between them are important for maintaining the
HCLP motif in a specific conformation to orientate their main-chain oxygen atoms towards
the active site. In all known transcarbamylase structures, a leucine residue corresponding to
Leu295 is in an energetically unfavorable conformation and the peptide bond between this
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leucine and Pro296 is in the cis conformation. In addition to the direct hydrogen bonding
interaction above, the carboxyl group on the modified Lys302 interacts with the α-amino
nitrogen atom of the AORN moiety of PALAO and Glu92 from the adjacent subunit via
water molecules.
When we revisited all previously determined AOTCase structures (see supplementary
Figure S1) we found: (1) Lys302 was carboxylated in the absence of substrate binding, but
substrate binding immobilizes the side-chain of Lys302 further by hydrogen bonding
interaction via water molecules. (2) Water-mediated hydrogen bonding promotes
interactions of carboxylated Lys302 with Glu92 from the adjacent subunit and the α-amino
nitrogen atom of AORN. (3) Similarly to AOTCase, in the structure of SOTCase E92Z (Z =
Ala, Ser, Pro, Val), mutant with N-succinyl-L-norvaline bound (22) the carboxylated Lys302
hydrogen bonds to the α-amino nitrogen atom and the succinyl carboxyl group of N-
succinyl-L-norvaline via water molecules (Figure S1).
To obtain direct, independent evidence for the carboxylation of Lys302, 13C NMR
experiments were carried out with both wild-type protein and the K302A mutant. As
observed for other proteins with carboxylated lysine (36,37), the strong 13C NMR signal at
164 ppm characteristic of a carboxyl group was clearly detectable in AOTCase wild-type
protein labeled by 13C-bicarbonate, in contrast to the K302A mutant where the signal was
weak (Figure 2). Since there are 17 other lysine residues in the protein, the weak signal seen
for the K302A mutant might be caused by the adventitious carboxylation of another lysine
with reduced pKa, as has been observed for the K392A mutant of the sensor domain of the
BlaR protein (38).
Functional and structural studies of Lys302 mutants
To investigate the effect of lysine carboxylation on enzyme activity, Lys302 was mutated to
alanine, glutamate or arginine. Each of these variants was expressed in E. coli and gave
similar yields. Enzymatic assays demonstrated a significant decrease in enzymatic activity in
all three mutants, reflecting the functional importance of Lys302 (Table 3). The level of
enzymatic activity for the wild-type (WT) and three mutants was WT > K302A > K302E ≫
K302R. To determine the structural basis of these results, the WT and mutant enzymes
bound with PALAO were crystallized and their structures were determined at 1.8–2.2 Å
resolution. Only the K302R mutation had and appreciable effect on the structure of the
protein. Since K302 is located near the AORN binding site, the mutations would weaken
AORN binding to the active site.
In the structure of the K302A mutant, three additional water molecules (labeled as w3, w4
and w5 in Figure 1B) replace the carboxylated lysine. The two water molecules (labeled w1
and w2 in Figure 1A–1D) that mediate the hydrogen bonding interaction of carboxylated
Lys302 with PALAO and Glu92 from the adjacent subunit are also found in the K302A
mutant structure. Furthermore, these water molecules maintain a similar hydrogen-bonding
network to the wild-type enzyme. These results might explain why the K302A mutant
retains significant catalytic activity (Table 3). To investigate whether adding short-chain
carboxylic acids to the K302A mutant increases its activity as other enzymes (14, 15, 39,
40), the activity of the K302A mutant was measured in the presence of high formate and
acetate concentration (0.5 M). Surprisingly, the activity of the K302A mutant was not
significantly improved. The crystal structure of the K302A mutant soaking with the
crystallization buffer in the presence of 0.5 M acetate was also determined (not shown) and
it was observed that the same five water molecules were present in the cavity that replaced
the side chain of the carboxylated lysine. This, the acetate’s inability to replace the water
molecules in the crystal structure, is consistent with the unchanged activity assay results.
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The side-chain of Glu302 in the K302E mutant structure is well defined and anchored by
hydrogen bonding interaction with the main-chain nitrogen atom of Arg298 and weakly
hydrogen bonded to the main-chain nitrogen atom of Ser253 (Figure 1C). Two of three
additional water molecules (w4 and w5) observed in the K302A mutant structure occupied
the same position as the carboxyl oxygen atoms of Glu302 and form a similar hydrogen-
bonding network. Relative to the PALAO-bound wild-type structure, there is only one more
water molecule (w3) at the position of the carboxyl group of the carboxylated Lys302. This
water molecule mediates a hydrogen bonding interaction between Glu302 and Lys252. Two
common water molecules (w1 and w2) that interact with PALAO and Glu92 from the
adjacent subunit, respectively, were also identified in the K302E structure.
Our observation that the K302E mutant had lower enzymatic activity than that of the K302A
mutant (Table 3) was surprising since the carboxyl group of the glutamate could conceivably
function similarly to a carboxylated lysine. The explanation may be that, in the K302A
mutant, the hydrogen bonding network is well maintained by water molecules in the cavity
that replaces the carboxylated lysine. In particular, w3 is optimally located for strong
hydrogen bonding to w1 (2.7 Å), which in turn binds AORN. The distances between w1 and
the carboxyl oxygen of carboxylated Lys302 in all wild-type crystal structures are within
2.4–2.7 Å, but the distance between w1 and w2 in the K302E structure is significantly
greater (3.2 Å). The weaker hydrogen bonding interaction may be a reason for lower
enzymatic activity of the K302E mutant.
In contrast to the K302A and K302E structures, the K302R structure shows a much larger
reduction in enzyme activity relative to the wild-type enzyme. The electron density for the
side-chain of Arg302 is weak and the temperature factor of its side-chain is 54.4 Å2,
significantly higher than those of carboxylated Lys302 (44.7 Å2) and Glu302 (33.4 Å2),
implying greater flexibility. Furthermore, the side-chain of Arg302 is oriented differently
from the carboxyl group of carboxylated Lys302 and pushes the nearby residues His180,
Pro181 and Lys182 outwards about 1.0 Å (Figure 1D). However, the water molecules
involved in hydrogen bonding to the α-amino nitrogen atom of PALAO (w1) and the side-
chains of Lys252 (w3) and Glu92 (w2) from the adjacent subunit are conserved. Consistent
with the K302E structure, the distance between w1 and w2 is even greater (3.4 Å) than in
the WT structure and the hydrogen bonding interaction between w2 and w3 is no longer
observed. Thus, the almost undetectable enzymatic activity of the K302R mutant probably
results from the changes at its active site, including the weakened hydrogen bonding
network involved in substrate binding.
DISCUSSION
Several lines of evidence clearly indicate that Lys302 in AOTCase is carboxylated. First, the
extra electron density indicates that the side-chain of Lys302 is modified. Second, the
hydrogen bonding environment of Lys302 for hydrogen bonding interactions is compatible
with a carboxyl group, but not for a positively charged lysine side-chain. Third, the
modification is labile at low pH, since mass spectroscopy of samples prepared at low pH
indicated that Lys302 was no longer modified. Fourth, the clear presence of the
indicative 13C NMR signal at 164 ppm for wild-type protein and its absence in the K302A
mutant confirms carboxylation of Lys302.
It is well known that lysine carboxylation is non-enzymatic and reversible, while other post-
translational modifications such as methylation, acetylation, and carbamylation are
irreversible and detectable by mass spectroscopy. Furthermore, lysine methylation and
acetylation usually require an enzyme-catalyzed reaction in vivo (41). Therefore, it is
unlikely that such lysine modifications will be observed in recombinant proteins
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overexpressed in a foreign host (e.g. E. coli). Lysine methylation can be achieved by using
special chemicals in vitro, but these chemicals are not present in vivo. Lysine carbamylation
and carboxylation use completely different mechanisms to form functionally different
groups (Figure 3). Carbamylation can be achieved by cyanate produced from
myeloperoxidase-catalyzed oxidation of thiocyanate, an anion abundant in plasma and
increased in smokers, or from urea in the plasma. Lysine carboxylation, on the other hand,
occurs readily in aqueous solution in the presence of carbon dioxide at a basic pH (32,42).
Even though carbamylation and carboxylation use very different mechanisms, the two are
confused in the literatures. Lysine carbamylation (or carbamoylation) is referred to in
several publications (15,32,42–44), when the actual reaction is in fact carboxylation.
The activity of the K302A mutant is almost half of that of the wild-type enzyme raising the
question of why AOTCase retains a lysine in this position. Perhaps this lysine was
maintained through evolution to distinguish AOTCase from SOTCase which uses N-
succinyl-L-ornithine (SORN) rather than AORN (22), and OTCase which uses L-ornithine.
An alternative explanation may be found in the very low activity of the K302R mutant. The
side-chain of arginine has a positive charge while carboxylated lysine has a negative charge.
The side chain of unmodified lysine is usually located in a similar position as that of
arginine, as observed in the structure of UV damage endonuclease (14). It would be
expected that the activity of AOTCase with an uncarboxylated lysine would be as low as the
K302R mutant’s. It could further be surmised that, the respective organisms need to use
carboxylation as a switch to turn “on” or “off” the arginine biosynthetic pathway. It has been
well known that rubulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) in plant cells
uses the carboxylation on Lys201 as a switch to turn the enzyme “on” during the day and
“off” at night by removing the carboxyl group (45,46). Carbon dioxide and bicarbonate have
been found to play an important biological role in modulating several biological processes
including photosynthetic carbon fixation (47), pH homeostasis (48), carbon metabolism
(49), activation of virulence in pathogenic organisms (50), sperm maturation (51),
stimulation of mammalian G-protein-responsive adenylyl cyclase (52), and as an alarmone
in Drosophila (53,54). Whether or not carboxylation of a key lysine in their related proteins
is used as an underlying regulatory mechanism should be investigated further.
There are 197 structures with carboxylated lysine residue (modified residue indicated as
Kcx) in the Protein Data Bank (PDB). If structures with 90% identity are counted only once,
there are still 52 unique structures remaining in this pool (Table 4). These proteins include
hydantoinase (40,55), folylpolyglutamate synthase (43), UV damage endonuclease (14),
OXA10, OXA-1 class D β-lactamase (38,56,57), urease (42), phosphotriesterase (58),
dihydroorotase (59), dihydropyrimidinase (60), organophosphate hydrolase (61) and MurE
and MurD ligases (44,62). In most of these proteins, the carboxylated lysine bridges two
metal ions, similar to the role of glutamate or aspartate in proteins with two metal-binding
sites (26 structures among 52). However, the urease apoenzyme can be activated in vitro
only in the presence of carbon dioxide prior to nickel binding (63), suggesting that the
carboxylated lysine may have other structural roles beyond binding metals. In some proteins
such as β-lactamase, UV damage endonuclease, Rubisco, MurD and MurE ligase and BlaR
signal transducer protein, a carboxylated lysine plays an essential catalytic role. More
interestingly, in three structures (PDB ID: 1HL9, 1PU6 and 2UYN for fucosiadase, 3-
methyladenine DNA glycosylase and TdcF protein of unknown function, respectively), the
carboxylated lysines are located near the surface of proteins, presumably playing primarily a
structure stabilizing role (64–66). Since the carboxyl group is labile at acidic pH, but easily
formed in the presence of carbon dioxide at basic pH, the number of proteins with lysine
carboxylation must be underestimated. Furthermore, the carboxylated lysine must be fixed
in place by metal ions (either one or two) or hydrogen bonding with other protein residues
(at least one). Therefore, any detection method involving denaturing the proteins will result
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in release of the carboxyl group. With current technology, 13C NMR (38) and
crystallography are the only methods that can detect this modification. However, these
methods are not amenable to high-throughput investigations. The majority (49 out of 52
structures in the PDB) of known lysine carboxylation modifications were found to be
located at or near the active site, probably because these sites receive the most attention.
Revisiting the structures in PDB with more attention to surface lysines might reveal more
structures with carboxylated lysines.
In conclusion, we have shown that Lys302 in AOTCase is post-translationally modified by
carboxylation and that this modification may be functionally important for enzymatic
activity. Lysine carboxylation is likely to be a more common event than currently
appreciated and may play a critical role in enzymatic activity and protein stability.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Abbreviations
ACIT
N-acteyl-L-citrulline
ANOR
N-acetyl-L-norvaline
AORN
N-acetyl-L-Ornithine
AOTCase
N-acetyl-L-ornithine transcarbamylase
ATCase
aspartate transcarbamlyase
OTCase
ornithine transcarbamylase
CP
carbamyl phosphate
ORN
L-ornithine
PALAO
Nδ-(phosphonacetyl)-Nα-acetyl-L-ornithine
SORN
N-succinyl-L-ornithine
WT
wild-type
xc
Xanthomonas campestris
Acknowledgments
We thank Dr. David Davies for facilitating our use of the diffraction equipment in the Molecular Structure Section
of the National Institute of Health and Dr. Fred Dyda for help in data collection and processing, and Dr. Yui-Fai
Lam in the University of Maryland for help in setting up NMR measurements.
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Figure 1.
Stereo view of the structure and hydrogen bonding network surrounding residue 302. A,
PALAO bound wild-type AOTCase, B, PALAO bound K302A AOTCase C, PALAO bound
K302E AOTCase, D, PALAO bound K302R AOTCase. Contours of the electron density
maps (2Fo-Fc) around PALAO, residue 302 and water molecules are shown as a brown cage
at 1.0σ. The final refined positions of the ligands and surrounding protein residues are
represented as colored sticks. The predicted hydrogen bonding interactions are in pink
dashed lines. The water molecules are represented as pink balls. The carbon of PALAO,
residue 302 and other protein residues are shown in pink, light blue and green sticks,
respectively.
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Figure 2.
13C NMR spectra of wild-type (upper panel) and K302A mutant (lower panel) AOTCase (1
mM). Experiments were performed in 100 mM Tris HCl, 50 mM NaCl, 7% D2O, pH 8.0,
supplemented with 20 mM NaH13CO3. The position of the resonance attributed to
carboxylated lysine in the enzyme is around 164 ppm.
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Figure 3.
Chemical structure of carbamylated vs. carboxylated lysine.
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Table 1
Data collection and refinement statistics
Dataset
PALAO
K302A
K302E
K302R
Space group
I213
I213
I213
I213
Resolution (Å)
2.2
1.9
1.85
2.2
Unit-cell parameters (Å)
a = b = c =128.88
a = b = c =128.92
a = b = c =129.29
a = b = c =127.39
Measurements
219,475
305,757
390,128
246,817
Unique reflections
18,269 (1,832) a
28,236 (1,365)
30,622 (1,456)
17,635 (879)
Redundancy
12.0 (11.8)
10.8(5.4)
12.8 (5.4)
14.0 (13.1)
Completeness (%)
99.8 (100.0)
100.0 (100.0)
99.7 (95.1)
100.0 (100.0)
<I/σ (I)>
15.0 (4.9)
16.4 (2.3)
19.8 (2.8)
8.7 (3.7)
Rmerg b
7.4 (48.4)
6.5(64.9)
5.2 (55.3)
9.8 (79.1)
Wilson B (Å2)
30.4
27.6
28.6
21.9
Refinement
Resolution range (Å)
50.0-2.2
50-1.9
50-1.85
50-2.2
No. of protein atoms
2620
2613
2617
2619
No. of water atoms
90
219
193
146
No. of hetero atoms
24
24
24
24
Rmsd of bond lengths (Å)
0.006
0.005
0.005
0.005
Rmsd of bond angle (°)
1.1
1.2
1.2
1.2
Rwork (%)c
20.0
19.8
20.0
18.9
Rfree (%)d
24.3
23.2
23.2
22.2
Average B factor (Å2)
41.7
32.2
32.3
35.3
aFigures in brackets apply to the highest-resolution shell.
bRmerg = ΣhΣi|I(h,i)-<I(h)>|/∑hΣiI(h,i), where I(h,i) is the intensity of the ith observation of reflection h, and < I(h)> is the average intensity of
redundant measurements of reflection h.
cRwork= Σh‖Fobs| – |Fcalc‖/Σh|Fobs|.
dRfree = Σh‖Fobs| – |Fcalc‖/Σh|Fobs| for 5% of the reserved reflections.
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Table 2
Interactions between carboxylated lysine and other residues at the active site of AOTCase
Kcx302
Other residues
Bound ligands
PALAO
CPa
AORNb
CP +ANORc
SO4+ACITd
OQ1
K252 NZ
2.6
2.6
2.7
2.6
2.6
OQ1
W1e
2.6
2.6
2.6
2.7
OQ2
S253 N
3.0
3.1
2.8
2.9
2.9
OQ2
H293 NE2
3.0
3.2
3.0
2.9
2.9
NZ
W2f
3.1
2.9
3.0
3.0
aThe values were calculated based on PDB ID 3KZM.
bThe values were calculated based on PDB ID 3KZN.
cThe values were calculated based on PDB ID 3KZO.
dThe values were calculated based on PDB ID 3KZK.
eThis water molecule hydrogen bonds to N1 atom of PALAO, AORN or ANOR, and backbone O atom of Pro296 as well.
fThis water molecule hydrogen bonds to OE1 atom of Glu92 from adjacent subunit as well.
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Table 3
Specific activity of wild-type and mutant AOTCase in the presence of acids (0.5M).
Compounds added
Specific activity(µmol/min/mg)
Wild-type
K302A
K302E
K302R
None
43.4 ± 0.4a
23.0 ± 0.5
7.1 ± 0.1
0.059±0.01
Formate
44.1 ± 1.2
26.4 ± 0.6
6.7 ± 0.2
0.093±0.01
Acetate
48.5 ± 1.1
21.2 ± 0.8
6.6 ± 0.5
0.104±0.03
aThe Mean ± S.D. are shown (n = 3).
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Table 4
Protein structures with lysine carboxylation modification
PDB ID
Enzyme name
Residue
Organism source
Funciton
2OGJ
Dihydroorotase
175
A.tumefaciens
Bridging two Zn(II)
2Z26
Dihydroorotase
102
E.coli
Bridging two Zn(II)
3JZE
Dihydroorotase
103
S.enterica
Bridging two Zn(II)
2GWN
Dihydroorotase
149
P. gingivalis
Bridging two Zn(II)
3F4C
Organophosphorus hydrolase
243
G. stearothermophilus
Bridging two Co(II)
3ICJ
Metal-dependent hydrolase
294
P. furiosus
Bridging two Zn(II)
3GTX
Organophosphorus hydrolase
243
D. radiodurans
Bridging two Co(II)
2QPX
Metal-dependent hydrolase
166
L. casei
Bridging two Zn(II)
2FTW
Dihydropyrimidinase
158
D. discoideum
Bridging two Zn(II)
2FVK
Dihydropyrimidinase
167
S. kluyveri
Bridging two Zn(II)
3DC8
Dihydropyrimidinase
147
S. meliloti
Bridging two Zn(II)
3GNH
L-Lys/Arg carboxypeptidase
211
C. crescentus cb15
Bridging two Zn(II)
3DUG
Arginine carboxypeptidase
182
Unidentified
Bridging two Zn(II)
2VC7
Phosphotriesterase
137
S. solfataricus
Bridging two Co(II)
2R1N
Metallophosphotriesterases
169
A. tumefaciens
Bridging two Co(II)
2OB3
Phosphotriesterase
169
B. diminuta
Bridging two Zn(II)
3E74
Allantoinase
146
E. coli
Bridging two Fe(III)
1EJX
Urease
217
K. aerogenes
Bridging two Ni(II)
1E9Z
Urease
219
H. pylori
Bridging two Ni(II)
4UBP
Urease
220
B. pasteurii
Bridging two Ni(II)
1ONW
Isoaspartyl dipeptidase
162
E. coli
Bridging two Zn(II)
1K1D
D-hydanroinase
150
G. stearothermophilus
Bridging two Zn(II)
1GKR
L-hydanroinase
147
A. aurescens
Bridging two Zn(II)
1GKP
D-hydanroinase
150
Thermus sp.
Bridging two Zn(II)
1NFG
D-hydantoinase
148
R. pickettii
Bridging two Zn(II)
2ICS
Adenine deaminase
154
E. faecalis
Bridging two Zn(II)
1RQB
Transcarboxylase
184
P. freudenreichii
Binding one Co(II)
2QF7
Pyruvate carboxylase
718
R. etli
Binding one Zn(II)
3BG3
Pyruvate carboxylase
741
H. sapiens
Binding one Mn(II)
2OEM
Rubisco-like protein
173
G. kaustophilus
Binding one Mg(II)
1WDD
Rubisco
201
O. sativa
Binding one Mg(II)
1GK8
Rubisco
201
C. reinhardtii
Binding one Mg(II)
1BWV
Rubisco
201
G. partita
Binding one Mg(II)
2WTZ
ATP-dependent MurE ligase
262
M. tuberculosis
Binding one Mg(II)
2JFG
MurD ligase
198
E. coli
Catalytic role?
1E8C
MurE ligase
224
E. coli
Catalytic role?
1JBW
Folypolyglutamate synthetase
185
L. casei
Catalytic role?
1W78
FolC bifunctional protein
188
E. coli
Binding one Mg(II)
3HBR
OXA-48 β-lactamase
73
K. pneumoniae
Catalytic role
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PDB ID
Enzyme name
Residue
Organism source
Funciton
3ISG
Class D β-lactamase
70
E. coli
Catalytic role
2P9V
AmpC beta-lactamase
315
E. coli
Catalytic role
1K55
OXA-10 β-lactamase
70
P. aeruginosa
Catalytic role
1K38
β-lactamase OXA-2
70
S. typhimurium
Catalytic role
1XQL
Alanine racemase
129
G. stearothermophilus
Binding substrate?
1VFS
Alanine racemase
129
S. lavendulae
Binding substrate?
1RCQ
Alanine racemase
122
P. aeruginosa
Binding substrate?
2J6V
UV damage endonuclease
229
T. thermophilus
Catalytic role
1H01
Cell division protein kinase 2
33
H. sapiens
Catalytic role?
2UYN
Protein TdcF
A58
E. coli
Structural role?
1HL9
Fucosidase
338
T. maritime
Structural role?
1PU6
3-methyladenine DNA glycosylase
205
H. pylori
Structural role?
Biochemistry. Author manuscript; available in PMC 2011 August 17.
|
3M5D
|
Crystal structure of N-acetyl-L-ornithine transcarbamylase K302R mutant complexed with PALAO
|
Reversible Post-Translational Carboxylation Modulates The
Enzymatic Activity Of N-Acetyl-L-Ornithine Transcarbamylase†
Yongdong Li1,2, Xiaolin Yu1, Jeremy Ho1, David Fushman3, Norma M. Allewell3, Mendel
Tuchman1, and Dashuang Shi1,‡
1Research Center for Genetic Medicine and Department of Integrative Systems Biology,
Children’s National Medical Center, The George Washington University, Washington, DC 20010,
USA.
2Key Laboratory of Organo-Pharmaceutical Chemistry, Jiangxi Province, Gannan Normal
University, Ganzhou 341000, China.
3Department of Chemistry and Biochemistry, College of Chemical and Life Sciences, University of
Maryland, College Park, MD 20742, USA.
Abstract
N-acetyl-L-ornithine transcarbamylase (AOTCase), rather than ornithine transcarbamylase
(OTCase), is the essential carbamylase enzyme in the arginine biosynthesis of several plant and
human pathogens. The specificity of this unique enzyme provides a potential target for controlling
the spread of these pathogens. Recently, several crystal structures of AOTCase from Xanthomonas
campestris (xc) have been determined. In these structures, an unexplained electron density at the
tip of Lys302 side-chain was observed. Using 13C NMR spectroscopy, we show herein that
Lys302 is post-translationally carboxylated. The structure of wild-type AOTCase complexed with
the bisubstrate analogue, Nδ-(phosphonoacetyl)-Nα-acetyl-L-ornithine (PALAO), indicates that the
carboxyl group on Lys302 forms a strong hydrogen bonding network with surrounding active site
residues, Lys252, Ser253, His293, and Glu92 from the adjacent subunit either directly or via a
water molecule. Furthermore, the carboxyl group is involved in binding N-acetyl-L-ornithine via a
water molecule. Activity assays with the wild-type enzyme and several mutants demonstrate that
the post translational modification of lysine 302 has an important role in catalysis.
Post-translational modification of the ε-amino group of lysine residues in proteins is a
common mechanism used by organisms to regulate protein functions including DNA-protein
interactions, subcellular localization, transcriptional activity, and protein stability and
activity (1). Lysine residues can be modified by the addition of functional groups to become
acetylated, methylated, carbamylated or carboxylated. The role of histone lysine acetylation
and methylation in affecting chromatin structure and gene expression has been well
established for more than a decade (2). However, the biological roles for lysine
carbamylation and carboxylation have rarely been investigated.
†This work was supported by Public Health Service grants DK-47870 (MT) and DK-067935 (DS) from the National Institute of
Diabetes, Digestive and Kidney Diseases. JH was supported by a Scholarship from the Doug and Lynn Parsons Family Foundation.
The Cornell High Energy Synchrotron Source (CHESS) is supported by the National Science Foundation under award DMR 0225180
and the Macromolecular Diffraction Facility at CHESS (MacCHESS) is supported by award RR-01646 from the National Institutes of
Health, through its National Center for Research Resources.
‡Corresponding author. dshi@cnmcresearch.org. Phone: 202-476-5817. Fax: 202-476-6014.
SUPPORTING INFORMATION AVAILABLE
Figure S1. Structure and hydrogen bonding network around residue 302 for previously determined AOTCase structures. This material
is available free of charge via the Internet at http://pubs.acs.org.
NIH Public Access
Author Manuscript
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Published in final edited form as:
Biochemistry. 2010 August 17; 49(32): 6887–6895. doi:10.1021/bi1007386.
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In vivo, lysine acetylation and methylation are usually carried out by acetyltransferase and
methyltransferase enzymes, respectively (3). In addition, some proteins such as hemoglobin
and human serum albumin can be acetylated non-enzymatically by chemicals such as
aspirin, methyl acetyl phosphate, and other acetylating agents such as acetyl-CoA (4–8).
Lysine can also be methylated by small chemicals in vitro, and this has routinely been used
as a rescue method for protein crystallization (9). Lysine carbamylation and lysine
carboxylation have only been achieved by using chemicals and no enzyme has yet been
found to catalyze these modifications. Lysine carbamylation was one of the earliest post-
translational modification of proteins to be elucidated when it was identified as a product of
reversible denaturation-renaturation studies of proteins with urea (10,11). This
carbamylation, which produces homocitrulline, has also been detected in uremic patients
(12) and in patients with elevated plasma and/or urinary lysine levels (13). In contrast, lysine
carboxylation is not as commonly reported, but has been identified in a number of proteins
via crystal structure determinations. In most of these proteins, the carboxyl groups of
modified lysines are involved in bridging two metal ions that play a structural role in the
active site. In several other proteins, however, a direct role for a carboxylated lysine in the
catalytic mechanism has been reported (14–16).
N-acetyl-L-ornithine transcarbamylase (AOTCase, EC 2.1.3.9) was recently discovered to
be part of a novel arginine biosynthesis pathway in plant pathogens of the
Xanthomonadaceae family such as Xylella and Xanthomonas (17–19). These pathogens
attack a variety of economically important crops including citrus fruits, cotton, tomatoes,
and rice (20,21). Genome sequence analyses showed that an AOTCase-like gene is also
present in some human pathogens such as Stenotrophomonas maltophilia and members of
the genus Bacteroides (22). In the case of Bacteroides fragilis, this gene was later confirmed
to encode another novel transcarbamylase, N-succinyl-L-ornithine transcarbamylase
(SOTCase, EC 2.1.3.11) (23). Crystal structures of both AOTCase and SOTCase bound with
substrate or substrate analogues have recently been determined (17,18,23). An extended
density at the side-chain tip of Lys302 in AOTCase was observed suggesting a post-
translational modification. Since Lys302 is located within the active site of AOTCase and is
not present in SOTCase, it was proposed as one of three key signature residues to
distinguish the two carbamylases (22). Here, we demonstrate that Lys302 is post-
translationally modified by carboxylation and that this change affects the catalytic function
of the enzyme.
MATERIALS AND METHODS
Materials
All chemicals were purchased from Sigma Chemical Company unless otherwise specified.
ANOR was purchased from Indofine Chemical Co., Inc. N-acetyl-L-citrulline was custom
synthesized and purified by Chiral Quest Company. PALAO (>95% purity) was synthesized
by IMI TAMI Institute of Research and Development Ltd. (19). xcAOTCase was prepared
and purified as previously described (18). Mutants K302A (primer: 5’-
CTGCGTCGCAACGTCGCGGCTACTGATGCGGTG-3’), K302E (primer:5’-
CTGCGTCGCAACGTCGAGGCTACTGATGCGGTG-3’) and K302R (primer: 5’-
CTGCGTCGCAACGTCAGGGCTACTGATGCGGTG-3’) were generated by site-directed
mutagenesis using the “Quik Change” mutagenesis kit (Stratagene) according to the
manufacturer’s protocol. The correct mutants were confirmed by DNA sequencing.
Recombinant mutant proteins were expressed and purified in the same manner as the wild-
type enzyme.
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Activity assay
The modified colorimetric assay method, which detects the formation of the ureido group
during the transcarbamylation reaction (24), was used to measure enzyme activity. CP and
AORN concentration were kept constant as 4.0 mM. After an incubation of 5 minutes, the
reaction was stopped by the addition of 1 ml of color reagent, as described previously (19).
A set of tubes containing known amounts of N-acetyl-L-citrulline was included with each
rack of enzyme assays to produce a standard curve for calculation of the enzyme specific
activity.
Mass spectrometric analysis
In order to identify the post-translational modification, mass spectrometric analysis was
carried out on a 4700 ABI TOF/TOF mass spectrometer (Applier Biosystems) based on the
method described previously (25). In brief, 10 µg of native protein were digested overnight
at 310 K using trypsin in 50 mM ammonium bicarbonate pH 7.4. After desalting using a
C18 ZipTip micropipette tip, the resulting peptides were eluted in 10 µl of acetonitrile/0.1%
TFA [70:30(v:v)]. The sample was mixed with matrix solution and spotted on a MALDI
plate to be submitted to the mass spectrometric analysis.
Chemical rescue experiments
The assay in the presence of various selected chemical was conducted as described above.
The stock solutions of small chemicals were titrated to the pH of the assay with KOH or
HCl.
13C NMR experiments
The wild-type and K302A mutant protein of AOTCase (~10 mg) was precipitated by
degassed buffer (pH 4.5) containing 25 mM sodium acetate. After centrifugation, the
precipitate was re-dissolved by adding a buffer containing 20 mM NaH13CO3, 100 mM Tris
HCl (pH 8.0) and 50 mM NaCl. Before NMR experiments, 40 µl D2O was added to 500 µl
protein sample. The 13C NMR spectra were collected on a Bruker Avance 600 spectrometer
(operating at 14.1 T) equipped with a direct 13C-detection probe at 298 K. The experimental
settings and processing parameters for the wild-type protein and K302A variant were
identical. 512 transients were collected with 4K time domain points and a spectral width of
3019 Hz centered at 160 ppm. The spectra were processed using exponential multiplication
with the line broadening factor set to 3Hz. The similarity of protein concentration in both
samples was verified by 1H NMR (not shown).
Crystallization, data collection and processing
PALAO-bound wild-type and mutant AOTCase crystals were grown using the hanging-drop
vapor diffusion method, with conditions similar to those used to produce native and ligand-
complexed AOTCase crystals (18,23). 2.0 µl of ~10 mg/ml solution of AOTCase were
mixed with 1.6 µl of reservoir solution and 0.4 µl PALAO solution (~0.01 M). The reservoir
solution contained 20% (w/v) PEG 3350, 0.2 M lithium sulfate and 0.1 M bis-Tris, pH 6.0.
Diffraction data for the PALAO-bound crystal were collected at 100 K at the F1 beam line
of the Cornell High Energy Synchrotron Source. Data sets for the PALAO-bound mutant
AOTCase crystals were obtained using a Rigaku anode x-ray generator in the Molecular
Structure Section of the National Institute of Health. All data were processed using
HKL2000 package (26) and reduced using the program TRUNCATE in the CCP4 suite (27).
Data collection parameters are listed in Table 1.
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Structure solution and refinement
Molecular replacement was used for phase determination of the PALAO-bound wild-type
and mutant AOTCase structures. The coordinates of AOTCase (PDB code: 3KZO) after
removal of ligands or water molecules were used for phase determination. Upon rigid-body
refinement, electron density corresponding to the ligands could be clearly visualized. The
ligands were built into the map using the graphics program O (28). Refinements were
carried out using molecular annealing, energy minimization and restrained B factor
refinement with the program CNS1.1 (29). During refinements, 5% of the reflections at
various resolutions were randomly selected to set aside to calculate Rfree to monitor the
progress of refinement (30). After every cycle of refinement, the model was manually
adjusted using the program O (28). Water molecules were automatically assigned using the
program WATERPICK of CNS. Model quality was checked using the program
PROCHECK (31) to ensure good stereochemistry for all three models. The final refinement
statistics are listed in Table 1.
Figures 1 was drawn using the programs Pymol (http://www.pymol.org). Figure 3 was
drawn using ChemDraw 8.0. The coordinates have been deposited with the RCSB PDB as
entries 3M4J, 3M4N, 3M5C and 3M5D.
RESULTS
Lys302 in AOTCase is carboxylated
To investigate the nature of the modification of Lys302 and how it affects catalytic activity,
we revisited all AOTCase structures. In the PALAO-bound AOTCase structure, the electron
density map clearly indicates that Lys302 is post-translationally modified (Figure 1A). The
type of modification can include methylation, acetylation, carbamylation, and carboxylation.
The shape of the electron density can been used to distinguish methyl groups from larger
functional groups, but it is difficult to distinguish between acetyl, carbamyl, and carboxyl
groups, all of which have three non-hydrogen atoms in a plane. Given the hydrogen bonding
network with surrounding residues (Lys252, Ser253 and His293, Table 2), a carboxylated
modification is the most likely choice for the modification of Lys302 in AOTCase. To
exclude that the modification’s identity represents chemically stable moieties (methyl,
acetyl, carbamyl), we analyzed trypsin digested fragments of purified AOTCase by TOF-
TOF mass spectroscopy. As expected, only a peptide fragment with an unmodified Lys302
was observed, consistent with the lability of the carboxylic group in acidic solutions. At low
pH, the carboxyl group is spontaneously released as carbon dioxide (14, 32), in contrast to
other modified groups that are stably bound and can be observed by mass spectrometry
analysis after proteolysis (33).
The putative carboxyl group on the modified Lys302 forms direct hydrogen bonds with
main-chain or side-chain nitrogen atoms of Lys252, Ser253 and His293 (Figure 1A and
Table 2). Among these, Lys252 is involved in direct hydrogen bonding to the carboxyl
group of the AORN moiety of PALAO, and His293 forms a strong hydrogen bond with the
main-chain nitrogen atom of Leu295 in the conserved His293-Cys294-Leu295-Pro206
(HCLP) motif. The hydrogen bonding network between the carboxyl group of modified
Lys302, His293 and the main-chain nitrogen atom of Leu295 is reminiscent of the similar
hydrogen bonding network, Glu310-His302-Leu304 and Glu299-Leu272-Leu274, found in
human and E. coli OTCase, respectively (34, 35). These three residues are conserved in all
OTCase sequences, and the interactions between them are important for maintaining the
HCLP motif in a specific conformation to orientate their main-chain oxygen atoms towards
the active site. In all known transcarbamylase structures, a leucine residue corresponding to
Leu295 is in an energetically unfavorable conformation and the peptide bond between this
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leucine and Pro296 is in the cis conformation. In addition to the direct hydrogen bonding
interaction above, the carboxyl group on the modified Lys302 interacts with the α-amino
nitrogen atom of the AORN moiety of PALAO and Glu92 from the adjacent subunit via
water molecules.
When we revisited all previously determined AOTCase structures (see supplementary
Figure S1) we found: (1) Lys302 was carboxylated in the absence of substrate binding, but
substrate binding immobilizes the side-chain of Lys302 further by hydrogen bonding
interaction via water molecules. (2) Water-mediated hydrogen bonding promotes
interactions of carboxylated Lys302 with Glu92 from the adjacent subunit and the α-amino
nitrogen atom of AORN. (3) Similarly to AOTCase, in the structure of SOTCase E92Z (Z =
Ala, Ser, Pro, Val), mutant with N-succinyl-L-norvaline bound (22) the carboxylated Lys302
hydrogen bonds to the α-amino nitrogen atom and the succinyl carboxyl group of N-
succinyl-L-norvaline via water molecules (Figure S1).
To obtain direct, independent evidence for the carboxylation of Lys302, 13C NMR
experiments were carried out with both wild-type protein and the K302A mutant. As
observed for other proteins with carboxylated lysine (36,37), the strong 13C NMR signal at
164 ppm characteristic of a carboxyl group was clearly detectable in AOTCase wild-type
protein labeled by 13C-bicarbonate, in contrast to the K302A mutant where the signal was
weak (Figure 2). Since there are 17 other lysine residues in the protein, the weak signal seen
for the K302A mutant might be caused by the adventitious carboxylation of another lysine
with reduced pKa, as has been observed for the K392A mutant of the sensor domain of the
BlaR protein (38).
Functional and structural studies of Lys302 mutants
To investigate the effect of lysine carboxylation on enzyme activity, Lys302 was mutated to
alanine, glutamate or arginine. Each of these variants was expressed in E. coli and gave
similar yields. Enzymatic assays demonstrated a significant decrease in enzymatic activity in
all three mutants, reflecting the functional importance of Lys302 (Table 3). The level of
enzymatic activity for the wild-type (WT) and three mutants was WT > K302A > K302E ≫
K302R. To determine the structural basis of these results, the WT and mutant enzymes
bound with PALAO were crystallized and their structures were determined at 1.8–2.2 Å
resolution. Only the K302R mutation had and appreciable effect on the structure of the
protein. Since K302 is located near the AORN binding site, the mutations would weaken
AORN binding to the active site.
In the structure of the K302A mutant, three additional water molecules (labeled as w3, w4
and w5 in Figure 1B) replace the carboxylated lysine. The two water molecules (labeled w1
and w2 in Figure 1A–1D) that mediate the hydrogen bonding interaction of carboxylated
Lys302 with PALAO and Glu92 from the adjacent subunit are also found in the K302A
mutant structure. Furthermore, these water molecules maintain a similar hydrogen-bonding
network to the wild-type enzyme. These results might explain why the K302A mutant
retains significant catalytic activity (Table 3). To investigate whether adding short-chain
carboxylic acids to the K302A mutant increases its activity as other enzymes (14, 15, 39,
40), the activity of the K302A mutant was measured in the presence of high formate and
acetate concentration (0.5 M). Surprisingly, the activity of the K302A mutant was not
significantly improved. The crystal structure of the K302A mutant soaking with the
crystallization buffer in the presence of 0.5 M acetate was also determined (not shown) and
it was observed that the same five water molecules were present in the cavity that replaced
the side chain of the carboxylated lysine. This, the acetate’s inability to replace the water
molecules in the crystal structure, is consistent with the unchanged activity assay results.
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The side-chain of Glu302 in the K302E mutant structure is well defined and anchored by
hydrogen bonding interaction with the main-chain nitrogen atom of Arg298 and weakly
hydrogen bonded to the main-chain nitrogen atom of Ser253 (Figure 1C). Two of three
additional water molecules (w4 and w5) observed in the K302A mutant structure occupied
the same position as the carboxyl oxygen atoms of Glu302 and form a similar hydrogen-
bonding network. Relative to the PALAO-bound wild-type structure, there is only one more
water molecule (w3) at the position of the carboxyl group of the carboxylated Lys302. This
water molecule mediates a hydrogen bonding interaction between Glu302 and Lys252. Two
common water molecules (w1 and w2) that interact with PALAO and Glu92 from the
adjacent subunit, respectively, were also identified in the K302E structure.
Our observation that the K302E mutant had lower enzymatic activity than that of the K302A
mutant (Table 3) was surprising since the carboxyl group of the glutamate could conceivably
function similarly to a carboxylated lysine. The explanation may be that, in the K302A
mutant, the hydrogen bonding network is well maintained by water molecules in the cavity
that replaces the carboxylated lysine. In particular, w3 is optimally located for strong
hydrogen bonding to w1 (2.7 Å), which in turn binds AORN. The distances between w1 and
the carboxyl oxygen of carboxylated Lys302 in all wild-type crystal structures are within
2.4–2.7 Å, but the distance between w1 and w2 in the K302E structure is significantly
greater (3.2 Å). The weaker hydrogen bonding interaction may be a reason for lower
enzymatic activity of the K302E mutant.
In contrast to the K302A and K302E structures, the K302R structure shows a much larger
reduction in enzyme activity relative to the wild-type enzyme. The electron density for the
side-chain of Arg302 is weak and the temperature factor of its side-chain is 54.4 Å2,
significantly higher than those of carboxylated Lys302 (44.7 Å2) and Glu302 (33.4 Å2),
implying greater flexibility. Furthermore, the side-chain of Arg302 is oriented differently
from the carboxyl group of carboxylated Lys302 and pushes the nearby residues His180,
Pro181 and Lys182 outwards about 1.0 Å (Figure 1D). However, the water molecules
involved in hydrogen bonding to the α-amino nitrogen atom of PALAO (w1) and the side-
chains of Lys252 (w3) and Glu92 (w2) from the adjacent subunit are conserved. Consistent
with the K302E structure, the distance between w1 and w2 is even greater (3.4 Å) than in
the WT structure and the hydrogen bonding interaction between w2 and w3 is no longer
observed. Thus, the almost undetectable enzymatic activity of the K302R mutant probably
results from the changes at its active site, including the weakened hydrogen bonding
network involved in substrate binding.
DISCUSSION
Several lines of evidence clearly indicate that Lys302 in AOTCase is carboxylated. First, the
extra electron density indicates that the side-chain of Lys302 is modified. Second, the
hydrogen bonding environment of Lys302 for hydrogen bonding interactions is compatible
with a carboxyl group, but not for a positively charged lysine side-chain. Third, the
modification is labile at low pH, since mass spectroscopy of samples prepared at low pH
indicated that Lys302 was no longer modified. Fourth, the clear presence of the
indicative 13C NMR signal at 164 ppm for wild-type protein and its absence in the K302A
mutant confirms carboxylation of Lys302.
It is well known that lysine carboxylation is non-enzymatic and reversible, while other post-
translational modifications such as methylation, acetylation, and carbamylation are
irreversible and detectable by mass spectroscopy. Furthermore, lysine methylation and
acetylation usually require an enzyme-catalyzed reaction in vivo (41). Therefore, it is
unlikely that such lysine modifications will be observed in recombinant proteins
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overexpressed in a foreign host (e.g. E. coli). Lysine methylation can be achieved by using
special chemicals in vitro, but these chemicals are not present in vivo. Lysine carbamylation
and carboxylation use completely different mechanisms to form functionally different
groups (Figure 3). Carbamylation can be achieved by cyanate produced from
myeloperoxidase-catalyzed oxidation of thiocyanate, an anion abundant in plasma and
increased in smokers, or from urea in the plasma. Lysine carboxylation, on the other hand,
occurs readily in aqueous solution in the presence of carbon dioxide at a basic pH (32,42).
Even though carbamylation and carboxylation use very different mechanisms, the two are
confused in the literatures. Lysine carbamylation (or carbamoylation) is referred to in
several publications (15,32,42–44), when the actual reaction is in fact carboxylation.
The activity of the K302A mutant is almost half of that of the wild-type enzyme raising the
question of why AOTCase retains a lysine in this position. Perhaps this lysine was
maintained through evolution to distinguish AOTCase from SOTCase which uses N-
succinyl-L-ornithine (SORN) rather than AORN (22), and OTCase which uses L-ornithine.
An alternative explanation may be found in the very low activity of the K302R mutant. The
side-chain of arginine has a positive charge while carboxylated lysine has a negative charge.
The side chain of unmodified lysine is usually located in a similar position as that of
arginine, as observed in the structure of UV damage endonuclease (14). It would be
expected that the activity of AOTCase with an uncarboxylated lysine would be as low as the
K302R mutant’s. It could further be surmised that, the respective organisms need to use
carboxylation as a switch to turn “on” or “off” the arginine biosynthetic pathway. It has been
well known that rubulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) in plant cells
uses the carboxylation on Lys201 as a switch to turn the enzyme “on” during the day and
“off” at night by removing the carboxyl group (45,46). Carbon dioxide and bicarbonate have
been found to play an important biological role in modulating several biological processes
including photosynthetic carbon fixation (47), pH homeostasis (48), carbon metabolism
(49), activation of virulence in pathogenic organisms (50), sperm maturation (51),
stimulation of mammalian G-protein-responsive adenylyl cyclase (52), and as an alarmone
in Drosophila (53,54). Whether or not carboxylation of a key lysine in their related proteins
is used as an underlying regulatory mechanism should be investigated further.
There are 197 structures with carboxylated lysine residue (modified residue indicated as
Kcx) in the Protein Data Bank (PDB). If structures with 90% identity are counted only once,
there are still 52 unique structures remaining in this pool (Table 4). These proteins include
hydantoinase (40,55), folylpolyglutamate synthase (43), UV damage endonuclease (14),
OXA10, OXA-1 class D β-lactamase (38,56,57), urease (42), phosphotriesterase (58),
dihydroorotase (59), dihydropyrimidinase (60), organophosphate hydrolase (61) and MurE
and MurD ligases (44,62). In most of these proteins, the carboxylated lysine bridges two
metal ions, similar to the role of glutamate or aspartate in proteins with two metal-binding
sites (26 structures among 52). However, the urease apoenzyme can be activated in vitro
only in the presence of carbon dioxide prior to nickel binding (63), suggesting that the
carboxylated lysine may have other structural roles beyond binding metals. In some proteins
such as β-lactamase, UV damage endonuclease, Rubisco, MurD and MurE ligase and BlaR
signal transducer protein, a carboxylated lysine plays an essential catalytic role. More
interestingly, in three structures (PDB ID: 1HL9, 1PU6 and 2UYN for fucosiadase, 3-
methyladenine DNA glycosylase and TdcF protein of unknown function, respectively), the
carboxylated lysines are located near the surface of proteins, presumably playing primarily a
structure stabilizing role (64–66). Since the carboxyl group is labile at acidic pH, but easily
formed in the presence of carbon dioxide at basic pH, the number of proteins with lysine
carboxylation must be underestimated. Furthermore, the carboxylated lysine must be fixed
in place by metal ions (either one or two) or hydrogen bonding with other protein residues
(at least one). Therefore, any detection method involving denaturing the proteins will result
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in release of the carboxyl group. With current technology, 13C NMR (38) and
crystallography are the only methods that can detect this modification. However, these
methods are not amenable to high-throughput investigations. The majority (49 out of 52
structures in the PDB) of known lysine carboxylation modifications were found to be
located at or near the active site, probably because these sites receive the most attention.
Revisiting the structures in PDB with more attention to surface lysines might reveal more
structures with carboxylated lysines.
In conclusion, we have shown that Lys302 in AOTCase is post-translationally modified by
carboxylation and that this modification may be functionally important for enzymatic
activity. Lysine carboxylation is likely to be a more common event than currently
appreciated and may play a critical role in enzymatic activity and protein stability.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Abbreviations
ACIT
N-acteyl-L-citrulline
ANOR
N-acetyl-L-norvaline
AORN
N-acetyl-L-Ornithine
AOTCase
N-acetyl-L-ornithine transcarbamylase
ATCase
aspartate transcarbamlyase
OTCase
ornithine transcarbamylase
CP
carbamyl phosphate
ORN
L-ornithine
PALAO
Nδ-(phosphonacetyl)-Nα-acetyl-L-ornithine
SORN
N-succinyl-L-ornithine
WT
wild-type
xc
Xanthomonas campestris
Acknowledgments
We thank Dr. David Davies for facilitating our use of the diffraction equipment in the Molecular Structure Section
of the National Institute of Health and Dr. Fred Dyda for help in data collection and processing, and Dr. Yui-Fai
Lam in the University of Maryland for help in setting up NMR measurements.
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Figure 1.
Stereo view of the structure and hydrogen bonding network surrounding residue 302. A,
PALAO bound wild-type AOTCase, B, PALAO bound K302A AOTCase C, PALAO bound
K302E AOTCase, D, PALAO bound K302R AOTCase. Contours of the electron density
maps (2Fo-Fc) around PALAO, residue 302 and water molecules are shown as a brown cage
at 1.0σ. The final refined positions of the ligands and surrounding protein residues are
represented as colored sticks. The predicted hydrogen bonding interactions are in pink
dashed lines. The water molecules are represented as pink balls. The carbon of PALAO,
residue 302 and other protein residues are shown in pink, light blue and green sticks,
respectively.
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Figure 2.
13C NMR spectra of wild-type (upper panel) and K302A mutant (lower panel) AOTCase (1
mM). Experiments were performed in 100 mM Tris HCl, 50 mM NaCl, 7% D2O, pH 8.0,
supplemented with 20 mM NaH13CO3. The position of the resonance attributed to
carboxylated lysine in the enzyme is around 164 ppm.
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Figure 3.
Chemical structure of carbamylated vs. carboxylated lysine.
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Table 1
Data collection and refinement statistics
Dataset
PALAO
K302A
K302E
K302R
Space group
I213
I213
I213
I213
Resolution (Å)
2.2
1.9
1.85
2.2
Unit-cell parameters (Å)
a = b = c =128.88
a = b = c =128.92
a = b = c =129.29
a = b = c =127.39
Measurements
219,475
305,757
390,128
246,817
Unique reflections
18,269 (1,832) a
28,236 (1,365)
30,622 (1,456)
17,635 (879)
Redundancy
12.0 (11.8)
10.8(5.4)
12.8 (5.4)
14.0 (13.1)
Completeness (%)
99.8 (100.0)
100.0 (100.0)
99.7 (95.1)
100.0 (100.0)
<I/σ (I)>
15.0 (4.9)
16.4 (2.3)
19.8 (2.8)
8.7 (3.7)
Rmerg b
7.4 (48.4)
6.5(64.9)
5.2 (55.3)
9.8 (79.1)
Wilson B (Å2)
30.4
27.6
28.6
21.9
Refinement
Resolution range (Å)
50.0-2.2
50-1.9
50-1.85
50-2.2
No. of protein atoms
2620
2613
2617
2619
No. of water atoms
90
219
193
146
No. of hetero atoms
24
24
24
24
Rmsd of bond lengths (Å)
0.006
0.005
0.005
0.005
Rmsd of bond angle (°)
1.1
1.2
1.2
1.2
Rwork (%)c
20.0
19.8
20.0
18.9
Rfree (%)d
24.3
23.2
23.2
22.2
Average B factor (Å2)
41.7
32.2
32.3
35.3
aFigures in brackets apply to the highest-resolution shell.
bRmerg = ΣhΣi|I(h,i)-<I(h)>|/∑hΣiI(h,i), where I(h,i) is the intensity of the ith observation of reflection h, and < I(h)> is the average intensity of
redundant measurements of reflection h.
cRwork= Σh‖Fobs| – |Fcalc‖/Σh|Fobs|.
dRfree = Σh‖Fobs| – |Fcalc‖/Σh|Fobs| for 5% of the reserved reflections.
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Table 2
Interactions between carboxylated lysine and other residues at the active site of AOTCase
Kcx302
Other residues
Bound ligands
PALAO
CPa
AORNb
CP +ANORc
SO4+ACITd
OQ1
K252 NZ
2.6
2.6
2.7
2.6
2.6
OQ1
W1e
2.6
2.6
2.6
2.7
OQ2
S253 N
3.0
3.1
2.8
2.9
2.9
OQ2
H293 NE2
3.0
3.2
3.0
2.9
2.9
NZ
W2f
3.1
2.9
3.0
3.0
aThe values were calculated based on PDB ID 3KZM.
bThe values were calculated based on PDB ID 3KZN.
cThe values were calculated based on PDB ID 3KZO.
dThe values were calculated based on PDB ID 3KZK.
eThis water molecule hydrogen bonds to N1 atom of PALAO, AORN or ANOR, and backbone O atom of Pro296 as well.
fThis water molecule hydrogen bonds to OE1 atom of Glu92 from adjacent subunit as well.
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Table 3
Specific activity of wild-type and mutant AOTCase in the presence of acids (0.5M).
Compounds added
Specific activity(µmol/min/mg)
Wild-type
K302A
K302E
K302R
None
43.4 ± 0.4a
23.0 ± 0.5
7.1 ± 0.1
0.059±0.01
Formate
44.1 ± 1.2
26.4 ± 0.6
6.7 ± 0.2
0.093±0.01
Acetate
48.5 ± 1.1
21.2 ± 0.8
6.6 ± 0.5
0.104±0.03
aThe Mean ± S.D. are shown (n = 3).
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Table 4
Protein structures with lysine carboxylation modification
PDB ID
Enzyme name
Residue
Organism source
Funciton
2OGJ
Dihydroorotase
175
A.tumefaciens
Bridging two Zn(II)
2Z26
Dihydroorotase
102
E.coli
Bridging two Zn(II)
3JZE
Dihydroorotase
103
S.enterica
Bridging two Zn(II)
2GWN
Dihydroorotase
149
P. gingivalis
Bridging two Zn(II)
3F4C
Organophosphorus hydrolase
243
G. stearothermophilus
Bridging two Co(II)
3ICJ
Metal-dependent hydrolase
294
P. furiosus
Bridging two Zn(II)
3GTX
Organophosphorus hydrolase
243
D. radiodurans
Bridging two Co(II)
2QPX
Metal-dependent hydrolase
166
L. casei
Bridging two Zn(II)
2FTW
Dihydropyrimidinase
158
D. discoideum
Bridging two Zn(II)
2FVK
Dihydropyrimidinase
167
S. kluyveri
Bridging two Zn(II)
3DC8
Dihydropyrimidinase
147
S. meliloti
Bridging two Zn(II)
3GNH
L-Lys/Arg carboxypeptidase
211
C. crescentus cb15
Bridging two Zn(II)
3DUG
Arginine carboxypeptidase
182
Unidentified
Bridging two Zn(II)
2VC7
Phosphotriesterase
137
S. solfataricus
Bridging two Co(II)
2R1N
Metallophosphotriesterases
169
A. tumefaciens
Bridging two Co(II)
2OB3
Phosphotriesterase
169
B. diminuta
Bridging two Zn(II)
3E74
Allantoinase
146
E. coli
Bridging two Fe(III)
1EJX
Urease
217
K. aerogenes
Bridging two Ni(II)
1E9Z
Urease
219
H. pylori
Bridging two Ni(II)
4UBP
Urease
220
B. pasteurii
Bridging two Ni(II)
1ONW
Isoaspartyl dipeptidase
162
E. coli
Bridging two Zn(II)
1K1D
D-hydanroinase
150
G. stearothermophilus
Bridging two Zn(II)
1GKR
L-hydanroinase
147
A. aurescens
Bridging two Zn(II)
1GKP
D-hydanroinase
150
Thermus sp.
Bridging two Zn(II)
1NFG
D-hydantoinase
148
R. pickettii
Bridging two Zn(II)
2ICS
Adenine deaminase
154
E. faecalis
Bridging two Zn(II)
1RQB
Transcarboxylase
184
P. freudenreichii
Binding one Co(II)
2QF7
Pyruvate carboxylase
718
R. etli
Binding one Zn(II)
3BG3
Pyruvate carboxylase
741
H. sapiens
Binding one Mn(II)
2OEM
Rubisco-like protein
173
G. kaustophilus
Binding one Mg(II)
1WDD
Rubisco
201
O. sativa
Binding one Mg(II)
1GK8
Rubisco
201
C. reinhardtii
Binding one Mg(II)
1BWV
Rubisco
201
G. partita
Binding one Mg(II)
2WTZ
ATP-dependent MurE ligase
262
M. tuberculosis
Binding one Mg(II)
2JFG
MurD ligase
198
E. coli
Catalytic role?
1E8C
MurE ligase
224
E. coli
Catalytic role?
1JBW
Folypolyglutamate synthetase
185
L. casei
Catalytic role?
1W78
FolC bifunctional protein
188
E. coli
Binding one Mg(II)
3HBR
OXA-48 β-lactamase
73
K. pneumoniae
Catalytic role
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PDB ID
Enzyme name
Residue
Organism source
Funciton
3ISG
Class D β-lactamase
70
E. coli
Catalytic role
2P9V
AmpC beta-lactamase
315
E. coli
Catalytic role
1K55
OXA-10 β-lactamase
70
P. aeruginosa
Catalytic role
1K38
β-lactamase OXA-2
70
S. typhimurium
Catalytic role
1XQL
Alanine racemase
129
G. stearothermophilus
Binding substrate?
1VFS
Alanine racemase
129
S. lavendulae
Binding substrate?
1RCQ
Alanine racemase
122
P. aeruginosa
Binding substrate?
2J6V
UV damage endonuclease
229
T. thermophilus
Catalytic role
1H01
Cell division protein kinase 2
33
H. sapiens
Catalytic role?
2UYN
Protein TdcF
A58
E. coli
Structural role?
1HL9
Fucosidase
338
T. maritime
Structural role?
1PU6
3-methyladenine DNA glycosylase
205
H. pylori
Structural role?
Biochemistry. Author manuscript; available in PMC 2011 August 17.
|
3M5G
|
Crystal structure of a H7 influenza virus hemagglutinin
|
Structures of Receptor Complexes of a North American
H7N2 Influenza Hemagglutinin with a Loop Deletion in
the Receptor Binding Site
Hua Yang, Li-Mei Chen, Paul J. Carney, Ruben O. Donis, James Stevens*
Influenza Division, Centers for Disease Control and Prevention, Atlanta, Georgia, United States of America
Abstract
Human infections with subtype H7 avian influenza viruses have been reported as early as 1979. In 1996, a genetically stable
24-nucleotide deletion emerged in North American H7 influenza virus hemagglutinins, resulting in an eight amino acid
deletion in the receptor-binding site. The continuous circulation of these viruses in live bird markets, as well as its
documented ability to infect humans, raises the question of how these viruses achieve structural stability and functionality.
Here we report a detailed molecular analysis of the receptor binding site of the North American lineage subtype H7N2 virus
A/New York/107/2003 (NY107), including complexes with an avian receptor analog (39-sialyl-N-acetyllactosamine, 39SLN)
and two human receptor analogs (69-sialyl-N-acetyllactosamine, 69SLN; sialyllacto-N-tetraose b, LSTb). Structural results
suggest a novel mechanism by which residues Arg220 and Arg229 (H3 numbering) are used to compensate for the deletion
of the 220-loop and form interactions with the receptor analogs. Glycan microarray results reveal that NY107 maintains an
avian-type (a2-3) receptor binding profile, with only moderate binding to human-type (a2-6) receptor. Thus despite its
dramatically altered receptor binding site, this HA maintains functionality and confirms a need for continued influenza virus
surveillance of avian and other animal reservoirs to define their zoonotic potential.
Citation: Yang H, Chen L-M, Carney PJ, Donis RO, Stevens J (2010) Structures of Receptor Complexes of a North American H7N2 Influenza Hemagglutinin with a
Loop Deletion in the Receptor Binding Site. PLoS Pathog 6(9): e1001081. doi:10.1371/journal.ppat.1001081
Editor: Fe´lix A. Rey, Institut Pasteur, France
Received March 16, 2010; Accepted July 28, 2010; Published September 2, 2010
This is an open-access article distributed under the terms of the Creative Commons Public Domain declaration which stipulates that, once placed in the public
domain, this work may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose.
Funding: This work was funded by the Centers for Disease Control and Prevention. Use of the Advanced Photon Source at Argonne National Laboratory was
supported by the U. S. Department of Energy, Office of Science, Office of Basic Energy Sciences, under Contract No. DE-AC02-06CH11357. Glycan microarrays as
well as glycans for direct binding experiments were produced for the Centers for Disease Control by the CFG funded by National Institute of General Medical
Sciences Grant GM62116. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: fwb4@cdc.gov
Introduction
Influenza is an acute respiratory virus that infects up to 20% of
the population in the United States, resulting in ,36,000 deaths
annually [1,2]. The two membrane glycoproteins on the surface of
influenza A virus, hemagglutinin (HA), which functions as the
receptor binding and membrane fusion glycoprotein in cell entry,
and neuraminidase (NA), which functions as the receptor destroying
enzyme in virus release, form the basis for defining subtypes [3]. To
date, 16 HA (H1–H16) and 9 NA (N1–N9) have been identified in
avian species [4], while in the last century, only three subtypes,
H1N1 in 1918 and 2009, H2N2 in 1957, and H3N2 in 1968 [5,6,7],
have successfully adapted to humans. Hemagglutinin binds to sialic
acid (SA) glycans present on host cell surfaces. The receptors on
epithelial cells of the human upper respiratory tract are mainly
a2-6-linked SA moieties [8]. Since avian influenza viruses pre-
dominately bind a2-3-linked SA, and human influenza viruses
preferentially bind to a2-6-linked SA, human infection by avian
influenza viruses is rare [9]. However, since 1997 a growing number
of human cases of avian influenza infection have been reported [10],
including H5N1, H7N2, H7N3, H7N7, and H9N2 strains [11].
Although the current situation with the pandemic H1N1 influenza
virus dominates public health efforts, the prospect of a novel
pandemic emerging from these isolated cases continues to be a
major public health threat around the world.
Early cases of human infection by H7 influenza viruses are reported
as far back as 1979 [12,13]. Since 2002, multiple outbreaks and
human infections of H7 subtype viruses; within both Eurasian and
North American lineages have been reported. In the Netherlands in
2003, a highly pathogenic avian influenza (HPAI) H7N7 outbreak
resulted in more than 80 cases of human infections, including one
fatality [14,15]. In New York in 2003, a single case of human
respiratory infection of H7N2 was reported [16] and in British
Columbia in 2004, an H7N3 virus caused two cases of conjunctivitis
[17,18]. More recently in 2007, the United Kingdom reported several
cases of low pathogenic avian influenza (LPAI) H7N2 virus infections
that caused influenza-like illness and conjunctivitis [19].
Since 1996, H7 viruses of the North American lineage have
been circulating in regional live bird markets [20], containing a
24-nucleotide deletion resulting in an eight amino acid deletion in
the receptor-binding site (RBS) of HA (Figure S1). The recent
human infections with H7 in North America have raised public
health concerns as to how these viruses adapt to such a dramatic
structural change while remaining one of the predominant
circulating viral strains. A recent study of H7 viruses isolated
from previous outbreaks revealed efficient replication in both
mouse and ferret animal models [21]. In particular, ferret studies
with A/New York/107/2003 (NY107), an H7N2 virus isolated
from a man in New York, not only showed efficient replication in
the upper respiratory tract of the ferret but also the capacity for
PLoS Pathogens | www.plospathogens.org
1
September 2010 | Volume 6 | Issue 9 | e1001081
intra-species transmission by direct contact [21,22]. Interestingly,
both an increased preference for a2-6 and decreased preference for
a2-3-linked sialosides of this virus compared to the other avian
influenza viruses was shown by previous glycan microarray analysis
but less so by a competitive solid-phase binding assay [22,23].
Here we report a detailed molecular analysis of the RBS of the
HA from North American lineage H7N2 virus, NY107, including
glycan microarray analyses and structural analyses of the HA in
complex with an avian receptor analog (39-Sialyl-N-acetyllactosa-
mine, 39SLN) and two human receptor analogs (69-Sialyl-N-
acetyllactosamine, 69SLN; Sialyllacto-N-tetraose b, LSTb). These
results provide important insight into the interaction of H7 HAs
with both avian and human hosts.
Results
Overall structure
By using x-ray crystallography, the structure of H7 HA from the
NY107 virus was determined to 2.6 A˚ resolution (Table 1). In
addition, we also report three H7 HA receptor complex structures,
with avian receptor analog (39SLN) to 2.7 A˚ resolution and with
human receptor analogs (69SLN and LSTb) to 3.0 A˚ and 2.6 A˚
resolution, respectively (Table 1). The overall structure of NY107
is similar to other reported HA structures with a globular head
containing the RBS and vestigial esterase domain, and a
membrane proximal domain with its distinctive, central helical
stalk and HA1/HA2 cleavage site (Figure 1A). Although five
asparagine-linked glycosylation sites are predicted in the NY107
HA monomer, interpretable electron density was observed at only
two sites, Asn38 in HA1 and Asn82 in HA2 (all residue numbers
are based on H3 numbering). At these sites, only one or two N-
acetyl glucosamines could be interpreted.
During viral replication, HA is synthesized as a single chain
precursor (HA0) and cleaved by specific host proteases into the
infectious HA1/HA2 form. In baculovirus expression systems,
highly pathogenic HAs, with a polybasic cleavage site, are
expressed as an HA1/HA2 form [24], whereas HAs with
monobasic cleavage sites (single Arg) from low pathogenic viruses
are expressed as the HA0 form [25]. NY107 is regarded as a low
pathogenic virus, and as expected, was produced in the HA0
form (Figure S2). However, subsequent digestion with thrombin
protease to remove the His-tag resulted in cleavage to a profile on
SDS-PAGE comparable to that of an HA1/HA2 form (Figure S2).
A comparison of the NY107 cleavage site with the consensus
cleavage pattern in the MEROPS database (http://merops.
sanger.ac.uk) suggests it to be a possible thrombin cleavage site.
Based on their molecular phylogenies, HAs are divided into two
groups and five clades: group 1 includes H8, H9, and H12; H1,
H2, H5, and H6; H11, H13 and H16; group 2 includes H3, H4,
and H14; H7, H10 and H15 [26]. Among all available HA
structures, we selected ten representative HAs from both avian and
human subtypes for structural analysis. As expected, NY107 HA is
structurally very similar to the Avian-H7 in all comparisons and
closely related to H3, the other group 2 members used in the
analyses (Tables S1 and S2).
The receptor binding site
The RBS is at the membrane distal end of each HA monomer
and its specificity for sialic acid and the nature of its linkage to a
vicinal galactose residue is a major determinant of host range-
restriction. The consensus RBS for all current HAs is composed of
three major structural elements: a 190-helix (residues 188–194), a
220-loop (residues 221–228), and a 130-loop (residues 134–138).
In addition, highly conserved residues (Tyr98, Trp153, His183,
and Tyr195) form the base of the pocket.
Although the NY107 RBS is similar to other subtypes (H1, H2,
H3, H5, and H9), a previously observed specific feature of H7 HAs,
is also observed in the NY107 150-loop region: two residues inserted
at position 158 result in this loop protruding more than 6A˚ towards
the binding site compared to other subtype HAs (Figure 1B and
Table S2) [27]. More interestingly, the eight amino acid deletion,
only found in the North American lineage H7s, from position 221 to
228 (Figure S1), resulted in a complete loss of the 220-loop
(Figure 1B). Sequence alignment shows that Arg220 and Arg229 are
conserved in all influenza A HA subtypes (Figure S1), but structural
alignment of NY107 HA shows Arg220 occupying the Gly228
position, and the much shorter loop turns at residue Pro217
(Figure 1C). The Ca distance between NY107 Arg220 and its
homolog in the Av-H7 structure (PDB: 1TI8) [27] is 5.8A˚ , and they
point in opposite directions (Figure 1C). The side chain direction of
Av-H7 Arg220 is almost parallel with the beta sheet after Arg229,
whereas the NY107 Arg220 points downward to the binding pocket.
The Ca position of Arg229 in both H7 structures remains the same,
except the side chain in the NY107 swings away by about 5.9A˚
(Figure 1C) and could help to stabilize this region by forming a
hydrogen bond to the mainchain carbonyl of Gln210 in the
neighboring monomer. In the absence of the 220-loop in NY107
HA, upon glycan binding the long side chain of Arg220 compensates
for its loss and is displaced 4A˚ upward to form hydrogen bonds with
receptor analogs inside the binding pocket (Figure 1D).
Effect of loop truncation on the receptor binding
specificity of NY107
Previously, mutations in the HA receptor binding domains of
H1N1 (Glu190Asp/Gly225Asp) and H2N2/H3N2 (Gln226Leu
and Gly228Ser) subtypes were responsible for adaptation of these
viruses to pandemic strains [24,28,29,30]. Due to missing residues
221–228 in the NY107 HA RBS, neither mechanism for
adaptation is possible. Thus, in order to look more closely at the
role of the missing loop and its effect on receptor specificity, we
first subjected the recombinant HA (recHA) to glycan microarray
analyses and compared it to a reverse genetics-derived NY107
virus, and a co-circulating Eurasian virus and recHA, A/
Author Summary
Influenza virus adaptation to different hosts usually results
in a switch in receptor specificity of the viral surface coat
protein, hemagglutinin. Indeed, the hemagglutinin sub-
types from the last two human influenza pandemics of the
20th Century (H2 in 1957 and H3 1968) both adapted
successfully to human-type receptor specificity through
only two amino acid mutations in the receptor binding
pocket (Glutamine226RLeucine and Glycine228RSerine).
The recent human infections reported with other avian
subtypes such as H5, H7 and H9 have raised public health
concerns and focused efforts on identifying potential
subtypes from which a future pandemic strain may
emerge. Since 1996, H7 viruses of the North American
lineage have been circulating in regional live bird markets,
containing an eight amino acid deletion in the receptor-
binding site of HA. Here we report a detailed structural
analysis of the receptor binding site of a hemagglutinin
from the North American lineage of H7N2 viruses, in
complex with avian and human receptor analogs, to
understand how these viruses have adapted to such a
dramatic structural change in the binding site while
remaining one of the predominant circulating viral strains.
Structure of a North American H7 Hemagglutinin
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Netherlands/219/2003 (NL219), that has the consensus avian
sequence in the 220-loop and it also infected a human [15].
Glycan microarray analysis of recombinant NY107 (Figure 2A
and Table 2) revealed a highly restricted binding profile with
strong binding to only a2-3 sulfated (#4–8), a2-3 branched (#9–
11) and mixed a2-3/a2-6 branched sialosides (#60–64) as well as
to the long linear sialyl di- and tri-lactosamines (#22, 24). Weak
binding was also observed (above background) to other a2-3
glycans on the array. The recombinant NY107 also revealed a
strict glycan binding preference to only one a2-6 glycan, the
internal structure, Galb1-3(Neu5Aca2-6)GlcNAcb1-3Galb1-4Glc
(#58; LSTb) (Figure 2A), a glycan highlighted in a previous study
[22]. The virus with higher valency and avidity revealed stronger
binding to all a2-3 groups, in addition to the branched di-sialyl a2-
6 biantennary structures (#46–48) as well the LSTb (#58)
(Figure 2B and Table 2). In contrast, the NL219 recHA (Figure 2C
and Table 2) bound well to only the avian a2-3 containing sialyl-
glycans (sulfated, branched, linear and fucosylated). Its corre-
sponding virus also reflected this specificity although it also
revealed strong binding to a2-3 N-glycolylneuraminic acid
(Neu5Gc) containing glycans (#66–70) (Figure 2D and Table 2).
To further assess the effect of the missing 220 loop on HA
structural stability and receptor specificity it was essential to
evaluate these functions on the ancestral HA containing the full
length 220-loop. To this end, we engineered an HA with an avian
H7 consensus (PQVNGQSG) 220-loop re-introduced (NY107-
220ins) into the NY107 HA and recovered this virus by reverse
genetics. Compared to the NY107 virus (Figure 2A) glycan
microarray analyses of the resulting NY107-220ins virus (Figure 3A
and Table 2) revealed a decrease in binding to branched (#9–11)
and linear (#12–27) a2-3 sialosides and a loss of binding to the
branched di-sialyl a2-6 biantennary structures (#46–48), LSTb
(#58) as well as the mixed a2-3/a2-6 branched sialosides (#60–
64). In addition, sequence analysis of the NY107-220ins HA
revealed the presence of quasispecies in the second position of the
inserted loop, P(Q/K)VNGQSG, suggesting that re-introduction
of the loop alone is not tolerated and does not create an avian-type
binding profile. Thus other amino acid substitutions in the HA
might have co-evolved with the deletion of the 220 loop to help
stabilize the RBS/HA to maintain functionality.
When viruses containing this 220-loop deletion emerged in North
America in the mid 90’s, four additional amino acid substitutions,
Gly114Arg, Asp119Gly, Gly186Glu and Gly205Arg, in the HA1 as
well as an Asp19Asn in the HA2 chain were also introduced to most
of the circulating isolates. Of these, Gly186Glu and Gly205Arg in
the HA1 are close to the RBS, at the monomer interface, and could
potentially modulate its structure and/or function. NY107 viruses
with a restored consensus 220-loop and a single Glu186Gly (NY107-
ins-186) or Arg205Gly (NY107-ins-205) substitution as well as the
Glu186Gly/Arg205Gly double substitution (NY107-ins-186/205)
were derived by reverse genetics and evaluated. Glycan microarray
analysis for the three resulting viruses revealed similar glycan
binding profiles with increased binding to a2-3 sialosides, including
mixed a2-3/a2-6 branched sialosides (#60–64), a2-3 Neu5Gc
(#66–70), but limited binding to the a2,6 sialosides (Figures 3B, 3C,
3D), resulting in a binding profile virtually identical to that of the
NL219 virus and other avian influenza viruses (Figure 2D) [30].
Sequence analysis of the three reverse genetics derived viruses
revealed no mutations/quasispecies in the HAs of either the NY107-
ins-186 or the NY107-ins-186/205 virus stocks, indicative of
replication fitness. For the NY107-ins-205 virus however, a
Glu186Gly substitution emerged in the HA after only two passages
in eggs following recovery from DNA transfection, indicating the
importance of the co-variant position 186 with respect to HA
functionality/glycan specificity. Altogether, the data indicates that
the H7 subtype avian influenza viruses that were circulating in
Table 1. Data collection and refinement statistics.
NY107
NY107+39SLN
NY107+69SLN
NY107+LSTb
Data collection
Space group
P212121
P212121
P212121
P212121
Cell dimensions (A˚)
66.96, 115.92, 251.61
67.80, 116.70, 249.84
66.60, 116.58, 250.68
67.08, 116.52, 251.95
Resolution (A˚)
50-2.6 (2.69-2.60)a
30-2.7 (2.80-2.70)
50-3.0 (3.11-3.0)
50-2.6 (2.69-2.60)
Rsym or Rmerge
10.6 (41.3)
14.6 (48.6)
14.3 (35.4)
12.2 (31.5)
I/s
39.6 (2.0)
24.3 (1.7)
34.2 (8.2)
40.5 (9.9)
Completeness (%)
99.2 (99.0)
99.3 (94.6)
92.3 (75.6)
91.3 (86.2)
Redundancy
7.2 (6.2)
5.8 (5.5)
4.9 (4.4)
10.9 (11.2)
Refinement
Resolution (A˚)
50-2.6 (2.67-2.60)
30-2.7 (2.77-2.70)
50-3.0 (3.08-3.00)
50-2.6 (2.67-2.60)
No. of reflections (total)
57285
51770
33421
53603
No. of reflections (test)
3053
2769
1779
2842
Rwork/Rfree
21.7/25.6
21.4/26.4
20.5/26.0
20.4/24.7
No. of atoms
11795
11878
11648
12108
r.m.s.d.- bond length (A˚)
0.006
0.006
0.008
0.006
r.m.s.d.- bond angle (u)
0.905
0.974
1.085
0.859
MolProbityb scores
Favored (%)
96.9
96.5
94.3
97.1
Outliers (%) (No. of residues)
0.1 (1/1434)
0.0 (0/1429)
0.1 (2/1433)
0.1 (2/1435)
aNumbers in parentheses refer to the highest resolution shell.
bReference [51].
doi:10.1371/journal.ppat.1001081.t001
Structure of a North American H7 Hemagglutinin
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aquatic birds and poultry in North America before 1996 exhibited a
classic avian a2-3 sialoside binding preference. In order for the 220-
loop deletion to be tolerated, concurrent Gly186Glu and Gly205Arg
substitutions in the vicinity of RBS of HA emerged to achieve a
restricted a2-3 binding profile and only a moderate/limited increase
in binding to branched di-sialyl a2-6 biantennary structures (#46–
48) as well the a2,6 internal sialoside, LSTb (#58).
NY107 avian receptor complex
To understand from a structural perspective how NY107
interacts with host receptors, we solved the structure of NY107
in complex with an avian and two human receptor analogs.
For the avian receptor analog, 39SLN, the electron density
maps revealed well-ordered features for the Sia-1, Gal-2, and
GlcNAc-3 in the NY107 HA complex structure (Figure 4A).
Structural comparison of NY107 HA binding to other, H1, H2,
H3, H5, and H9 subtypes (Figure S2A) revealed that 39SLN
binding to NY107 resembled binding of the other published
HAs. Indeed, the terminal Sia-1 moiety is positioned almost
identically in all structures, and forms the majority of hydrogen
bonds and contacts with residues in the RBS (Figure 4A and
Table S3).
Figure 1. NY107 HA monomer and comparison of its RBS to other HA structures. (A) One monomer is shown with the HA1 chain colored in
green and the HA2 chain in cyan. The location of the receptor binding site and the HA1/HA2 cleavage site are circled. (B) The superposition of
receptor binding domains of NY107 (green), Av-H7 (marine), 1918-Hu-H1 (magenta), Hu-H5 (yellow), Hu-H3 (orange), and Sw-H9 (grey). The proximity
of Arg220 and Gln226 are highlighted. Three structural elements comprising this binding site are labeled. The two major differences are the extended
150-loop and the deletion of 220-loop of NY107. (C) Overlap of NY107 (green) and Av-H7 (marine) (PDB: 1TI8) illustrates the compensatory effect of
R220 bringing it close to the position occupied by G228 in the avian HA. (D) Overlap of the NY107 (green), NY107- 39SLN (orange), NY107-69SLN (red),
and NY107-LSTb (magenta) structures. All the figures were generated and rendered with the use of MacPyMOL [56].
doi:10.1371/journal.ppat.1001081.g001
Structure of a North American H7 Hemagglutinin
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Published avian HA structures with an intact 220-loop form
very close interactions with Gal-2 of 39SLN via residue Gln226
which is important in receptor specificity and host adaptation. For
example, in the avian H7/39SLN HA structure it interacts with
Gal-2 O4 [31]. In the NY107 HA structure, although Gln226 is
absent and no other residue occupies the same space as Gln226
(Figure 1B), Arg220 does forms a hydrogen bond between Arg220
NH2 and Gal-2 O4 (Figure 4A). Interestingly, although there was
interpretable density for the GlcNAc-3 (Figure 4A and Figure
S4B), no hydrogen bonding was apparent between the HA and the
GlcNAc-3, which is consistent with other reported structures [32].
Thus, for binding to avian receptors, the trans conformation of
a2-3 linkages is essential and perhaps only the first two saccharides
are required. Indeed, due to the absence of 220-loop in the NY107
HA structure, the ‘‘aperture’’ of the RBS formed by 220-loop and
130-loop in regular HAs is increased by ,10 A˚ , so that the
branched, internal, and perhaps more complicated glycans might
be accommodated more efficiently.
NY107 human receptor complexes
In the NY107/69SLN complex, only Sia-1 and Gal-2 are
ordered (Figure 4B). The Sia-1 remains in the same position as
previously analyzed glycan/HA complexes from H1, H2, H3, H5,
and H9 (Figure S3B), whereas the Av-H7 complex structure with
Sialyllacto-N-tetraose c (LSTc) did not reveal any density for the
Sia-1 in the receptor binding site [31]. The Gal-2 position varies
significantly among different subtypes. Compared to the human-
adapted H1 HA [32], Gal-2 in the NY107 HA is 3A˚ higher, and
thus is further from the protein (Figure S3B). In NY107, the Gal-2
only forms an intramolecular, saccharide-saccharide interaction
with Sia-1. The poor electron density map and fewer interactions
with protein residues suggest that the cis conformation of a2-6
Figure 2. Receptor specificity of NY107 recHA and virus. Glycan microarray analysis of recombinant NY107 HA (A) and NY107 virus (B)
compared to the recHA (C) and virus (D) from a Eurasian lineage A/Netherlands/219/2003 H7 influenza virus that was circulating in the same year and
also infected a human. Colored bars highlight glycans that contain a2-3 SA (blue) and a2-6 SA (red), a2-6/a2-3 mixed SA (purple), N-glycolyl SA
(green), a2-8 SA (brown), b2-6 and 9-O-acetyl SA, and non-SA (grey). Error bars reflect the standard error in the signal for six independent replicates
on the array. Structures of each of the numbered glycans are found in Table S4.
doi:10.1371/journal.ppat.1001081.g002
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linkages in 69SLN trisaccharides show a reduced binding affinity
with NY107.
Glycan array results with NY107 revealed a strong binding
signal for the internal a2-6 sialoside, LSTb. To further investigate
this interaction, we solved the structure of the NY107/LSTb
complex. The final model contained Sia-1, NAG-2, Gal-3, and
Gal-5 in the RBS. Although glycan microarray data indicated
NY107 to have a specific affinity for LSTb, few interactions were
apparent from the crystal structure. Sia-1 still forms multiple
hydrogen bonds with residues in the RBS (Table S3 & Figure 4C).
The branched Gal-5 interacts with Ser137, to help stabilize the
LSTb binding. However, Arg220 and Lys193, the two residues
showing close binding with 39SLN, did not form any hydrogen
bonds with LSTb. In the structure, Gal-5 also interacts with a
crystal packing symmetry mate and thus the flexibility of whole
LSTb may be restricted. In solution, with more freedom, the
LSTb should be able to tilt closer to the RBS, and thus Glc-4 may
have more interactions with the 190-helix than seen in the crystal
structure.
Discussion
Human infections by avian influenza viruses, including H7
subtypes, continue to pose a major public health threat. Although
the
species
barrier
prevents
avian
influenza
viruses
from
widespread infection of the human population, the molecular
determinants of efficient interspecies transmission and pathoge-
nicity are still poorly understood. The viral coat protein HA
however, is perhaps a critical molecule since previous pandemic
viruses modified their receptor specificity and overcame the
interspecies barrier to spread in the human population. Although
HA structures alone and in complex with receptor analogs provide
considerable insight into receptor binding, it is clear that HAs from
different species and subtypes have significant structural variation.
Indeed, low-pathogenic H7N2 avian influenza viruses with an 8
amino acid deletion within its RBS started to circulate in live-bird
markets in the northeast United States in 1996. Despite what one
would consider a debilitating mutation, these viruses have been
reported as the predominant isolate [33]. Whether such a deletion
contributed to their evolutionary success and how are an
important questions, especially in light of NY107’s ability to
produce respiratory illness in humans [16], as well as its reported
increased affinity for human-type receptors and ability for contact
transmission in ferrets [21]. To try to help answer these questions,
we have analyzed the molecular structures of NY107 and its
complexes with receptor analogs to explain receptor specificity at
the molecular level.
The crystal structures of NY107 and its complexes with both
avian and human receptor analogs describe a mechanism as to
how an influenza virus might adapt by dramatically altering its
RBS, and still be functional. Arg220 of the HA1 chain of NY107
compensates for the loss of the 220-loop, by forming hydrogen
bonds with Gal-2 from the avian analog (binding was not observed
in either of the structures complexes with the human analogs).
However, in the LSTb complex, branched Gal-5 forms extra
interactions with the 130-loop, thus improving the binding
preference for this particular glycan. Consistent with the structural
evidence, glycan microarray analyses of NY107 revealed a strong
binding preference for the branched a2-6 sialoside, LSTb. Except
for the absence of the 220-loop, other key residues within the RBS
are conserved in NY107 and thus, direct interactions with sialic
acid are maintained.
The 220-loop is recognized as one of the three crucial structural
elements in the RBS. Aside from the North American lineage
H7N2 viruses, which have been circulating with a deletion (221–
228) in this loop, there has been one other report describing a
seven amino acid deletion (224–230) in a laboratory generated
H3N2 escape mutant which was reported to have a slightly
Table 2. Comparison of the sialoside receptor specificity of the HAs from H7 influenza viruses.
Glycan Group
Graph
Numbera
NY107
RecHA
NY107
Virus
NY107-ins
Virus
NY107-ins
E186G Virus
NY107-ins
R205G Virus
NY107-ins E186G/
R205G Virus
NL219
RecHA
NL219
Virus
a2-3
Sulfated
4–8
+++b
+++
+++
+++
+++
+++
+++
+++
Branched
9–11
+++
+++
+
+++
+++
+++
+++
+++
Linear
12–27
+
+++
+
+++
+++
+++
+++
+++
Fucosylated
28–34
2
+++
+++
+++
+++
+++
+++
+++
a2-6
Sulfated
41
2
2
2
2
2
2
2
2
Branched mono-sialyl
42–45, 49
2
2
2
2
2
2
2
2
Branched di-sialyl
46–48
2
+++
2
2
2
2
2
2
Linear
50–56
2
2
2
2
2
2
2
2
Internal
58–59
+++
+++
2
2
2
2
2
2
Other
Sialic acid
1–2
2
+++
+
2
2
2
2
2
a2-3/a2-6 Branched
60–64
2
2
2
+++
+++
+++
+++
+++
Neu5Gcc
65–72
2
2
2
+++
+++
+++
2
+++
aMembers of each group are identified according to the graph number used in the microarray data in Figures 2 and 3 and correspond to numbers in the complete
glycan list (Table S4).
bBinding of samples to glycan subclasses are qualitatively estimated based on relative strength of the signal for the data shown in Figures 2 and 3: strong (+++), weak
(+), absent (2).
cN-glycolylneuraminic acid.
doi:10.1371/journal.ppat.1001081.t002
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increased affinity for a2-3-linked glycans by hemagglutination
assay [34]. Meanwhile, the equivalent region in the hemaggluti-
nin-esterase-fusion (HEF) protein of influenza C virus reveals a
rearrangement resulting in a truncated 260-loop in its RBS (Figure
S5) [35]. However, without structural data with appropriate
receptor analogs, it is not possible to compare the role of these loop
variants in receptor binding to the H7 HA structure described
here.
When compared to NL219, another co-circulating H7 avian
virus HA (Figure 2C and D), overall binding to a2-3-linked
glycans was markedly reduced, while increased binding to a2-6-
linked receptors was only marginal. However, these results focus
attention on only 2 sub-classes of human-type receptors that may
be important for infection (and transmission in ferrets). The
NY107 virus interaction with biantennary glycans (Figure 2B),
although weak (not seen in Figure 2A with recHA), is a possible
route for virus entry as biantennary structures are common on
tissues, i.e. glycan profiling data from human lung tissue on the
Consortium for Functional Glycomics (CFG) web site. In addition,
the internal sialoside, LSTb, was observed in both virus and
recHA microarray data, suggesting this type of glycan has good
affinity for this HA. The significance of this is unknown since
LSTb has only been described in human milk [36].
Interestingly, NY107 and NL219 virus receptor binding and
specificity has been addressed previously using glycan microarray
analysis that reported a significantly increased preference for a2-6
and decreased preference for a2-3-linked sialosides [22]. In
addition, the same viruses were also included in a recent study
from Gambaryan et al. using a competitive solid-phase binding
assay [23]. Our findings confirm and extend the receptor binding
specificity reported by these authors in that they reported both
viruses binding to sulfated sialylglycans with a lactosamine (Galb1-
4GlcNAc core and reported only a moderate binding affinity for
a2-6-sialyllactosamine, the human-type receptor analog used in
their assay.
The 220-loop is an integral feature of the receptor binding site,
and thus one would predict that such a deletion might have
compromised this strain to be deleted from the population of
circulating viruses. However, this was not the case [33] and its
existence appears to be in part due to the additional mutations at
Figure 3. Effect of 220-loop deletion and additional RBS mutations on NY107 receptor specificity. NY107 was engineered to restore the
220-loop to a consensus full-length HA from 1996 (A) and additional co-variant amino acid substitutions, Glu186Gly (B), Arg205Gly (C) and the double
mutant Glu186Gly/Arg205Gly (D) to restore, on the NY107 framework, an HA RBS found in viruses prior to the introduction of the deletion in North
American viruses. Colored bars group glycans as described in Figure 3. Error bars reflect the standard error in the signal for six independent replicates
on the array. Structures of each of the numbered glycans are found in Table S4.
doi:10.1371/journal.ppat.1001081.g003
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positions 186 and 205. Restoration of the loop with either or both
residues mutated back to the pre-1994 consensus sequence
resulted in a classic avian influenza virus binding profile. The
emergence of the Glu186Gly mutation in the HA of the NY107-
ins-205 mutant after only two passages of the rescued virus in
eggs, also indicates the importance of these positions for HA
functionality/glycan specificity. Analysis of the structural data
reveals that positions 186 and 205 are on opposite sides of a
monomer but are both close to the 220-loop deletion region in
the trimeric form. The Glu at position 186 is close to Arg220 and
may interact with Arg220 when binding avian receptors. Position
205 in the neighboring monomer may be important in trimer
stability and maintaining RBS functionality. If one models the pre-
1996 220-loop restored into the NY107 structure, Arg205, Glu186
and the loop all clash, thus explaining the Glu186Gly mutation
that emerged in the NY107-ins-205 virus HA after limited egg
passage.
The NY107 RBS with its more restricted a2-3 glycan binding
preference and weak/moderate increase in a2-6 binding may have
given the virus a selective advantage to be maintained in poultry at
live bird markets and supplying farms. Certain terrestrial birds,
such as quails and chickens, have recently been shown to present
both human and avian types of receptors in the trachea and
intestine [37,38,39]. Although it is not known what specific glycans
are presented in these animals, it is conceivable that a virus with
mixed specificity might have a distinct advantage over avian
viruses that have specific avian receptor requirements, particularly
in bird markets where multiple species coalesce. Previous results
with H7N2, H9N2 and H5N1 viruses all highlight the fact that an
increase in a2-6-binding preference is not sufficient for efficient
transmission of avian influenza viruses to humans [22,40,41].
Although it remains to be seen whether prolonged circulation of
viruses in terrestrial birds, such as domestic chickens, can provide a
possible route for viruses to adapt for efficient human infection
[11], continued surveillance of influenza viruses from avian and
other animal reservoirs is urgently needed to define their zoonotic
potential.
Materials and Methods
Cloning
Based on H3 numbering [42], cDNA corresponding to residues
11–329 (HA1) and 1–176 (HA2) of the ectodomain of the
hemagglutinin
(HA)
from
A/New
York/107/2003
(H7N2;
Genbank:ACC55270)
and
A/Netherlands/219/2003
(H7N7;
Genebank: AAR02640) was cloned into the baculovirus transfer
vector, pAcGP67-A (BD Biosciences), incorporating a C-terminal
thrombin cleavage site, a ‘‘foldon’’ sequence [43] and a His-tag at
the extreme C-terminus of the construct to enable protein
purification [25,44]. Transfection and virus amplification were
carried out according to the baculovirus expression system manual
(BD Biosciences Pharmingen).
Figure 4. Glycan interactions within the NY107 RBS. The top panel shows the interactions of NY107 with (A) 39SLN, (B) 69SLN and (C) LSTb.
NY107 is shown in orange/red/magenta cartoon respectively. The interacting HA residues are shown as green sticks. The bottom panel shows the
electron density map of the ligands. The NY107 is shown in the same colors as above, and the ligands are shown as green sticks, the 2fo-fc electron
density maps (contoured at 1s) are shown in grey. Simulated annealing omit maps are shown in supplementary Figure S4.
doi:10.1371/journal.ppat.1001081.g004
Structure of a North American H7 Hemagglutinin
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September 2010 | Volume 6 | Issue 9 | e1001081
Protein expression and purification
Soluble NY107 was recovered from the cell supernatant by metal
affinity chromatography using Ni-NTA resin (Qiagen Inc.). Frac-
tions containing NY107 were pooled and dialyzed against 10 mM
Tris-HCl, 50 mM NaCl, pH 8.0, then subjected to ion-exchange
chromatography (IEX) using a Mono-Q HR 10/10 column (GE
Healthcare). IEX purified NY107 was subjected to thrombin digest
(3 units/mg protein; overnight at 4uC) and purified by gel filtra-
tion chromatography using a Superdex-200 16/60 column (GE
Healthcare) and 50 mM Tris-HCl, 100 mM NaCl, pH 8.0 as
running buffer. Protein eluting as a trimer was buffer exchanged
into 10 mM Tris-HCl, 50 mM NaCl, pH 8.0 and concentrated to
14.5 mg/ml for crystallization trials. At this stage, the protein
sample still contained the additional plasmid-encoded residues at
both the N (ADPG) and C terminus (SGRLVPR).
Crystallization, ligand soaking and data collection
Initial crystallization trials were set up using a Topaz Free
Interface Diffusion (FID) Crystallizer system (Fluidigm Corpora-
tion, San Francisco, CA). Crystals were observed in several
conditions containing PEG 3350 or PEG 4000. Following opti-
mization, diffraction quality crystals for NY107 were obtained at
room temperature using a modified method for microbath under
oil [45], by mixing the protein with reservoir solution containing
20% PEG 3350, 0.2 M magnesium chloride at pH 7.2. For
receptor analog complexes, crystals were soaked for 3 hours in the
crystallization buffer containing 10 mM 39SLN or 69SLN (V-labs
Inc., Covington, LA), or overnight in 10mM LSTb (Sigma, St.
Louis, MO). All crystals were flash-cooled at 100K using 20%
glycerol as the cryo-protectant. Datasets were collected at
Advanced Photon Source (APS) beamlines 22 ID and BM at
100K. Data were processed with the DENZO-SACLEPACK suite
[46]. Statistics for data collection are presented in Table 1.
Structure determination and refinement
The
structure
of
NY107
was
determined
by
molecular
replacement with Phaser [47] using the structure of the avian
H7 (Av-H7) from A/turkey/Italy/2002, pdb:1TI8 (HA1, 78%
identity; HA2, 90% identity) as the searching model. One HA
trimer occupies the asymmetric unit with an estimated solvent
content of 58% based on a Matthews’ coefficient (Vm) of 2.9 A˚ 3/
Da. Rigid body refinement of the trimer led to an overall R/Rfree
of 28.6%/37.4%. The model was then ‘‘mutated’’ to the correct
sequence and rebuilt by Coot [48], then the protein structures
were refined with REFMAC [49] using TLS refinement [50]. The
final models were assessed using MolProbity [51]. The three
complex structures were refined and evaluated using the same
strategy. All statistics for data processing and refinement are
presented in Table 1. Electron density maps (2fo-fc) were
generated in Refmac [49] while simulated annealing omit maps
were generated by sa-omit-map, a part of the Crystallography and
NMR System (CNS) software [52].
Virus generation
Wild type and mutant viruses of NY107 (H7N2) and A/
Netherland/219/2003 (H7N7) were generated from plasmids by a
reverse genetics approach [53]. To generate viruses with amino
acid insertion or substitution in the HA, mutations were
introduced into plasmid DNA with an overlap extension PCR
approach [54]. Viruses derived by plasmid transfection of HK293
cells were propagated in eggs. The genomes of resulting virus
stocks were sequenced to detect the emergence of possible variants
during amplification.
Glycan binding analyses
Glycan microarray printing and recHA analyses have been
described previously [24,30,44,55] (see Table 2 for glycans used
for analyses in these experiments). Virus were analyzed on the
microarray as described previously [30].
PDB accession codes
The atomic coordinates and structure factors of NY107 are
available from the RCSB PDB under accession codes 3M5G for
the unliganded NY107, 3M5H for the NY107 with 39-SLN and
3M5I and 3M5J for NY107 with 69SLN and LSTb, respectively.
Accession/ID numbers for genes/proteins used in this
work
A/New York/107/03 (H7N2), Genbank: ACC55270; A/
Netherlands/219/03 (H7N7), Genbank: AAR02640; A/Hong
Kong/1-9/68 (H3N2), 2HMG; A/Duck/Ukraine/1/63 (H3N8),
PDB: 1MQL; A/South Carolina/1/18 (H1N1), PDB: 1RD8; A/
Puerto Rico/8/34 (H1N1), PDB: 1RU7; A/Swine/Iowa/15/30
(H1N1), PDB: 1RUY; A/Singapore/1/1957
(H2N2), PDB:
2WRC; A/Viet Nam/1203/04 (H5N1), PDB: 2FK0; A/Duck/
Singapore/3/97 (H5N3), PDB: 1JSM; A/Swine/Hong Kong/9/
98 (H9N2), PDB: 1JSD; A/Turkey/Italy/8000/02 (H7N3), PDB:
1TI8; C/Johannesburg/1/66, 1FLC.
Supporting Information
Figure S1
Sequence alignment of selected structurally available
HAs. Human H3 (PDB: 2HMG), Avian H3 (PDB: 1MQL), 1918-
Human H1 (PDB: 1RD8), 1934-Human H1 (PDB: 1RU7), Swine
H1 (PDB: 1RUY), 1957-Huamn H2 (PDB: 2WRC), Human H5
(PDB: 2FK0), Avian H5 (PDB: 1JSM), Swine H9 (PDB: 1JSD),
and Avian H7 (PDB: 1TI8) were used in the alignments. The
fusion domain of HA1 is highlighted in magenta, the vestigial
esterase domain is highlighted in green, the receptor binding
domain is highlighted in blue, and the fusion domain of HA2 is
highlighted in red. Residue numbering is based on the H3 HA
sequence.
Found at: doi:10.1371/journal.ppat.1001081.s001 (2.84 MB TIF)
Figure S2
Expression and purification of NY107. SDS-PAGE
reveals that NY107 was expressed as the HA0 form with a mass
approximately 60kDa (middle lane). Thrombin cleavage resulted in
an unexpected reduction in band size to a HA1/HA2 profile (right
lane) with possible multiple glycoforms for the HA2 clearly present.
Found at: doi:10.1371/journal.ppat.1001081.s002 (0.23 MB TIF)
Figure S3
Comparison of glycan binding to NY107 with other
HAs. A. Overlap of a2-3 ligands binding in the receptor binding
site from NY-H7 (green), Av-H3 (orange), 1930-Hu-H1 (magenta),
1957-Hu-H2 (cyan), Av-H5 (yellow), and Sw-H9 (grey). B.
Overlap of a2-6 linkage ligands binding in the receptor binding
site from NY-H7 (green), Av-H3 (orange), 1930-Hu-H1 (magenta),
1957-Hu-H2 (cyan), Av-H5 (yellow), and Sw-H9 (grey).
Found at: doi:10.1371/journal.ppat.1001081.s003 (2.55 MB TIF)
Figure S4
Simulated annealing omit maps of the receptor
binding site (contoured at 1s). A. NY107 (blue), B. NY107-39SLN
(orange), C. NY107-69SLN (red), and D. NY107-LSTb (magenta).
The protein model is shown in cartoon, and the residues involved
in the binding to receptor analogs were shown in sticks. Maps were
generated using version 1.2 of the Crystallography and NMR
System (CNS) software.
Found
at:
doi:10.1371/journal.ppat.1001081.s004
(1.93
MB
TIF)
Structure of a North American H7 Hemagglutinin
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9
September 2010 | Volume 6 | Issue 9 | e1001081
Figure S5
Comparison of NY107 RBS to HEF. Overlap of RBS
from NY107 (green), Av-H7 (marine) and HEF (magenta).
Found at: doi:10.1371/journal.ppat.1001081.s005 (1.12 MB TIF)
Table S1
Comparison of r.m.s.d. (A˚ ) for different HA domains.
For analyzing differences in the overall structure, r.m.s.d. values
were calculated between monomers or domains of different HA’s,
after the Ca atoms of the HA2 domains were superposed by
sequence and structural alignment onto the equivalent domains of
NY107.
Found
at:
doi:10.1371/journal.ppat.1001081.s006
(0.04
MB
DOC)
Table S2
Comparison of r.m.s.d. (A˚ ) for individual domains.
Each domain was superimposed separately to determine how the
individual NY107 domains compared to equivalent domains in the
other structures.
Found
at:
doi:10.1371/journal.ppat.1001081.s007
(0.04
MB
DOC)
Table S3
Molecular interactions between NY107 and receptor
analogs. The hydrogen bond cutoff is 3.8 A˚
for the listing
interactions.
Found
at:
doi:10.1371/journal.ppat.1001081.s008
(0.07
MB
DOC)
Table S4
Glycan array differences between NY107, the fully
restored NY107-ins, and NL219 (virus and rHA). The color
coding in the left hand column reflects the same coloring scheme
used in Figures 2 and 3. Significant binding of samples to glycans
are qualitatively estimated based on relative strength of the signal
for the data shown in Figures 2 and 3 Strong (+++), weak (+).
Found
at:
doi:10.1371/journal.ppat.1001081.s009
(0.19
MB
DOC)
Acknowledgments
The authors would like to thank the staff of SER-CAT sector 22 for their
help in data collection. We also thank WHO Global Influenza Surveillance
Network for providing NY107 and NL219 viruses from which the reverse
genetics viruses were generated. Glycan microarray data presented here
will be made available on-line through the CFG web site upon publication
(www.functionalglycomics.org). The findings and conclusions in this report
are those of the authors and do not necessarily represent the views of the
Centers for Disease Control and Prevention or the Agency for Toxic
Substances and Disease Registry.
Author Contributions
Conceived and designed the experiments: HY LMC PJC ROD JS.
Performed the experiments: HY LMC PJC JS. Analyzed the data: HY
LMC PJC ROD JS. Wrote the paper: HY LMC PJC ROD JS.
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Structure of a North American H7 Hemagglutinin
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|
3M5H
|
Crystal structure of a H7 influenza virus hemagglutinin complexed with 3SLN
|
Structures of Receptor Complexes of a North American
H7N2 Influenza Hemagglutinin with a Loop Deletion in
the Receptor Binding Site
Hua Yang, Li-Mei Chen, Paul J. Carney, Ruben O. Donis, James Stevens*
Influenza Division, Centers for Disease Control and Prevention, Atlanta, Georgia, United States of America
Abstract
Human infections with subtype H7 avian influenza viruses have been reported as early as 1979. In 1996, a genetically stable
24-nucleotide deletion emerged in North American H7 influenza virus hemagglutinins, resulting in an eight amino acid
deletion in the receptor-binding site. The continuous circulation of these viruses in live bird markets, as well as its
documented ability to infect humans, raises the question of how these viruses achieve structural stability and functionality.
Here we report a detailed molecular analysis of the receptor binding site of the North American lineage subtype H7N2 virus
A/New York/107/2003 (NY107), including complexes with an avian receptor analog (39-sialyl-N-acetyllactosamine, 39SLN)
and two human receptor analogs (69-sialyl-N-acetyllactosamine, 69SLN; sialyllacto-N-tetraose b, LSTb). Structural results
suggest a novel mechanism by which residues Arg220 and Arg229 (H3 numbering) are used to compensate for the deletion
of the 220-loop and form interactions with the receptor analogs. Glycan microarray results reveal that NY107 maintains an
avian-type (a2-3) receptor binding profile, with only moderate binding to human-type (a2-6) receptor. Thus despite its
dramatically altered receptor binding site, this HA maintains functionality and confirms a need for continued influenza virus
surveillance of avian and other animal reservoirs to define their zoonotic potential.
Citation: Yang H, Chen L-M, Carney PJ, Donis RO, Stevens J (2010) Structures of Receptor Complexes of a North American H7N2 Influenza Hemagglutinin with a
Loop Deletion in the Receptor Binding Site. PLoS Pathog 6(9): e1001081. doi:10.1371/journal.ppat.1001081
Editor: Fe´lix A. Rey, Institut Pasteur, France
Received March 16, 2010; Accepted July 28, 2010; Published September 2, 2010
This is an open-access article distributed under the terms of the Creative Commons Public Domain declaration which stipulates that, once placed in the public
domain, this work may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose.
Funding: This work was funded by the Centers for Disease Control and Prevention. Use of the Advanced Photon Source at Argonne National Laboratory was
supported by the U. S. Department of Energy, Office of Science, Office of Basic Energy Sciences, under Contract No. DE-AC02-06CH11357. Glycan microarrays as
well as glycans for direct binding experiments were produced for the Centers for Disease Control by the CFG funded by National Institute of General Medical
Sciences Grant GM62116. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: fwb4@cdc.gov
Introduction
Influenza is an acute respiratory virus that infects up to 20% of
the population in the United States, resulting in ,36,000 deaths
annually [1,2]. The two membrane glycoproteins on the surface of
influenza A virus, hemagglutinin (HA), which functions as the
receptor binding and membrane fusion glycoprotein in cell entry,
and neuraminidase (NA), which functions as the receptor destroying
enzyme in virus release, form the basis for defining subtypes [3]. To
date, 16 HA (H1–H16) and 9 NA (N1–N9) have been identified in
avian species [4], while in the last century, only three subtypes,
H1N1 in 1918 and 2009, H2N2 in 1957, and H3N2 in 1968 [5,6,7],
have successfully adapted to humans. Hemagglutinin binds to sialic
acid (SA) glycans present on host cell surfaces. The receptors on
epithelial cells of the human upper respiratory tract are mainly
a2-6-linked SA moieties [8]. Since avian influenza viruses pre-
dominately bind a2-3-linked SA, and human influenza viruses
preferentially bind to a2-6-linked SA, human infection by avian
influenza viruses is rare [9]. However, since 1997 a growing number
of human cases of avian influenza infection have been reported [10],
including H5N1, H7N2, H7N3, H7N7, and H9N2 strains [11].
Although the current situation with the pandemic H1N1 influenza
virus dominates public health efforts, the prospect of a novel
pandemic emerging from these isolated cases continues to be a
major public health threat around the world.
Early cases of human infection by H7 influenza viruses are reported
as far back as 1979 [12,13]. Since 2002, multiple outbreaks and
human infections of H7 subtype viruses; within both Eurasian and
North American lineages have been reported. In the Netherlands in
2003, a highly pathogenic avian influenza (HPAI) H7N7 outbreak
resulted in more than 80 cases of human infections, including one
fatality [14,15]. In New York in 2003, a single case of human
respiratory infection of H7N2 was reported [16] and in British
Columbia in 2004, an H7N3 virus caused two cases of conjunctivitis
[17,18]. More recently in 2007, the United Kingdom reported several
cases of low pathogenic avian influenza (LPAI) H7N2 virus infections
that caused influenza-like illness and conjunctivitis [19].
Since 1996, H7 viruses of the North American lineage have
been circulating in regional live bird markets [20], containing a
24-nucleotide deletion resulting in an eight amino acid deletion in
the receptor-binding site (RBS) of HA (Figure S1). The recent
human infections with H7 in North America have raised public
health concerns as to how these viruses adapt to such a dramatic
structural change while remaining one of the predominant
circulating viral strains. A recent study of H7 viruses isolated
from previous outbreaks revealed efficient replication in both
mouse and ferret animal models [21]. In particular, ferret studies
with A/New York/107/2003 (NY107), an H7N2 virus isolated
from a man in New York, not only showed efficient replication in
the upper respiratory tract of the ferret but also the capacity for
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intra-species transmission by direct contact [21,22]. Interestingly,
both an increased preference for a2-6 and decreased preference for
a2-3-linked sialosides of this virus compared to the other avian
influenza viruses was shown by previous glycan microarray analysis
but less so by a competitive solid-phase binding assay [22,23].
Here we report a detailed molecular analysis of the RBS of the
HA from North American lineage H7N2 virus, NY107, including
glycan microarray analyses and structural analyses of the HA in
complex with an avian receptor analog (39-Sialyl-N-acetyllactosa-
mine, 39SLN) and two human receptor analogs (69-Sialyl-N-
acetyllactosamine, 69SLN; Sialyllacto-N-tetraose b, LSTb). These
results provide important insight into the interaction of H7 HAs
with both avian and human hosts.
Results
Overall structure
By using x-ray crystallography, the structure of H7 HA from the
NY107 virus was determined to 2.6 A˚ resolution (Table 1). In
addition, we also report three H7 HA receptor complex structures,
with avian receptor analog (39SLN) to 2.7 A˚ resolution and with
human receptor analogs (69SLN and LSTb) to 3.0 A˚ and 2.6 A˚
resolution, respectively (Table 1). The overall structure of NY107
is similar to other reported HA structures with a globular head
containing the RBS and vestigial esterase domain, and a
membrane proximal domain with its distinctive, central helical
stalk and HA1/HA2 cleavage site (Figure 1A). Although five
asparagine-linked glycosylation sites are predicted in the NY107
HA monomer, interpretable electron density was observed at only
two sites, Asn38 in HA1 and Asn82 in HA2 (all residue numbers
are based on H3 numbering). At these sites, only one or two N-
acetyl glucosamines could be interpreted.
During viral replication, HA is synthesized as a single chain
precursor (HA0) and cleaved by specific host proteases into the
infectious HA1/HA2 form. In baculovirus expression systems,
highly pathogenic HAs, with a polybasic cleavage site, are
expressed as an HA1/HA2 form [24], whereas HAs with
monobasic cleavage sites (single Arg) from low pathogenic viruses
are expressed as the HA0 form [25]. NY107 is regarded as a low
pathogenic virus, and as expected, was produced in the HA0
form (Figure S2). However, subsequent digestion with thrombin
protease to remove the His-tag resulted in cleavage to a profile on
SDS-PAGE comparable to that of an HA1/HA2 form (Figure S2).
A comparison of the NY107 cleavage site with the consensus
cleavage pattern in the MEROPS database (http://merops.
sanger.ac.uk) suggests it to be a possible thrombin cleavage site.
Based on their molecular phylogenies, HAs are divided into two
groups and five clades: group 1 includes H8, H9, and H12; H1,
H2, H5, and H6; H11, H13 and H16; group 2 includes H3, H4,
and H14; H7, H10 and H15 [26]. Among all available HA
structures, we selected ten representative HAs from both avian and
human subtypes for structural analysis. As expected, NY107 HA is
structurally very similar to the Avian-H7 in all comparisons and
closely related to H3, the other group 2 members used in the
analyses (Tables S1 and S2).
The receptor binding site
The RBS is at the membrane distal end of each HA monomer
and its specificity for sialic acid and the nature of its linkage to a
vicinal galactose residue is a major determinant of host range-
restriction. The consensus RBS for all current HAs is composed of
three major structural elements: a 190-helix (residues 188–194), a
220-loop (residues 221–228), and a 130-loop (residues 134–138).
In addition, highly conserved residues (Tyr98, Trp153, His183,
and Tyr195) form the base of the pocket.
Although the NY107 RBS is similar to other subtypes (H1, H2,
H3, H5, and H9), a previously observed specific feature of H7 HAs,
is also observed in the NY107 150-loop region: two residues inserted
at position 158 result in this loop protruding more than 6A˚ towards
the binding site compared to other subtype HAs (Figure 1B and
Table S2) [27]. More interestingly, the eight amino acid deletion,
only found in the North American lineage H7s, from position 221 to
228 (Figure S1), resulted in a complete loss of the 220-loop
(Figure 1B). Sequence alignment shows that Arg220 and Arg229 are
conserved in all influenza A HA subtypes (Figure S1), but structural
alignment of NY107 HA shows Arg220 occupying the Gly228
position, and the much shorter loop turns at residue Pro217
(Figure 1C). The Ca distance between NY107 Arg220 and its
homolog in the Av-H7 structure (PDB: 1TI8) [27] is 5.8A˚ , and they
point in opposite directions (Figure 1C). The side chain direction of
Av-H7 Arg220 is almost parallel with the beta sheet after Arg229,
whereas the NY107 Arg220 points downward to the binding pocket.
The Ca position of Arg229 in both H7 structures remains the same,
except the side chain in the NY107 swings away by about 5.9A˚
(Figure 1C) and could help to stabilize this region by forming a
hydrogen bond to the mainchain carbonyl of Gln210 in the
neighboring monomer. In the absence of the 220-loop in NY107
HA, upon glycan binding the long side chain of Arg220 compensates
for its loss and is displaced 4A˚ upward to form hydrogen bonds with
receptor analogs inside the binding pocket (Figure 1D).
Effect of loop truncation on the receptor binding
specificity of NY107
Previously, mutations in the HA receptor binding domains of
H1N1 (Glu190Asp/Gly225Asp) and H2N2/H3N2 (Gln226Leu
and Gly228Ser) subtypes were responsible for adaptation of these
viruses to pandemic strains [24,28,29,30]. Due to missing residues
221–228 in the NY107 HA RBS, neither mechanism for
adaptation is possible. Thus, in order to look more closely at the
role of the missing loop and its effect on receptor specificity, we
first subjected the recombinant HA (recHA) to glycan microarray
analyses and compared it to a reverse genetics-derived NY107
virus, and a co-circulating Eurasian virus and recHA, A/
Author Summary
Influenza virus adaptation to different hosts usually results
in a switch in receptor specificity of the viral surface coat
protein, hemagglutinin. Indeed, the hemagglutinin sub-
types from the last two human influenza pandemics of the
20th Century (H2 in 1957 and H3 1968) both adapted
successfully to human-type receptor specificity through
only two amino acid mutations in the receptor binding
pocket (Glutamine226RLeucine and Glycine228RSerine).
The recent human infections reported with other avian
subtypes such as H5, H7 and H9 have raised public health
concerns and focused efforts on identifying potential
subtypes from which a future pandemic strain may
emerge. Since 1996, H7 viruses of the North American
lineage have been circulating in regional live bird markets,
containing an eight amino acid deletion in the receptor-
binding site of HA. Here we report a detailed structural
analysis of the receptor binding site of a hemagglutinin
from the North American lineage of H7N2 viruses, in
complex with avian and human receptor analogs, to
understand how these viruses have adapted to such a
dramatic structural change in the binding site while
remaining one of the predominant circulating viral strains.
Structure of a North American H7 Hemagglutinin
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Netherlands/219/2003 (NL219), that has the consensus avian
sequence in the 220-loop and it also infected a human [15].
Glycan microarray analysis of recombinant NY107 (Figure 2A
and Table 2) revealed a highly restricted binding profile with
strong binding to only a2-3 sulfated (#4–8), a2-3 branched (#9–
11) and mixed a2-3/a2-6 branched sialosides (#60–64) as well as
to the long linear sialyl di- and tri-lactosamines (#22, 24). Weak
binding was also observed (above background) to other a2-3
glycans on the array. The recombinant NY107 also revealed a
strict glycan binding preference to only one a2-6 glycan, the
internal structure, Galb1-3(Neu5Aca2-6)GlcNAcb1-3Galb1-4Glc
(#58; LSTb) (Figure 2A), a glycan highlighted in a previous study
[22]. The virus with higher valency and avidity revealed stronger
binding to all a2-3 groups, in addition to the branched di-sialyl a2-
6 biantennary structures (#46–48) as well the LSTb (#58)
(Figure 2B and Table 2). In contrast, the NL219 recHA (Figure 2C
and Table 2) bound well to only the avian a2-3 containing sialyl-
glycans (sulfated, branched, linear and fucosylated). Its corre-
sponding virus also reflected this specificity although it also
revealed strong binding to a2-3 N-glycolylneuraminic acid
(Neu5Gc) containing glycans (#66–70) (Figure 2D and Table 2).
To further assess the effect of the missing 220 loop on HA
structural stability and receptor specificity it was essential to
evaluate these functions on the ancestral HA containing the full
length 220-loop. To this end, we engineered an HA with an avian
H7 consensus (PQVNGQSG) 220-loop re-introduced (NY107-
220ins) into the NY107 HA and recovered this virus by reverse
genetics. Compared to the NY107 virus (Figure 2A) glycan
microarray analyses of the resulting NY107-220ins virus (Figure 3A
and Table 2) revealed a decrease in binding to branched (#9–11)
and linear (#12–27) a2-3 sialosides and a loss of binding to the
branched di-sialyl a2-6 biantennary structures (#46–48), LSTb
(#58) as well as the mixed a2-3/a2-6 branched sialosides (#60–
64). In addition, sequence analysis of the NY107-220ins HA
revealed the presence of quasispecies in the second position of the
inserted loop, P(Q/K)VNGQSG, suggesting that re-introduction
of the loop alone is not tolerated and does not create an avian-type
binding profile. Thus other amino acid substitutions in the HA
might have co-evolved with the deletion of the 220 loop to help
stabilize the RBS/HA to maintain functionality.
When viruses containing this 220-loop deletion emerged in North
America in the mid 90’s, four additional amino acid substitutions,
Gly114Arg, Asp119Gly, Gly186Glu and Gly205Arg, in the HA1 as
well as an Asp19Asn in the HA2 chain were also introduced to most
of the circulating isolates. Of these, Gly186Glu and Gly205Arg in
the HA1 are close to the RBS, at the monomer interface, and could
potentially modulate its structure and/or function. NY107 viruses
with a restored consensus 220-loop and a single Glu186Gly (NY107-
ins-186) or Arg205Gly (NY107-ins-205) substitution as well as the
Glu186Gly/Arg205Gly double substitution (NY107-ins-186/205)
were derived by reverse genetics and evaluated. Glycan microarray
analysis for the three resulting viruses revealed similar glycan
binding profiles with increased binding to a2-3 sialosides, including
mixed a2-3/a2-6 branched sialosides (#60–64), a2-3 Neu5Gc
(#66–70), but limited binding to the a2,6 sialosides (Figures 3B, 3C,
3D), resulting in a binding profile virtually identical to that of the
NL219 virus and other avian influenza viruses (Figure 2D) [30].
Sequence analysis of the three reverse genetics derived viruses
revealed no mutations/quasispecies in the HAs of either the NY107-
ins-186 or the NY107-ins-186/205 virus stocks, indicative of
replication fitness. For the NY107-ins-205 virus however, a
Glu186Gly substitution emerged in the HA after only two passages
in eggs following recovery from DNA transfection, indicating the
importance of the co-variant position 186 with respect to HA
functionality/glycan specificity. Altogether, the data indicates that
the H7 subtype avian influenza viruses that were circulating in
Table 1. Data collection and refinement statistics.
NY107
NY107+39SLN
NY107+69SLN
NY107+LSTb
Data collection
Space group
P212121
P212121
P212121
P212121
Cell dimensions (A˚)
66.96, 115.92, 251.61
67.80, 116.70, 249.84
66.60, 116.58, 250.68
67.08, 116.52, 251.95
Resolution (A˚)
50-2.6 (2.69-2.60)a
30-2.7 (2.80-2.70)
50-3.0 (3.11-3.0)
50-2.6 (2.69-2.60)
Rsym or Rmerge
10.6 (41.3)
14.6 (48.6)
14.3 (35.4)
12.2 (31.5)
I/s
39.6 (2.0)
24.3 (1.7)
34.2 (8.2)
40.5 (9.9)
Completeness (%)
99.2 (99.0)
99.3 (94.6)
92.3 (75.6)
91.3 (86.2)
Redundancy
7.2 (6.2)
5.8 (5.5)
4.9 (4.4)
10.9 (11.2)
Refinement
Resolution (A˚)
50-2.6 (2.67-2.60)
30-2.7 (2.77-2.70)
50-3.0 (3.08-3.00)
50-2.6 (2.67-2.60)
No. of reflections (total)
57285
51770
33421
53603
No. of reflections (test)
3053
2769
1779
2842
Rwork/Rfree
21.7/25.6
21.4/26.4
20.5/26.0
20.4/24.7
No. of atoms
11795
11878
11648
12108
r.m.s.d.- bond length (A˚)
0.006
0.006
0.008
0.006
r.m.s.d.- bond angle (u)
0.905
0.974
1.085
0.859
MolProbityb scores
Favored (%)
96.9
96.5
94.3
97.1
Outliers (%) (No. of residues)
0.1 (1/1434)
0.0 (0/1429)
0.1 (2/1433)
0.1 (2/1435)
aNumbers in parentheses refer to the highest resolution shell.
bReference [51].
doi:10.1371/journal.ppat.1001081.t001
Structure of a North American H7 Hemagglutinin
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aquatic birds and poultry in North America before 1996 exhibited a
classic avian a2-3 sialoside binding preference. In order for the 220-
loop deletion to be tolerated, concurrent Gly186Glu and Gly205Arg
substitutions in the vicinity of RBS of HA emerged to achieve a
restricted a2-3 binding profile and only a moderate/limited increase
in binding to branched di-sialyl a2-6 biantennary structures (#46–
48) as well the a2,6 internal sialoside, LSTb (#58).
NY107 avian receptor complex
To understand from a structural perspective how NY107
interacts with host receptors, we solved the structure of NY107
in complex with an avian and two human receptor analogs.
For the avian receptor analog, 39SLN, the electron density
maps revealed well-ordered features for the Sia-1, Gal-2, and
GlcNAc-3 in the NY107 HA complex structure (Figure 4A).
Structural comparison of NY107 HA binding to other, H1, H2,
H3, H5, and H9 subtypes (Figure S2A) revealed that 39SLN
binding to NY107 resembled binding of the other published
HAs. Indeed, the terminal Sia-1 moiety is positioned almost
identically in all structures, and forms the majority of hydrogen
bonds and contacts with residues in the RBS (Figure 4A and
Table S3).
Figure 1. NY107 HA monomer and comparison of its RBS to other HA structures. (A) One monomer is shown with the HA1 chain colored in
green and the HA2 chain in cyan. The location of the receptor binding site and the HA1/HA2 cleavage site are circled. (B) The superposition of
receptor binding domains of NY107 (green), Av-H7 (marine), 1918-Hu-H1 (magenta), Hu-H5 (yellow), Hu-H3 (orange), and Sw-H9 (grey). The proximity
of Arg220 and Gln226 are highlighted. Three structural elements comprising this binding site are labeled. The two major differences are the extended
150-loop and the deletion of 220-loop of NY107. (C) Overlap of NY107 (green) and Av-H7 (marine) (PDB: 1TI8) illustrates the compensatory effect of
R220 bringing it close to the position occupied by G228 in the avian HA. (D) Overlap of the NY107 (green), NY107- 39SLN (orange), NY107-69SLN (red),
and NY107-LSTb (magenta) structures. All the figures were generated and rendered with the use of MacPyMOL [56].
doi:10.1371/journal.ppat.1001081.g001
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Published avian HA structures with an intact 220-loop form
very close interactions with Gal-2 of 39SLN via residue Gln226
which is important in receptor specificity and host adaptation. For
example, in the avian H7/39SLN HA structure it interacts with
Gal-2 O4 [31]. In the NY107 HA structure, although Gln226 is
absent and no other residue occupies the same space as Gln226
(Figure 1B), Arg220 does forms a hydrogen bond between Arg220
NH2 and Gal-2 O4 (Figure 4A). Interestingly, although there was
interpretable density for the GlcNAc-3 (Figure 4A and Figure
S4B), no hydrogen bonding was apparent between the HA and the
GlcNAc-3, which is consistent with other reported structures [32].
Thus, for binding to avian receptors, the trans conformation of
a2-3 linkages is essential and perhaps only the first two saccharides
are required. Indeed, due to the absence of 220-loop in the NY107
HA structure, the ‘‘aperture’’ of the RBS formed by 220-loop and
130-loop in regular HAs is increased by ,10 A˚ , so that the
branched, internal, and perhaps more complicated glycans might
be accommodated more efficiently.
NY107 human receptor complexes
In the NY107/69SLN complex, only Sia-1 and Gal-2 are
ordered (Figure 4B). The Sia-1 remains in the same position as
previously analyzed glycan/HA complexes from H1, H2, H3, H5,
and H9 (Figure S3B), whereas the Av-H7 complex structure with
Sialyllacto-N-tetraose c (LSTc) did not reveal any density for the
Sia-1 in the receptor binding site [31]. The Gal-2 position varies
significantly among different subtypes. Compared to the human-
adapted H1 HA [32], Gal-2 in the NY107 HA is 3A˚ higher, and
thus is further from the protein (Figure S3B). In NY107, the Gal-2
only forms an intramolecular, saccharide-saccharide interaction
with Sia-1. The poor electron density map and fewer interactions
with protein residues suggest that the cis conformation of a2-6
Figure 2. Receptor specificity of NY107 recHA and virus. Glycan microarray analysis of recombinant NY107 HA (A) and NY107 virus (B)
compared to the recHA (C) and virus (D) from a Eurasian lineage A/Netherlands/219/2003 H7 influenza virus that was circulating in the same year and
also infected a human. Colored bars highlight glycans that contain a2-3 SA (blue) and a2-6 SA (red), a2-6/a2-3 mixed SA (purple), N-glycolyl SA
(green), a2-8 SA (brown), b2-6 and 9-O-acetyl SA, and non-SA (grey). Error bars reflect the standard error in the signal for six independent replicates
on the array. Structures of each of the numbered glycans are found in Table S4.
doi:10.1371/journal.ppat.1001081.g002
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linkages in 69SLN trisaccharides show a reduced binding affinity
with NY107.
Glycan array results with NY107 revealed a strong binding
signal for the internal a2-6 sialoside, LSTb. To further investigate
this interaction, we solved the structure of the NY107/LSTb
complex. The final model contained Sia-1, NAG-2, Gal-3, and
Gal-5 in the RBS. Although glycan microarray data indicated
NY107 to have a specific affinity for LSTb, few interactions were
apparent from the crystal structure. Sia-1 still forms multiple
hydrogen bonds with residues in the RBS (Table S3 & Figure 4C).
The branched Gal-5 interacts with Ser137, to help stabilize the
LSTb binding. However, Arg220 and Lys193, the two residues
showing close binding with 39SLN, did not form any hydrogen
bonds with LSTb. In the structure, Gal-5 also interacts with a
crystal packing symmetry mate and thus the flexibility of whole
LSTb may be restricted. In solution, with more freedom, the
LSTb should be able to tilt closer to the RBS, and thus Glc-4 may
have more interactions with the 190-helix than seen in the crystal
structure.
Discussion
Human infections by avian influenza viruses, including H7
subtypes, continue to pose a major public health threat. Although
the
species
barrier
prevents
avian
influenza
viruses
from
widespread infection of the human population, the molecular
determinants of efficient interspecies transmission and pathoge-
nicity are still poorly understood. The viral coat protein HA
however, is perhaps a critical molecule since previous pandemic
viruses modified their receptor specificity and overcame the
interspecies barrier to spread in the human population. Although
HA structures alone and in complex with receptor analogs provide
considerable insight into receptor binding, it is clear that HAs from
different species and subtypes have significant structural variation.
Indeed, low-pathogenic H7N2 avian influenza viruses with an 8
amino acid deletion within its RBS started to circulate in live-bird
markets in the northeast United States in 1996. Despite what one
would consider a debilitating mutation, these viruses have been
reported as the predominant isolate [33]. Whether such a deletion
contributed to their evolutionary success and how are an
important questions, especially in light of NY107’s ability to
produce respiratory illness in humans [16], as well as its reported
increased affinity for human-type receptors and ability for contact
transmission in ferrets [21]. To try to help answer these questions,
we have analyzed the molecular structures of NY107 and its
complexes with receptor analogs to explain receptor specificity at
the molecular level.
The crystal structures of NY107 and its complexes with both
avian and human receptor analogs describe a mechanism as to
how an influenza virus might adapt by dramatically altering its
RBS, and still be functional. Arg220 of the HA1 chain of NY107
compensates for the loss of the 220-loop, by forming hydrogen
bonds with Gal-2 from the avian analog (binding was not observed
in either of the structures complexes with the human analogs).
However, in the LSTb complex, branched Gal-5 forms extra
interactions with the 130-loop, thus improving the binding
preference for this particular glycan. Consistent with the structural
evidence, glycan microarray analyses of NY107 revealed a strong
binding preference for the branched a2-6 sialoside, LSTb. Except
for the absence of the 220-loop, other key residues within the RBS
are conserved in NY107 and thus, direct interactions with sialic
acid are maintained.
The 220-loop is recognized as one of the three crucial structural
elements in the RBS. Aside from the North American lineage
H7N2 viruses, which have been circulating with a deletion (221–
228) in this loop, there has been one other report describing a
seven amino acid deletion (224–230) in a laboratory generated
H3N2 escape mutant which was reported to have a slightly
Table 2. Comparison of the sialoside receptor specificity of the HAs from H7 influenza viruses.
Glycan Group
Graph
Numbera
NY107
RecHA
NY107
Virus
NY107-ins
Virus
NY107-ins
E186G Virus
NY107-ins
R205G Virus
NY107-ins E186G/
R205G Virus
NL219
RecHA
NL219
Virus
a2-3
Sulfated
4–8
+++b
+++
+++
+++
+++
+++
+++
+++
Branched
9–11
+++
+++
+
+++
+++
+++
+++
+++
Linear
12–27
+
+++
+
+++
+++
+++
+++
+++
Fucosylated
28–34
2
+++
+++
+++
+++
+++
+++
+++
a2-6
Sulfated
41
2
2
2
2
2
2
2
2
Branched mono-sialyl
42–45, 49
2
2
2
2
2
2
2
2
Branched di-sialyl
46–48
2
+++
2
2
2
2
2
2
Linear
50–56
2
2
2
2
2
2
2
2
Internal
58–59
+++
+++
2
2
2
2
2
2
Other
Sialic acid
1–2
2
+++
+
2
2
2
2
2
a2-3/a2-6 Branched
60–64
2
2
2
+++
+++
+++
+++
+++
Neu5Gcc
65–72
2
2
2
+++
+++
+++
2
+++
aMembers of each group are identified according to the graph number used in the microarray data in Figures 2 and 3 and correspond to numbers in the complete
glycan list (Table S4).
bBinding of samples to glycan subclasses are qualitatively estimated based on relative strength of the signal for the data shown in Figures 2 and 3: strong (+++), weak
(+), absent (2).
cN-glycolylneuraminic acid.
doi:10.1371/journal.ppat.1001081.t002
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increased affinity for a2-3-linked glycans by hemagglutination
assay [34]. Meanwhile, the equivalent region in the hemaggluti-
nin-esterase-fusion (HEF) protein of influenza C virus reveals a
rearrangement resulting in a truncated 260-loop in its RBS (Figure
S5) [35]. However, without structural data with appropriate
receptor analogs, it is not possible to compare the role of these loop
variants in receptor binding to the H7 HA structure described
here.
When compared to NL219, another co-circulating H7 avian
virus HA (Figure 2C and D), overall binding to a2-3-linked
glycans was markedly reduced, while increased binding to a2-6-
linked receptors was only marginal. However, these results focus
attention on only 2 sub-classes of human-type receptors that may
be important for infection (and transmission in ferrets). The
NY107 virus interaction with biantennary glycans (Figure 2B),
although weak (not seen in Figure 2A with recHA), is a possible
route for virus entry as biantennary structures are common on
tissues, i.e. glycan profiling data from human lung tissue on the
Consortium for Functional Glycomics (CFG) web site. In addition,
the internal sialoside, LSTb, was observed in both virus and
recHA microarray data, suggesting this type of glycan has good
affinity for this HA. The significance of this is unknown since
LSTb has only been described in human milk [36].
Interestingly, NY107 and NL219 virus receptor binding and
specificity has been addressed previously using glycan microarray
analysis that reported a significantly increased preference for a2-6
and decreased preference for a2-3-linked sialosides [22]. In
addition, the same viruses were also included in a recent study
from Gambaryan et al. using a competitive solid-phase binding
assay [23]. Our findings confirm and extend the receptor binding
specificity reported by these authors in that they reported both
viruses binding to sulfated sialylglycans with a lactosamine (Galb1-
4GlcNAc core and reported only a moderate binding affinity for
a2-6-sialyllactosamine, the human-type receptor analog used in
their assay.
The 220-loop is an integral feature of the receptor binding site,
and thus one would predict that such a deletion might have
compromised this strain to be deleted from the population of
circulating viruses. However, this was not the case [33] and its
existence appears to be in part due to the additional mutations at
Figure 3. Effect of 220-loop deletion and additional RBS mutations on NY107 receptor specificity. NY107 was engineered to restore the
220-loop to a consensus full-length HA from 1996 (A) and additional co-variant amino acid substitutions, Glu186Gly (B), Arg205Gly (C) and the double
mutant Glu186Gly/Arg205Gly (D) to restore, on the NY107 framework, an HA RBS found in viruses prior to the introduction of the deletion in North
American viruses. Colored bars group glycans as described in Figure 3. Error bars reflect the standard error in the signal for six independent replicates
on the array. Structures of each of the numbered glycans are found in Table S4.
doi:10.1371/journal.ppat.1001081.g003
Structure of a North American H7 Hemagglutinin
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7
September 2010 | Volume 6 | Issue 9 | e1001081
positions 186 and 205. Restoration of the loop with either or both
residues mutated back to the pre-1994 consensus sequence
resulted in a classic avian influenza virus binding profile. The
emergence of the Glu186Gly mutation in the HA of the NY107-
ins-205 mutant after only two passages of the rescued virus in
eggs, also indicates the importance of these positions for HA
functionality/glycan specificity. Analysis of the structural data
reveals that positions 186 and 205 are on opposite sides of a
monomer but are both close to the 220-loop deletion region in
the trimeric form. The Glu at position 186 is close to Arg220 and
may interact with Arg220 when binding avian receptors. Position
205 in the neighboring monomer may be important in trimer
stability and maintaining RBS functionality. If one models the pre-
1996 220-loop restored into the NY107 structure, Arg205, Glu186
and the loop all clash, thus explaining the Glu186Gly mutation
that emerged in the NY107-ins-205 virus HA after limited egg
passage.
The NY107 RBS with its more restricted a2-3 glycan binding
preference and weak/moderate increase in a2-6 binding may have
given the virus a selective advantage to be maintained in poultry at
live bird markets and supplying farms. Certain terrestrial birds,
such as quails and chickens, have recently been shown to present
both human and avian types of receptors in the trachea and
intestine [37,38,39]. Although it is not known what specific glycans
are presented in these animals, it is conceivable that a virus with
mixed specificity might have a distinct advantage over avian
viruses that have specific avian receptor requirements, particularly
in bird markets where multiple species coalesce. Previous results
with H7N2, H9N2 and H5N1 viruses all highlight the fact that an
increase in a2-6-binding preference is not sufficient for efficient
transmission of avian influenza viruses to humans [22,40,41].
Although it remains to be seen whether prolonged circulation of
viruses in terrestrial birds, such as domestic chickens, can provide a
possible route for viruses to adapt for efficient human infection
[11], continued surveillance of influenza viruses from avian and
other animal reservoirs is urgently needed to define their zoonotic
potential.
Materials and Methods
Cloning
Based on H3 numbering [42], cDNA corresponding to residues
11–329 (HA1) and 1–176 (HA2) of the ectodomain of the
hemagglutinin
(HA)
from
A/New
York/107/2003
(H7N2;
Genbank:ACC55270)
and
A/Netherlands/219/2003
(H7N7;
Genebank: AAR02640) was cloned into the baculovirus transfer
vector, pAcGP67-A (BD Biosciences), incorporating a C-terminal
thrombin cleavage site, a ‘‘foldon’’ sequence [43] and a His-tag at
the extreme C-terminus of the construct to enable protein
purification [25,44]. Transfection and virus amplification were
carried out according to the baculovirus expression system manual
(BD Biosciences Pharmingen).
Figure 4. Glycan interactions within the NY107 RBS. The top panel shows the interactions of NY107 with (A) 39SLN, (B) 69SLN and (C) LSTb.
NY107 is shown in orange/red/magenta cartoon respectively. The interacting HA residues are shown as green sticks. The bottom panel shows the
electron density map of the ligands. The NY107 is shown in the same colors as above, and the ligands are shown as green sticks, the 2fo-fc electron
density maps (contoured at 1s) are shown in grey. Simulated annealing omit maps are shown in supplementary Figure S4.
doi:10.1371/journal.ppat.1001081.g004
Structure of a North American H7 Hemagglutinin
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Protein expression and purification
Soluble NY107 was recovered from the cell supernatant by metal
affinity chromatography using Ni-NTA resin (Qiagen Inc.). Frac-
tions containing NY107 were pooled and dialyzed against 10 mM
Tris-HCl, 50 mM NaCl, pH 8.0, then subjected to ion-exchange
chromatography (IEX) using a Mono-Q HR 10/10 column (GE
Healthcare). IEX purified NY107 was subjected to thrombin digest
(3 units/mg protein; overnight at 4uC) and purified by gel filtra-
tion chromatography using a Superdex-200 16/60 column (GE
Healthcare) and 50 mM Tris-HCl, 100 mM NaCl, pH 8.0 as
running buffer. Protein eluting as a trimer was buffer exchanged
into 10 mM Tris-HCl, 50 mM NaCl, pH 8.0 and concentrated to
14.5 mg/ml for crystallization trials. At this stage, the protein
sample still contained the additional plasmid-encoded residues at
both the N (ADPG) and C terminus (SGRLVPR).
Crystallization, ligand soaking and data collection
Initial crystallization trials were set up using a Topaz Free
Interface Diffusion (FID) Crystallizer system (Fluidigm Corpora-
tion, San Francisco, CA). Crystals were observed in several
conditions containing PEG 3350 or PEG 4000. Following opti-
mization, diffraction quality crystals for NY107 were obtained at
room temperature using a modified method for microbath under
oil [45], by mixing the protein with reservoir solution containing
20% PEG 3350, 0.2 M magnesium chloride at pH 7.2. For
receptor analog complexes, crystals were soaked for 3 hours in the
crystallization buffer containing 10 mM 39SLN or 69SLN (V-labs
Inc., Covington, LA), or overnight in 10mM LSTb (Sigma, St.
Louis, MO). All crystals were flash-cooled at 100K using 20%
glycerol as the cryo-protectant. Datasets were collected at
Advanced Photon Source (APS) beamlines 22 ID and BM at
100K. Data were processed with the DENZO-SACLEPACK suite
[46]. Statistics for data collection are presented in Table 1.
Structure determination and refinement
The
structure
of
NY107
was
determined
by
molecular
replacement with Phaser [47] using the structure of the avian
H7 (Av-H7) from A/turkey/Italy/2002, pdb:1TI8 (HA1, 78%
identity; HA2, 90% identity) as the searching model. One HA
trimer occupies the asymmetric unit with an estimated solvent
content of 58% based on a Matthews’ coefficient (Vm) of 2.9 A˚ 3/
Da. Rigid body refinement of the trimer led to an overall R/Rfree
of 28.6%/37.4%. The model was then ‘‘mutated’’ to the correct
sequence and rebuilt by Coot [48], then the protein structures
were refined with REFMAC [49] using TLS refinement [50]. The
final models were assessed using MolProbity [51]. The three
complex structures were refined and evaluated using the same
strategy. All statistics for data processing and refinement are
presented in Table 1. Electron density maps (2fo-fc) were
generated in Refmac [49] while simulated annealing omit maps
were generated by sa-omit-map, a part of the Crystallography and
NMR System (CNS) software [52].
Virus generation
Wild type and mutant viruses of NY107 (H7N2) and A/
Netherland/219/2003 (H7N7) were generated from plasmids by a
reverse genetics approach [53]. To generate viruses with amino
acid insertion or substitution in the HA, mutations were
introduced into plasmid DNA with an overlap extension PCR
approach [54]. Viruses derived by plasmid transfection of HK293
cells were propagated in eggs. The genomes of resulting virus
stocks were sequenced to detect the emergence of possible variants
during amplification.
Glycan binding analyses
Glycan microarray printing and recHA analyses have been
described previously [24,30,44,55] (see Table 2 for glycans used
for analyses in these experiments). Virus were analyzed on the
microarray as described previously [30].
PDB accession codes
The atomic coordinates and structure factors of NY107 are
available from the RCSB PDB under accession codes 3M5G for
the unliganded NY107, 3M5H for the NY107 with 39-SLN and
3M5I and 3M5J for NY107 with 69SLN and LSTb, respectively.
Accession/ID numbers for genes/proteins used in this
work
A/New York/107/03 (H7N2), Genbank: ACC55270; A/
Netherlands/219/03 (H7N7), Genbank: AAR02640; A/Hong
Kong/1-9/68 (H3N2), 2HMG; A/Duck/Ukraine/1/63 (H3N8),
PDB: 1MQL; A/South Carolina/1/18 (H1N1), PDB: 1RD8; A/
Puerto Rico/8/34 (H1N1), PDB: 1RU7; A/Swine/Iowa/15/30
(H1N1), PDB: 1RUY; A/Singapore/1/1957
(H2N2), PDB:
2WRC; A/Viet Nam/1203/04 (H5N1), PDB: 2FK0; A/Duck/
Singapore/3/97 (H5N3), PDB: 1JSM; A/Swine/Hong Kong/9/
98 (H9N2), PDB: 1JSD; A/Turkey/Italy/8000/02 (H7N3), PDB:
1TI8; C/Johannesburg/1/66, 1FLC.
Supporting Information
Figure S1
Sequence alignment of selected structurally available
HAs. Human H3 (PDB: 2HMG), Avian H3 (PDB: 1MQL), 1918-
Human H1 (PDB: 1RD8), 1934-Human H1 (PDB: 1RU7), Swine
H1 (PDB: 1RUY), 1957-Huamn H2 (PDB: 2WRC), Human H5
(PDB: 2FK0), Avian H5 (PDB: 1JSM), Swine H9 (PDB: 1JSD),
and Avian H7 (PDB: 1TI8) were used in the alignments. The
fusion domain of HA1 is highlighted in magenta, the vestigial
esterase domain is highlighted in green, the receptor binding
domain is highlighted in blue, and the fusion domain of HA2 is
highlighted in red. Residue numbering is based on the H3 HA
sequence.
Found at: doi:10.1371/journal.ppat.1001081.s001 (2.84 MB TIF)
Figure S2
Expression and purification of NY107. SDS-PAGE
reveals that NY107 was expressed as the HA0 form with a mass
approximately 60kDa (middle lane). Thrombin cleavage resulted in
an unexpected reduction in band size to a HA1/HA2 profile (right
lane) with possible multiple glycoforms for the HA2 clearly present.
Found at: doi:10.1371/journal.ppat.1001081.s002 (0.23 MB TIF)
Figure S3
Comparison of glycan binding to NY107 with other
HAs. A. Overlap of a2-3 ligands binding in the receptor binding
site from NY-H7 (green), Av-H3 (orange), 1930-Hu-H1 (magenta),
1957-Hu-H2 (cyan), Av-H5 (yellow), and Sw-H9 (grey). B.
Overlap of a2-6 linkage ligands binding in the receptor binding
site from NY-H7 (green), Av-H3 (orange), 1930-Hu-H1 (magenta),
1957-Hu-H2 (cyan), Av-H5 (yellow), and Sw-H9 (grey).
Found at: doi:10.1371/journal.ppat.1001081.s003 (2.55 MB TIF)
Figure S4
Simulated annealing omit maps of the receptor
binding site (contoured at 1s). A. NY107 (blue), B. NY107-39SLN
(orange), C. NY107-69SLN (red), and D. NY107-LSTb (magenta).
The protein model is shown in cartoon, and the residues involved
in the binding to receptor analogs were shown in sticks. Maps were
generated using version 1.2 of the Crystallography and NMR
System (CNS) software.
Found
at:
doi:10.1371/journal.ppat.1001081.s004
(1.93
MB
TIF)
Structure of a North American H7 Hemagglutinin
PLoS Pathogens | www.plospathogens.org
9
September 2010 | Volume 6 | Issue 9 | e1001081
Figure S5
Comparison of NY107 RBS to HEF. Overlap of RBS
from NY107 (green), Av-H7 (marine) and HEF (magenta).
Found at: doi:10.1371/journal.ppat.1001081.s005 (1.12 MB TIF)
Table S1
Comparison of r.m.s.d. (A˚ ) for different HA domains.
For analyzing differences in the overall structure, r.m.s.d. values
were calculated between monomers or domains of different HA’s,
after the Ca atoms of the HA2 domains were superposed by
sequence and structural alignment onto the equivalent domains of
NY107.
Found
at:
doi:10.1371/journal.ppat.1001081.s006
(0.04
MB
DOC)
Table S2
Comparison of r.m.s.d. (A˚ ) for individual domains.
Each domain was superimposed separately to determine how the
individual NY107 domains compared to equivalent domains in the
other structures.
Found
at:
doi:10.1371/journal.ppat.1001081.s007
(0.04
MB
DOC)
Table S3
Molecular interactions between NY107 and receptor
analogs. The hydrogen bond cutoff is 3.8 A˚
for the listing
interactions.
Found
at:
doi:10.1371/journal.ppat.1001081.s008
(0.07
MB
DOC)
Table S4
Glycan array differences between NY107, the fully
restored NY107-ins, and NL219 (virus and rHA). The color
coding in the left hand column reflects the same coloring scheme
used in Figures 2 and 3. Significant binding of samples to glycans
are qualitatively estimated based on relative strength of the signal
for the data shown in Figures 2 and 3 Strong (+++), weak (+).
Found
at:
doi:10.1371/journal.ppat.1001081.s009
(0.19
MB
DOC)
Acknowledgments
The authors would like to thank the staff of SER-CAT sector 22 for their
help in data collection. We also thank WHO Global Influenza Surveillance
Network for providing NY107 and NL219 viruses from which the reverse
genetics viruses were generated. Glycan microarray data presented here
will be made available on-line through the CFG web site upon publication
(www.functionalglycomics.org). The findings and conclusions in this report
are those of the authors and do not necessarily represent the views of the
Centers for Disease Control and Prevention or the Agency for Toxic
Substances and Disease Registry.
Author Contributions
Conceived and designed the experiments: HY LMC PJC ROD JS.
Performed the experiments: HY LMC PJC JS. Analyzed the data: HY
LMC PJC ROD JS. Wrote the paper: HY LMC PJC ROD JS.
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Structure of a North American H7 Hemagglutinin
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September 2010 | Volume 6 | Issue 9 | e1001081
|
3M5I
|
Crystal structure of a H7 influenza virus hemagglutinin complexed with 6SLN
|
Structures of Receptor Complexes of a North American
H7N2 Influenza Hemagglutinin with a Loop Deletion in
the Receptor Binding Site
Hua Yang, Li-Mei Chen, Paul J. Carney, Ruben O. Donis, James Stevens*
Influenza Division, Centers for Disease Control and Prevention, Atlanta, Georgia, United States of America
Abstract
Human infections with subtype H7 avian influenza viruses have been reported as early as 1979. In 1996, a genetically stable
24-nucleotide deletion emerged in North American H7 influenza virus hemagglutinins, resulting in an eight amino acid
deletion in the receptor-binding site. The continuous circulation of these viruses in live bird markets, as well as its
documented ability to infect humans, raises the question of how these viruses achieve structural stability and functionality.
Here we report a detailed molecular analysis of the receptor binding site of the North American lineage subtype H7N2 virus
A/New York/107/2003 (NY107), including complexes with an avian receptor analog (39-sialyl-N-acetyllactosamine, 39SLN)
and two human receptor analogs (69-sialyl-N-acetyllactosamine, 69SLN; sialyllacto-N-tetraose b, LSTb). Structural results
suggest a novel mechanism by which residues Arg220 and Arg229 (H3 numbering) are used to compensate for the deletion
of the 220-loop and form interactions with the receptor analogs. Glycan microarray results reveal that NY107 maintains an
avian-type (a2-3) receptor binding profile, with only moderate binding to human-type (a2-6) receptor. Thus despite its
dramatically altered receptor binding site, this HA maintains functionality and confirms a need for continued influenza virus
surveillance of avian and other animal reservoirs to define their zoonotic potential.
Citation: Yang H, Chen L-M, Carney PJ, Donis RO, Stevens J (2010) Structures of Receptor Complexes of a North American H7N2 Influenza Hemagglutinin with a
Loop Deletion in the Receptor Binding Site. PLoS Pathog 6(9): e1001081. doi:10.1371/journal.ppat.1001081
Editor: Fe´lix A. Rey, Institut Pasteur, France
Received March 16, 2010; Accepted July 28, 2010; Published September 2, 2010
This is an open-access article distributed under the terms of the Creative Commons Public Domain declaration which stipulates that, once placed in the public
domain, this work may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose.
Funding: This work was funded by the Centers for Disease Control and Prevention. Use of the Advanced Photon Source at Argonne National Laboratory was
supported by the U. S. Department of Energy, Office of Science, Office of Basic Energy Sciences, under Contract No. DE-AC02-06CH11357. Glycan microarrays as
well as glycans for direct binding experiments were produced for the Centers for Disease Control by the CFG funded by National Institute of General Medical
Sciences Grant GM62116. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: fwb4@cdc.gov
Introduction
Influenza is an acute respiratory virus that infects up to 20% of
the population in the United States, resulting in ,36,000 deaths
annually [1,2]. The two membrane glycoproteins on the surface of
influenza A virus, hemagglutinin (HA), which functions as the
receptor binding and membrane fusion glycoprotein in cell entry,
and neuraminidase (NA), which functions as the receptor destroying
enzyme in virus release, form the basis for defining subtypes [3]. To
date, 16 HA (H1–H16) and 9 NA (N1–N9) have been identified in
avian species [4], while in the last century, only three subtypes,
H1N1 in 1918 and 2009, H2N2 in 1957, and H3N2 in 1968 [5,6,7],
have successfully adapted to humans. Hemagglutinin binds to sialic
acid (SA) glycans present on host cell surfaces. The receptors on
epithelial cells of the human upper respiratory tract are mainly
a2-6-linked SA moieties [8]. Since avian influenza viruses pre-
dominately bind a2-3-linked SA, and human influenza viruses
preferentially bind to a2-6-linked SA, human infection by avian
influenza viruses is rare [9]. However, since 1997 a growing number
of human cases of avian influenza infection have been reported [10],
including H5N1, H7N2, H7N3, H7N7, and H9N2 strains [11].
Although the current situation with the pandemic H1N1 influenza
virus dominates public health efforts, the prospect of a novel
pandemic emerging from these isolated cases continues to be a
major public health threat around the world.
Early cases of human infection by H7 influenza viruses are reported
as far back as 1979 [12,13]. Since 2002, multiple outbreaks and
human infections of H7 subtype viruses; within both Eurasian and
North American lineages have been reported. In the Netherlands in
2003, a highly pathogenic avian influenza (HPAI) H7N7 outbreak
resulted in more than 80 cases of human infections, including one
fatality [14,15]. In New York in 2003, a single case of human
respiratory infection of H7N2 was reported [16] and in British
Columbia in 2004, an H7N3 virus caused two cases of conjunctivitis
[17,18]. More recently in 2007, the United Kingdom reported several
cases of low pathogenic avian influenza (LPAI) H7N2 virus infections
that caused influenza-like illness and conjunctivitis [19].
Since 1996, H7 viruses of the North American lineage have
been circulating in regional live bird markets [20], containing a
24-nucleotide deletion resulting in an eight amino acid deletion in
the receptor-binding site (RBS) of HA (Figure S1). The recent
human infections with H7 in North America have raised public
health concerns as to how these viruses adapt to such a dramatic
structural change while remaining one of the predominant
circulating viral strains. A recent study of H7 viruses isolated
from previous outbreaks revealed efficient replication in both
mouse and ferret animal models [21]. In particular, ferret studies
with A/New York/107/2003 (NY107), an H7N2 virus isolated
from a man in New York, not only showed efficient replication in
the upper respiratory tract of the ferret but also the capacity for
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intra-species transmission by direct contact [21,22]. Interestingly,
both an increased preference for a2-6 and decreased preference for
a2-3-linked sialosides of this virus compared to the other avian
influenza viruses was shown by previous glycan microarray analysis
but less so by a competitive solid-phase binding assay [22,23].
Here we report a detailed molecular analysis of the RBS of the
HA from North American lineage H7N2 virus, NY107, including
glycan microarray analyses and structural analyses of the HA in
complex with an avian receptor analog (39-Sialyl-N-acetyllactosa-
mine, 39SLN) and two human receptor analogs (69-Sialyl-N-
acetyllactosamine, 69SLN; Sialyllacto-N-tetraose b, LSTb). These
results provide important insight into the interaction of H7 HAs
with both avian and human hosts.
Results
Overall structure
By using x-ray crystallography, the structure of H7 HA from the
NY107 virus was determined to 2.6 A˚ resolution (Table 1). In
addition, we also report three H7 HA receptor complex structures,
with avian receptor analog (39SLN) to 2.7 A˚ resolution and with
human receptor analogs (69SLN and LSTb) to 3.0 A˚ and 2.6 A˚
resolution, respectively (Table 1). The overall structure of NY107
is similar to other reported HA structures with a globular head
containing the RBS and vestigial esterase domain, and a
membrane proximal domain with its distinctive, central helical
stalk and HA1/HA2 cleavage site (Figure 1A). Although five
asparagine-linked glycosylation sites are predicted in the NY107
HA monomer, interpretable electron density was observed at only
two sites, Asn38 in HA1 and Asn82 in HA2 (all residue numbers
are based on H3 numbering). At these sites, only one or two N-
acetyl glucosamines could be interpreted.
During viral replication, HA is synthesized as a single chain
precursor (HA0) and cleaved by specific host proteases into the
infectious HA1/HA2 form. In baculovirus expression systems,
highly pathogenic HAs, with a polybasic cleavage site, are
expressed as an HA1/HA2 form [24], whereas HAs with
monobasic cleavage sites (single Arg) from low pathogenic viruses
are expressed as the HA0 form [25]. NY107 is regarded as a low
pathogenic virus, and as expected, was produced in the HA0
form (Figure S2). However, subsequent digestion with thrombin
protease to remove the His-tag resulted in cleavage to a profile on
SDS-PAGE comparable to that of an HA1/HA2 form (Figure S2).
A comparison of the NY107 cleavage site with the consensus
cleavage pattern in the MEROPS database (http://merops.
sanger.ac.uk) suggests it to be a possible thrombin cleavage site.
Based on their molecular phylogenies, HAs are divided into two
groups and five clades: group 1 includes H8, H9, and H12; H1,
H2, H5, and H6; H11, H13 and H16; group 2 includes H3, H4,
and H14; H7, H10 and H15 [26]. Among all available HA
structures, we selected ten representative HAs from both avian and
human subtypes for structural analysis. As expected, NY107 HA is
structurally very similar to the Avian-H7 in all comparisons and
closely related to H3, the other group 2 members used in the
analyses (Tables S1 and S2).
The receptor binding site
The RBS is at the membrane distal end of each HA monomer
and its specificity for sialic acid and the nature of its linkage to a
vicinal galactose residue is a major determinant of host range-
restriction. The consensus RBS for all current HAs is composed of
three major structural elements: a 190-helix (residues 188–194), a
220-loop (residues 221–228), and a 130-loop (residues 134–138).
In addition, highly conserved residues (Tyr98, Trp153, His183,
and Tyr195) form the base of the pocket.
Although the NY107 RBS is similar to other subtypes (H1, H2,
H3, H5, and H9), a previously observed specific feature of H7 HAs,
is also observed in the NY107 150-loop region: two residues inserted
at position 158 result in this loop protruding more than 6A˚ towards
the binding site compared to other subtype HAs (Figure 1B and
Table S2) [27]. More interestingly, the eight amino acid deletion,
only found in the North American lineage H7s, from position 221 to
228 (Figure S1), resulted in a complete loss of the 220-loop
(Figure 1B). Sequence alignment shows that Arg220 and Arg229 are
conserved in all influenza A HA subtypes (Figure S1), but structural
alignment of NY107 HA shows Arg220 occupying the Gly228
position, and the much shorter loop turns at residue Pro217
(Figure 1C). The Ca distance between NY107 Arg220 and its
homolog in the Av-H7 structure (PDB: 1TI8) [27] is 5.8A˚ , and they
point in opposite directions (Figure 1C). The side chain direction of
Av-H7 Arg220 is almost parallel with the beta sheet after Arg229,
whereas the NY107 Arg220 points downward to the binding pocket.
The Ca position of Arg229 in both H7 structures remains the same,
except the side chain in the NY107 swings away by about 5.9A˚
(Figure 1C) and could help to stabilize this region by forming a
hydrogen bond to the mainchain carbonyl of Gln210 in the
neighboring monomer. In the absence of the 220-loop in NY107
HA, upon glycan binding the long side chain of Arg220 compensates
for its loss and is displaced 4A˚ upward to form hydrogen bonds with
receptor analogs inside the binding pocket (Figure 1D).
Effect of loop truncation on the receptor binding
specificity of NY107
Previously, mutations in the HA receptor binding domains of
H1N1 (Glu190Asp/Gly225Asp) and H2N2/H3N2 (Gln226Leu
and Gly228Ser) subtypes were responsible for adaptation of these
viruses to pandemic strains [24,28,29,30]. Due to missing residues
221–228 in the NY107 HA RBS, neither mechanism for
adaptation is possible. Thus, in order to look more closely at the
role of the missing loop and its effect on receptor specificity, we
first subjected the recombinant HA (recHA) to glycan microarray
analyses and compared it to a reverse genetics-derived NY107
virus, and a co-circulating Eurasian virus and recHA, A/
Author Summary
Influenza virus adaptation to different hosts usually results
in a switch in receptor specificity of the viral surface coat
protein, hemagglutinin. Indeed, the hemagglutinin sub-
types from the last two human influenza pandemics of the
20th Century (H2 in 1957 and H3 1968) both adapted
successfully to human-type receptor specificity through
only two amino acid mutations in the receptor binding
pocket (Glutamine226RLeucine and Glycine228RSerine).
The recent human infections reported with other avian
subtypes such as H5, H7 and H9 have raised public health
concerns and focused efforts on identifying potential
subtypes from which a future pandemic strain may
emerge. Since 1996, H7 viruses of the North American
lineage have been circulating in regional live bird markets,
containing an eight amino acid deletion in the receptor-
binding site of HA. Here we report a detailed structural
analysis of the receptor binding site of a hemagglutinin
from the North American lineage of H7N2 viruses, in
complex with avian and human receptor analogs, to
understand how these viruses have adapted to such a
dramatic structural change in the binding site while
remaining one of the predominant circulating viral strains.
Structure of a North American H7 Hemagglutinin
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Netherlands/219/2003 (NL219), that has the consensus avian
sequence in the 220-loop and it also infected a human [15].
Glycan microarray analysis of recombinant NY107 (Figure 2A
and Table 2) revealed a highly restricted binding profile with
strong binding to only a2-3 sulfated (#4–8), a2-3 branched (#9–
11) and mixed a2-3/a2-6 branched sialosides (#60–64) as well as
to the long linear sialyl di- and tri-lactosamines (#22, 24). Weak
binding was also observed (above background) to other a2-3
glycans on the array. The recombinant NY107 also revealed a
strict glycan binding preference to only one a2-6 glycan, the
internal structure, Galb1-3(Neu5Aca2-6)GlcNAcb1-3Galb1-4Glc
(#58; LSTb) (Figure 2A), a glycan highlighted in a previous study
[22]. The virus with higher valency and avidity revealed stronger
binding to all a2-3 groups, in addition to the branched di-sialyl a2-
6 biantennary structures (#46–48) as well the LSTb (#58)
(Figure 2B and Table 2). In contrast, the NL219 recHA (Figure 2C
and Table 2) bound well to only the avian a2-3 containing sialyl-
glycans (sulfated, branched, linear and fucosylated). Its corre-
sponding virus also reflected this specificity although it also
revealed strong binding to a2-3 N-glycolylneuraminic acid
(Neu5Gc) containing glycans (#66–70) (Figure 2D and Table 2).
To further assess the effect of the missing 220 loop on HA
structural stability and receptor specificity it was essential to
evaluate these functions on the ancestral HA containing the full
length 220-loop. To this end, we engineered an HA with an avian
H7 consensus (PQVNGQSG) 220-loop re-introduced (NY107-
220ins) into the NY107 HA and recovered this virus by reverse
genetics. Compared to the NY107 virus (Figure 2A) glycan
microarray analyses of the resulting NY107-220ins virus (Figure 3A
and Table 2) revealed a decrease in binding to branched (#9–11)
and linear (#12–27) a2-3 sialosides and a loss of binding to the
branched di-sialyl a2-6 biantennary structures (#46–48), LSTb
(#58) as well as the mixed a2-3/a2-6 branched sialosides (#60–
64). In addition, sequence analysis of the NY107-220ins HA
revealed the presence of quasispecies in the second position of the
inserted loop, P(Q/K)VNGQSG, suggesting that re-introduction
of the loop alone is not tolerated and does not create an avian-type
binding profile. Thus other amino acid substitutions in the HA
might have co-evolved with the deletion of the 220 loop to help
stabilize the RBS/HA to maintain functionality.
When viruses containing this 220-loop deletion emerged in North
America in the mid 90’s, four additional amino acid substitutions,
Gly114Arg, Asp119Gly, Gly186Glu and Gly205Arg, in the HA1 as
well as an Asp19Asn in the HA2 chain were also introduced to most
of the circulating isolates. Of these, Gly186Glu and Gly205Arg in
the HA1 are close to the RBS, at the monomer interface, and could
potentially modulate its structure and/or function. NY107 viruses
with a restored consensus 220-loop and a single Glu186Gly (NY107-
ins-186) or Arg205Gly (NY107-ins-205) substitution as well as the
Glu186Gly/Arg205Gly double substitution (NY107-ins-186/205)
were derived by reverse genetics and evaluated. Glycan microarray
analysis for the three resulting viruses revealed similar glycan
binding profiles with increased binding to a2-3 sialosides, including
mixed a2-3/a2-6 branched sialosides (#60–64), a2-3 Neu5Gc
(#66–70), but limited binding to the a2,6 sialosides (Figures 3B, 3C,
3D), resulting in a binding profile virtually identical to that of the
NL219 virus and other avian influenza viruses (Figure 2D) [30].
Sequence analysis of the three reverse genetics derived viruses
revealed no mutations/quasispecies in the HAs of either the NY107-
ins-186 or the NY107-ins-186/205 virus stocks, indicative of
replication fitness. For the NY107-ins-205 virus however, a
Glu186Gly substitution emerged in the HA after only two passages
in eggs following recovery from DNA transfection, indicating the
importance of the co-variant position 186 with respect to HA
functionality/glycan specificity. Altogether, the data indicates that
the H7 subtype avian influenza viruses that were circulating in
Table 1. Data collection and refinement statistics.
NY107
NY107+39SLN
NY107+69SLN
NY107+LSTb
Data collection
Space group
P212121
P212121
P212121
P212121
Cell dimensions (A˚)
66.96, 115.92, 251.61
67.80, 116.70, 249.84
66.60, 116.58, 250.68
67.08, 116.52, 251.95
Resolution (A˚)
50-2.6 (2.69-2.60)a
30-2.7 (2.80-2.70)
50-3.0 (3.11-3.0)
50-2.6 (2.69-2.60)
Rsym or Rmerge
10.6 (41.3)
14.6 (48.6)
14.3 (35.4)
12.2 (31.5)
I/s
39.6 (2.0)
24.3 (1.7)
34.2 (8.2)
40.5 (9.9)
Completeness (%)
99.2 (99.0)
99.3 (94.6)
92.3 (75.6)
91.3 (86.2)
Redundancy
7.2 (6.2)
5.8 (5.5)
4.9 (4.4)
10.9 (11.2)
Refinement
Resolution (A˚)
50-2.6 (2.67-2.60)
30-2.7 (2.77-2.70)
50-3.0 (3.08-3.00)
50-2.6 (2.67-2.60)
No. of reflections (total)
57285
51770
33421
53603
No. of reflections (test)
3053
2769
1779
2842
Rwork/Rfree
21.7/25.6
21.4/26.4
20.5/26.0
20.4/24.7
No. of atoms
11795
11878
11648
12108
r.m.s.d.- bond length (A˚)
0.006
0.006
0.008
0.006
r.m.s.d.- bond angle (u)
0.905
0.974
1.085
0.859
MolProbityb scores
Favored (%)
96.9
96.5
94.3
97.1
Outliers (%) (No. of residues)
0.1 (1/1434)
0.0 (0/1429)
0.1 (2/1433)
0.1 (2/1435)
aNumbers in parentheses refer to the highest resolution shell.
bReference [51].
doi:10.1371/journal.ppat.1001081.t001
Structure of a North American H7 Hemagglutinin
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aquatic birds and poultry in North America before 1996 exhibited a
classic avian a2-3 sialoside binding preference. In order for the 220-
loop deletion to be tolerated, concurrent Gly186Glu and Gly205Arg
substitutions in the vicinity of RBS of HA emerged to achieve a
restricted a2-3 binding profile and only a moderate/limited increase
in binding to branched di-sialyl a2-6 biantennary structures (#46–
48) as well the a2,6 internal sialoside, LSTb (#58).
NY107 avian receptor complex
To understand from a structural perspective how NY107
interacts with host receptors, we solved the structure of NY107
in complex with an avian and two human receptor analogs.
For the avian receptor analog, 39SLN, the electron density
maps revealed well-ordered features for the Sia-1, Gal-2, and
GlcNAc-3 in the NY107 HA complex structure (Figure 4A).
Structural comparison of NY107 HA binding to other, H1, H2,
H3, H5, and H9 subtypes (Figure S2A) revealed that 39SLN
binding to NY107 resembled binding of the other published
HAs. Indeed, the terminal Sia-1 moiety is positioned almost
identically in all structures, and forms the majority of hydrogen
bonds and contacts with residues in the RBS (Figure 4A and
Table S3).
Figure 1. NY107 HA monomer and comparison of its RBS to other HA structures. (A) One monomer is shown with the HA1 chain colored in
green and the HA2 chain in cyan. The location of the receptor binding site and the HA1/HA2 cleavage site are circled. (B) The superposition of
receptor binding domains of NY107 (green), Av-H7 (marine), 1918-Hu-H1 (magenta), Hu-H5 (yellow), Hu-H3 (orange), and Sw-H9 (grey). The proximity
of Arg220 and Gln226 are highlighted. Three structural elements comprising this binding site are labeled. The two major differences are the extended
150-loop and the deletion of 220-loop of NY107. (C) Overlap of NY107 (green) and Av-H7 (marine) (PDB: 1TI8) illustrates the compensatory effect of
R220 bringing it close to the position occupied by G228 in the avian HA. (D) Overlap of the NY107 (green), NY107- 39SLN (orange), NY107-69SLN (red),
and NY107-LSTb (magenta) structures. All the figures were generated and rendered with the use of MacPyMOL [56].
doi:10.1371/journal.ppat.1001081.g001
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Published avian HA structures with an intact 220-loop form
very close interactions with Gal-2 of 39SLN via residue Gln226
which is important in receptor specificity and host adaptation. For
example, in the avian H7/39SLN HA structure it interacts with
Gal-2 O4 [31]. In the NY107 HA structure, although Gln226 is
absent and no other residue occupies the same space as Gln226
(Figure 1B), Arg220 does forms a hydrogen bond between Arg220
NH2 and Gal-2 O4 (Figure 4A). Interestingly, although there was
interpretable density for the GlcNAc-3 (Figure 4A and Figure
S4B), no hydrogen bonding was apparent between the HA and the
GlcNAc-3, which is consistent with other reported structures [32].
Thus, for binding to avian receptors, the trans conformation of
a2-3 linkages is essential and perhaps only the first two saccharides
are required. Indeed, due to the absence of 220-loop in the NY107
HA structure, the ‘‘aperture’’ of the RBS formed by 220-loop and
130-loop in regular HAs is increased by ,10 A˚ , so that the
branched, internal, and perhaps more complicated glycans might
be accommodated more efficiently.
NY107 human receptor complexes
In the NY107/69SLN complex, only Sia-1 and Gal-2 are
ordered (Figure 4B). The Sia-1 remains in the same position as
previously analyzed glycan/HA complexes from H1, H2, H3, H5,
and H9 (Figure S3B), whereas the Av-H7 complex structure with
Sialyllacto-N-tetraose c (LSTc) did not reveal any density for the
Sia-1 in the receptor binding site [31]. The Gal-2 position varies
significantly among different subtypes. Compared to the human-
adapted H1 HA [32], Gal-2 in the NY107 HA is 3A˚ higher, and
thus is further from the protein (Figure S3B). In NY107, the Gal-2
only forms an intramolecular, saccharide-saccharide interaction
with Sia-1. The poor electron density map and fewer interactions
with protein residues suggest that the cis conformation of a2-6
Figure 2. Receptor specificity of NY107 recHA and virus. Glycan microarray analysis of recombinant NY107 HA (A) and NY107 virus (B)
compared to the recHA (C) and virus (D) from a Eurasian lineage A/Netherlands/219/2003 H7 influenza virus that was circulating in the same year and
also infected a human. Colored bars highlight glycans that contain a2-3 SA (blue) and a2-6 SA (red), a2-6/a2-3 mixed SA (purple), N-glycolyl SA
(green), a2-8 SA (brown), b2-6 and 9-O-acetyl SA, and non-SA (grey). Error bars reflect the standard error in the signal for six independent replicates
on the array. Structures of each of the numbered glycans are found in Table S4.
doi:10.1371/journal.ppat.1001081.g002
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linkages in 69SLN trisaccharides show a reduced binding affinity
with NY107.
Glycan array results with NY107 revealed a strong binding
signal for the internal a2-6 sialoside, LSTb. To further investigate
this interaction, we solved the structure of the NY107/LSTb
complex. The final model contained Sia-1, NAG-2, Gal-3, and
Gal-5 in the RBS. Although glycan microarray data indicated
NY107 to have a specific affinity for LSTb, few interactions were
apparent from the crystal structure. Sia-1 still forms multiple
hydrogen bonds with residues in the RBS (Table S3 & Figure 4C).
The branched Gal-5 interacts with Ser137, to help stabilize the
LSTb binding. However, Arg220 and Lys193, the two residues
showing close binding with 39SLN, did not form any hydrogen
bonds with LSTb. In the structure, Gal-5 also interacts with a
crystal packing symmetry mate and thus the flexibility of whole
LSTb may be restricted. In solution, with more freedom, the
LSTb should be able to tilt closer to the RBS, and thus Glc-4 may
have more interactions with the 190-helix than seen in the crystal
structure.
Discussion
Human infections by avian influenza viruses, including H7
subtypes, continue to pose a major public health threat. Although
the
species
barrier
prevents
avian
influenza
viruses
from
widespread infection of the human population, the molecular
determinants of efficient interspecies transmission and pathoge-
nicity are still poorly understood. The viral coat protein HA
however, is perhaps a critical molecule since previous pandemic
viruses modified their receptor specificity and overcame the
interspecies barrier to spread in the human population. Although
HA structures alone and in complex with receptor analogs provide
considerable insight into receptor binding, it is clear that HAs from
different species and subtypes have significant structural variation.
Indeed, low-pathogenic H7N2 avian influenza viruses with an 8
amino acid deletion within its RBS started to circulate in live-bird
markets in the northeast United States in 1996. Despite what one
would consider a debilitating mutation, these viruses have been
reported as the predominant isolate [33]. Whether such a deletion
contributed to their evolutionary success and how are an
important questions, especially in light of NY107’s ability to
produce respiratory illness in humans [16], as well as its reported
increased affinity for human-type receptors and ability for contact
transmission in ferrets [21]. To try to help answer these questions,
we have analyzed the molecular structures of NY107 and its
complexes with receptor analogs to explain receptor specificity at
the molecular level.
The crystal structures of NY107 and its complexes with both
avian and human receptor analogs describe a mechanism as to
how an influenza virus might adapt by dramatically altering its
RBS, and still be functional. Arg220 of the HA1 chain of NY107
compensates for the loss of the 220-loop, by forming hydrogen
bonds with Gal-2 from the avian analog (binding was not observed
in either of the structures complexes with the human analogs).
However, in the LSTb complex, branched Gal-5 forms extra
interactions with the 130-loop, thus improving the binding
preference for this particular glycan. Consistent with the structural
evidence, glycan microarray analyses of NY107 revealed a strong
binding preference for the branched a2-6 sialoside, LSTb. Except
for the absence of the 220-loop, other key residues within the RBS
are conserved in NY107 and thus, direct interactions with sialic
acid are maintained.
The 220-loop is recognized as one of the three crucial structural
elements in the RBS. Aside from the North American lineage
H7N2 viruses, which have been circulating with a deletion (221–
228) in this loop, there has been one other report describing a
seven amino acid deletion (224–230) in a laboratory generated
H3N2 escape mutant which was reported to have a slightly
Table 2. Comparison of the sialoside receptor specificity of the HAs from H7 influenza viruses.
Glycan Group
Graph
Numbera
NY107
RecHA
NY107
Virus
NY107-ins
Virus
NY107-ins
E186G Virus
NY107-ins
R205G Virus
NY107-ins E186G/
R205G Virus
NL219
RecHA
NL219
Virus
a2-3
Sulfated
4–8
+++b
+++
+++
+++
+++
+++
+++
+++
Branched
9–11
+++
+++
+
+++
+++
+++
+++
+++
Linear
12–27
+
+++
+
+++
+++
+++
+++
+++
Fucosylated
28–34
2
+++
+++
+++
+++
+++
+++
+++
a2-6
Sulfated
41
2
2
2
2
2
2
2
2
Branched mono-sialyl
42–45, 49
2
2
2
2
2
2
2
2
Branched di-sialyl
46–48
2
+++
2
2
2
2
2
2
Linear
50–56
2
2
2
2
2
2
2
2
Internal
58–59
+++
+++
2
2
2
2
2
2
Other
Sialic acid
1–2
2
+++
+
2
2
2
2
2
a2-3/a2-6 Branched
60–64
2
2
2
+++
+++
+++
+++
+++
Neu5Gcc
65–72
2
2
2
+++
+++
+++
2
+++
aMembers of each group are identified according to the graph number used in the microarray data in Figures 2 and 3 and correspond to numbers in the complete
glycan list (Table S4).
bBinding of samples to glycan subclasses are qualitatively estimated based on relative strength of the signal for the data shown in Figures 2 and 3: strong (+++), weak
(+), absent (2).
cN-glycolylneuraminic acid.
doi:10.1371/journal.ppat.1001081.t002
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increased affinity for a2-3-linked glycans by hemagglutination
assay [34]. Meanwhile, the equivalent region in the hemaggluti-
nin-esterase-fusion (HEF) protein of influenza C virus reveals a
rearrangement resulting in a truncated 260-loop in its RBS (Figure
S5) [35]. However, without structural data with appropriate
receptor analogs, it is not possible to compare the role of these loop
variants in receptor binding to the H7 HA structure described
here.
When compared to NL219, another co-circulating H7 avian
virus HA (Figure 2C and D), overall binding to a2-3-linked
glycans was markedly reduced, while increased binding to a2-6-
linked receptors was only marginal. However, these results focus
attention on only 2 sub-classes of human-type receptors that may
be important for infection (and transmission in ferrets). The
NY107 virus interaction with biantennary glycans (Figure 2B),
although weak (not seen in Figure 2A with recHA), is a possible
route for virus entry as biantennary structures are common on
tissues, i.e. glycan profiling data from human lung tissue on the
Consortium for Functional Glycomics (CFG) web site. In addition,
the internal sialoside, LSTb, was observed in both virus and
recHA microarray data, suggesting this type of glycan has good
affinity for this HA. The significance of this is unknown since
LSTb has only been described in human milk [36].
Interestingly, NY107 and NL219 virus receptor binding and
specificity has been addressed previously using glycan microarray
analysis that reported a significantly increased preference for a2-6
and decreased preference for a2-3-linked sialosides [22]. In
addition, the same viruses were also included in a recent study
from Gambaryan et al. using a competitive solid-phase binding
assay [23]. Our findings confirm and extend the receptor binding
specificity reported by these authors in that they reported both
viruses binding to sulfated sialylglycans with a lactosamine (Galb1-
4GlcNAc core and reported only a moderate binding affinity for
a2-6-sialyllactosamine, the human-type receptor analog used in
their assay.
The 220-loop is an integral feature of the receptor binding site,
and thus one would predict that such a deletion might have
compromised this strain to be deleted from the population of
circulating viruses. However, this was not the case [33] and its
existence appears to be in part due to the additional mutations at
Figure 3. Effect of 220-loop deletion and additional RBS mutations on NY107 receptor specificity. NY107 was engineered to restore the
220-loop to a consensus full-length HA from 1996 (A) and additional co-variant amino acid substitutions, Glu186Gly (B), Arg205Gly (C) and the double
mutant Glu186Gly/Arg205Gly (D) to restore, on the NY107 framework, an HA RBS found in viruses prior to the introduction of the deletion in North
American viruses. Colored bars group glycans as described in Figure 3. Error bars reflect the standard error in the signal for six independent replicates
on the array. Structures of each of the numbered glycans are found in Table S4.
doi:10.1371/journal.ppat.1001081.g003
Structure of a North American H7 Hemagglutinin
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positions 186 and 205. Restoration of the loop with either or both
residues mutated back to the pre-1994 consensus sequence
resulted in a classic avian influenza virus binding profile. The
emergence of the Glu186Gly mutation in the HA of the NY107-
ins-205 mutant after only two passages of the rescued virus in
eggs, also indicates the importance of these positions for HA
functionality/glycan specificity. Analysis of the structural data
reveals that positions 186 and 205 are on opposite sides of a
monomer but are both close to the 220-loop deletion region in
the trimeric form. The Glu at position 186 is close to Arg220 and
may interact with Arg220 when binding avian receptors. Position
205 in the neighboring monomer may be important in trimer
stability and maintaining RBS functionality. If one models the pre-
1996 220-loop restored into the NY107 structure, Arg205, Glu186
and the loop all clash, thus explaining the Glu186Gly mutation
that emerged in the NY107-ins-205 virus HA after limited egg
passage.
The NY107 RBS with its more restricted a2-3 glycan binding
preference and weak/moderate increase in a2-6 binding may have
given the virus a selective advantage to be maintained in poultry at
live bird markets and supplying farms. Certain terrestrial birds,
such as quails and chickens, have recently been shown to present
both human and avian types of receptors in the trachea and
intestine [37,38,39]. Although it is not known what specific glycans
are presented in these animals, it is conceivable that a virus with
mixed specificity might have a distinct advantage over avian
viruses that have specific avian receptor requirements, particularly
in bird markets where multiple species coalesce. Previous results
with H7N2, H9N2 and H5N1 viruses all highlight the fact that an
increase in a2-6-binding preference is not sufficient for efficient
transmission of avian influenza viruses to humans [22,40,41].
Although it remains to be seen whether prolonged circulation of
viruses in terrestrial birds, such as domestic chickens, can provide a
possible route for viruses to adapt for efficient human infection
[11], continued surveillance of influenza viruses from avian and
other animal reservoirs is urgently needed to define their zoonotic
potential.
Materials and Methods
Cloning
Based on H3 numbering [42], cDNA corresponding to residues
11–329 (HA1) and 1–176 (HA2) of the ectodomain of the
hemagglutinin
(HA)
from
A/New
York/107/2003
(H7N2;
Genbank:ACC55270)
and
A/Netherlands/219/2003
(H7N7;
Genebank: AAR02640) was cloned into the baculovirus transfer
vector, pAcGP67-A (BD Biosciences), incorporating a C-terminal
thrombin cleavage site, a ‘‘foldon’’ sequence [43] and a His-tag at
the extreme C-terminus of the construct to enable protein
purification [25,44]. Transfection and virus amplification were
carried out according to the baculovirus expression system manual
(BD Biosciences Pharmingen).
Figure 4. Glycan interactions within the NY107 RBS. The top panel shows the interactions of NY107 with (A) 39SLN, (B) 69SLN and (C) LSTb.
NY107 is shown in orange/red/magenta cartoon respectively. The interacting HA residues are shown as green sticks. The bottom panel shows the
electron density map of the ligands. The NY107 is shown in the same colors as above, and the ligands are shown as green sticks, the 2fo-fc electron
density maps (contoured at 1s) are shown in grey. Simulated annealing omit maps are shown in supplementary Figure S4.
doi:10.1371/journal.ppat.1001081.g004
Structure of a North American H7 Hemagglutinin
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Protein expression and purification
Soluble NY107 was recovered from the cell supernatant by metal
affinity chromatography using Ni-NTA resin (Qiagen Inc.). Frac-
tions containing NY107 were pooled and dialyzed against 10 mM
Tris-HCl, 50 mM NaCl, pH 8.0, then subjected to ion-exchange
chromatography (IEX) using a Mono-Q HR 10/10 column (GE
Healthcare). IEX purified NY107 was subjected to thrombin digest
(3 units/mg protein; overnight at 4uC) and purified by gel filtra-
tion chromatography using a Superdex-200 16/60 column (GE
Healthcare) and 50 mM Tris-HCl, 100 mM NaCl, pH 8.0 as
running buffer. Protein eluting as a trimer was buffer exchanged
into 10 mM Tris-HCl, 50 mM NaCl, pH 8.0 and concentrated to
14.5 mg/ml for crystallization trials. At this stage, the protein
sample still contained the additional plasmid-encoded residues at
both the N (ADPG) and C terminus (SGRLVPR).
Crystallization, ligand soaking and data collection
Initial crystallization trials were set up using a Topaz Free
Interface Diffusion (FID) Crystallizer system (Fluidigm Corpora-
tion, San Francisco, CA). Crystals were observed in several
conditions containing PEG 3350 or PEG 4000. Following opti-
mization, diffraction quality crystals for NY107 were obtained at
room temperature using a modified method for microbath under
oil [45], by mixing the protein with reservoir solution containing
20% PEG 3350, 0.2 M magnesium chloride at pH 7.2. For
receptor analog complexes, crystals were soaked for 3 hours in the
crystallization buffer containing 10 mM 39SLN or 69SLN (V-labs
Inc., Covington, LA), or overnight in 10mM LSTb (Sigma, St.
Louis, MO). All crystals were flash-cooled at 100K using 20%
glycerol as the cryo-protectant. Datasets were collected at
Advanced Photon Source (APS) beamlines 22 ID and BM at
100K. Data were processed with the DENZO-SACLEPACK suite
[46]. Statistics for data collection are presented in Table 1.
Structure determination and refinement
The
structure
of
NY107
was
determined
by
molecular
replacement with Phaser [47] using the structure of the avian
H7 (Av-H7) from A/turkey/Italy/2002, pdb:1TI8 (HA1, 78%
identity; HA2, 90% identity) as the searching model. One HA
trimer occupies the asymmetric unit with an estimated solvent
content of 58% based on a Matthews’ coefficient (Vm) of 2.9 A˚ 3/
Da. Rigid body refinement of the trimer led to an overall R/Rfree
of 28.6%/37.4%. The model was then ‘‘mutated’’ to the correct
sequence and rebuilt by Coot [48], then the protein structures
were refined with REFMAC [49] using TLS refinement [50]. The
final models were assessed using MolProbity [51]. The three
complex structures were refined and evaluated using the same
strategy. All statistics for data processing and refinement are
presented in Table 1. Electron density maps (2fo-fc) were
generated in Refmac [49] while simulated annealing omit maps
were generated by sa-omit-map, a part of the Crystallography and
NMR System (CNS) software [52].
Virus generation
Wild type and mutant viruses of NY107 (H7N2) and A/
Netherland/219/2003 (H7N7) were generated from plasmids by a
reverse genetics approach [53]. To generate viruses with amino
acid insertion or substitution in the HA, mutations were
introduced into plasmid DNA with an overlap extension PCR
approach [54]. Viruses derived by plasmid transfection of HK293
cells were propagated in eggs. The genomes of resulting virus
stocks were sequenced to detect the emergence of possible variants
during amplification.
Glycan binding analyses
Glycan microarray printing and recHA analyses have been
described previously [24,30,44,55] (see Table 2 for glycans used
for analyses in these experiments). Virus were analyzed on the
microarray as described previously [30].
PDB accession codes
The atomic coordinates and structure factors of NY107 are
available from the RCSB PDB under accession codes 3M5G for
the unliganded NY107, 3M5H for the NY107 with 39-SLN and
3M5I and 3M5J for NY107 with 69SLN and LSTb, respectively.
Accession/ID numbers for genes/proteins used in this
work
A/New York/107/03 (H7N2), Genbank: ACC55270; A/
Netherlands/219/03 (H7N7), Genbank: AAR02640; A/Hong
Kong/1-9/68 (H3N2), 2HMG; A/Duck/Ukraine/1/63 (H3N8),
PDB: 1MQL; A/South Carolina/1/18 (H1N1), PDB: 1RD8; A/
Puerto Rico/8/34 (H1N1), PDB: 1RU7; A/Swine/Iowa/15/30
(H1N1), PDB: 1RUY; A/Singapore/1/1957
(H2N2), PDB:
2WRC; A/Viet Nam/1203/04 (H5N1), PDB: 2FK0; A/Duck/
Singapore/3/97 (H5N3), PDB: 1JSM; A/Swine/Hong Kong/9/
98 (H9N2), PDB: 1JSD; A/Turkey/Italy/8000/02 (H7N3), PDB:
1TI8; C/Johannesburg/1/66, 1FLC.
Supporting Information
Figure S1
Sequence alignment of selected structurally available
HAs. Human H3 (PDB: 2HMG), Avian H3 (PDB: 1MQL), 1918-
Human H1 (PDB: 1RD8), 1934-Human H1 (PDB: 1RU7), Swine
H1 (PDB: 1RUY), 1957-Huamn H2 (PDB: 2WRC), Human H5
(PDB: 2FK0), Avian H5 (PDB: 1JSM), Swine H9 (PDB: 1JSD),
and Avian H7 (PDB: 1TI8) were used in the alignments. The
fusion domain of HA1 is highlighted in magenta, the vestigial
esterase domain is highlighted in green, the receptor binding
domain is highlighted in blue, and the fusion domain of HA2 is
highlighted in red. Residue numbering is based on the H3 HA
sequence.
Found at: doi:10.1371/journal.ppat.1001081.s001 (2.84 MB TIF)
Figure S2
Expression and purification of NY107. SDS-PAGE
reveals that NY107 was expressed as the HA0 form with a mass
approximately 60kDa (middle lane). Thrombin cleavage resulted in
an unexpected reduction in band size to a HA1/HA2 profile (right
lane) with possible multiple glycoforms for the HA2 clearly present.
Found at: doi:10.1371/journal.ppat.1001081.s002 (0.23 MB TIF)
Figure S3
Comparison of glycan binding to NY107 with other
HAs. A. Overlap of a2-3 ligands binding in the receptor binding
site from NY-H7 (green), Av-H3 (orange), 1930-Hu-H1 (magenta),
1957-Hu-H2 (cyan), Av-H5 (yellow), and Sw-H9 (grey). B.
Overlap of a2-6 linkage ligands binding in the receptor binding
site from NY-H7 (green), Av-H3 (orange), 1930-Hu-H1 (magenta),
1957-Hu-H2 (cyan), Av-H5 (yellow), and Sw-H9 (grey).
Found at: doi:10.1371/journal.ppat.1001081.s003 (2.55 MB TIF)
Figure S4
Simulated annealing omit maps of the receptor
binding site (contoured at 1s). A. NY107 (blue), B. NY107-39SLN
(orange), C. NY107-69SLN (red), and D. NY107-LSTb (magenta).
The protein model is shown in cartoon, and the residues involved
in the binding to receptor analogs were shown in sticks. Maps were
generated using version 1.2 of the Crystallography and NMR
System (CNS) software.
Found
at:
doi:10.1371/journal.ppat.1001081.s004
(1.93
MB
TIF)
Structure of a North American H7 Hemagglutinin
PLoS Pathogens | www.plospathogens.org
9
September 2010 | Volume 6 | Issue 9 | e1001081
Figure S5
Comparison of NY107 RBS to HEF. Overlap of RBS
from NY107 (green), Av-H7 (marine) and HEF (magenta).
Found at: doi:10.1371/journal.ppat.1001081.s005 (1.12 MB TIF)
Table S1
Comparison of r.m.s.d. (A˚ ) for different HA domains.
For analyzing differences in the overall structure, r.m.s.d. values
were calculated between monomers or domains of different HA’s,
after the Ca atoms of the HA2 domains were superposed by
sequence and structural alignment onto the equivalent domains of
NY107.
Found
at:
doi:10.1371/journal.ppat.1001081.s006
(0.04
MB
DOC)
Table S2
Comparison of r.m.s.d. (A˚ ) for individual domains.
Each domain was superimposed separately to determine how the
individual NY107 domains compared to equivalent domains in the
other structures.
Found
at:
doi:10.1371/journal.ppat.1001081.s007
(0.04
MB
DOC)
Table S3
Molecular interactions between NY107 and receptor
analogs. The hydrogen bond cutoff is 3.8 A˚
for the listing
interactions.
Found
at:
doi:10.1371/journal.ppat.1001081.s008
(0.07
MB
DOC)
Table S4
Glycan array differences between NY107, the fully
restored NY107-ins, and NL219 (virus and rHA). The color
coding in the left hand column reflects the same coloring scheme
used in Figures 2 and 3. Significant binding of samples to glycans
are qualitatively estimated based on relative strength of the signal
for the data shown in Figures 2 and 3 Strong (+++), weak (+).
Found
at:
doi:10.1371/journal.ppat.1001081.s009
(0.19
MB
DOC)
Acknowledgments
The authors would like to thank the staff of SER-CAT sector 22 for their
help in data collection. We also thank WHO Global Influenza Surveillance
Network for providing NY107 and NL219 viruses from which the reverse
genetics viruses were generated. Glycan microarray data presented here
will be made available on-line through the CFG web site upon publication
(www.functionalglycomics.org). The findings and conclusions in this report
are those of the authors and do not necessarily represent the views of the
Centers for Disease Control and Prevention or the Agency for Toxic
Substances and Disease Registry.
Author Contributions
Conceived and designed the experiments: HY LMC PJC ROD JS.
Performed the experiments: HY LMC PJC JS. Analyzed the data: HY
LMC PJC ROD JS. Wrote the paper: HY LMC PJC ROD JS.
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Structure of a North American H7 Hemagglutinin
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|
3M5J
|
Crystal structure of a H7 influenza virus hemagglutinin complexed with LSTb
|
Structures of Receptor Complexes of a North American
H7N2 Influenza Hemagglutinin with a Loop Deletion in
the Receptor Binding Site
Hua Yang, Li-Mei Chen, Paul J. Carney, Ruben O. Donis, James Stevens*
Influenza Division, Centers for Disease Control and Prevention, Atlanta, Georgia, United States of America
Abstract
Human infections with subtype H7 avian influenza viruses have been reported as early as 1979. In 1996, a genetically stable
24-nucleotide deletion emerged in North American H7 influenza virus hemagglutinins, resulting in an eight amino acid
deletion in the receptor-binding site. The continuous circulation of these viruses in live bird markets, as well as its
documented ability to infect humans, raises the question of how these viruses achieve structural stability and functionality.
Here we report a detailed molecular analysis of the receptor binding site of the North American lineage subtype H7N2 virus
A/New York/107/2003 (NY107), including complexes with an avian receptor analog (39-sialyl-N-acetyllactosamine, 39SLN)
and two human receptor analogs (69-sialyl-N-acetyllactosamine, 69SLN; sialyllacto-N-tetraose b, LSTb). Structural results
suggest a novel mechanism by which residues Arg220 and Arg229 (H3 numbering) are used to compensate for the deletion
of the 220-loop and form interactions with the receptor analogs. Glycan microarray results reveal that NY107 maintains an
avian-type (a2-3) receptor binding profile, with only moderate binding to human-type (a2-6) receptor. Thus despite its
dramatically altered receptor binding site, this HA maintains functionality and confirms a need for continued influenza virus
surveillance of avian and other animal reservoirs to define their zoonotic potential.
Citation: Yang H, Chen L-M, Carney PJ, Donis RO, Stevens J (2010) Structures of Receptor Complexes of a North American H7N2 Influenza Hemagglutinin with a
Loop Deletion in the Receptor Binding Site. PLoS Pathog 6(9): e1001081. doi:10.1371/journal.ppat.1001081
Editor: Fe´lix A. Rey, Institut Pasteur, France
Received March 16, 2010; Accepted July 28, 2010; Published September 2, 2010
This is an open-access article distributed under the terms of the Creative Commons Public Domain declaration which stipulates that, once placed in the public
domain, this work may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose.
Funding: This work was funded by the Centers for Disease Control and Prevention. Use of the Advanced Photon Source at Argonne National Laboratory was
supported by the U. S. Department of Energy, Office of Science, Office of Basic Energy Sciences, under Contract No. DE-AC02-06CH11357. Glycan microarrays as
well as glycans for direct binding experiments were produced for the Centers for Disease Control by the CFG funded by National Institute of General Medical
Sciences Grant GM62116. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: fwb4@cdc.gov
Introduction
Influenza is an acute respiratory virus that infects up to 20% of
the population in the United States, resulting in ,36,000 deaths
annually [1,2]. The two membrane glycoproteins on the surface of
influenza A virus, hemagglutinin (HA), which functions as the
receptor binding and membrane fusion glycoprotein in cell entry,
and neuraminidase (NA), which functions as the receptor destroying
enzyme in virus release, form the basis for defining subtypes [3]. To
date, 16 HA (H1–H16) and 9 NA (N1–N9) have been identified in
avian species [4], while in the last century, only three subtypes,
H1N1 in 1918 and 2009, H2N2 in 1957, and H3N2 in 1968 [5,6,7],
have successfully adapted to humans. Hemagglutinin binds to sialic
acid (SA) glycans present on host cell surfaces. The receptors on
epithelial cells of the human upper respiratory tract are mainly
a2-6-linked SA moieties [8]. Since avian influenza viruses pre-
dominately bind a2-3-linked SA, and human influenza viruses
preferentially bind to a2-6-linked SA, human infection by avian
influenza viruses is rare [9]. However, since 1997 a growing number
of human cases of avian influenza infection have been reported [10],
including H5N1, H7N2, H7N3, H7N7, and H9N2 strains [11].
Although the current situation with the pandemic H1N1 influenza
virus dominates public health efforts, the prospect of a novel
pandemic emerging from these isolated cases continues to be a
major public health threat around the world.
Early cases of human infection by H7 influenza viruses are reported
as far back as 1979 [12,13]. Since 2002, multiple outbreaks and
human infections of H7 subtype viruses; within both Eurasian and
North American lineages have been reported. In the Netherlands in
2003, a highly pathogenic avian influenza (HPAI) H7N7 outbreak
resulted in more than 80 cases of human infections, including one
fatality [14,15]. In New York in 2003, a single case of human
respiratory infection of H7N2 was reported [16] and in British
Columbia in 2004, an H7N3 virus caused two cases of conjunctivitis
[17,18]. More recently in 2007, the United Kingdom reported several
cases of low pathogenic avian influenza (LPAI) H7N2 virus infections
that caused influenza-like illness and conjunctivitis [19].
Since 1996, H7 viruses of the North American lineage have
been circulating in regional live bird markets [20], containing a
24-nucleotide deletion resulting in an eight amino acid deletion in
the receptor-binding site (RBS) of HA (Figure S1). The recent
human infections with H7 in North America have raised public
health concerns as to how these viruses adapt to such a dramatic
structural change while remaining one of the predominant
circulating viral strains. A recent study of H7 viruses isolated
from previous outbreaks revealed efficient replication in both
mouse and ferret animal models [21]. In particular, ferret studies
with A/New York/107/2003 (NY107), an H7N2 virus isolated
from a man in New York, not only showed efficient replication in
the upper respiratory tract of the ferret but also the capacity for
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intra-species transmission by direct contact [21,22]. Interestingly,
both an increased preference for a2-6 and decreased preference for
a2-3-linked sialosides of this virus compared to the other avian
influenza viruses was shown by previous glycan microarray analysis
but less so by a competitive solid-phase binding assay [22,23].
Here we report a detailed molecular analysis of the RBS of the
HA from North American lineage H7N2 virus, NY107, including
glycan microarray analyses and structural analyses of the HA in
complex with an avian receptor analog (39-Sialyl-N-acetyllactosa-
mine, 39SLN) and two human receptor analogs (69-Sialyl-N-
acetyllactosamine, 69SLN; Sialyllacto-N-tetraose b, LSTb). These
results provide important insight into the interaction of H7 HAs
with both avian and human hosts.
Results
Overall structure
By using x-ray crystallography, the structure of H7 HA from the
NY107 virus was determined to 2.6 A˚ resolution (Table 1). In
addition, we also report three H7 HA receptor complex structures,
with avian receptor analog (39SLN) to 2.7 A˚ resolution and with
human receptor analogs (69SLN and LSTb) to 3.0 A˚ and 2.6 A˚
resolution, respectively (Table 1). The overall structure of NY107
is similar to other reported HA structures with a globular head
containing the RBS and vestigial esterase domain, and a
membrane proximal domain with its distinctive, central helical
stalk and HA1/HA2 cleavage site (Figure 1A). Although five
asparagine-linked glycosylation sites are predicted in the NY107
HA monomer, interpretable electron density was observed at only
two sites, Asn38 in HA1 and Asn82 in HA2 (all residue numbers
are based on H3 numbering). At these sites, only one or two N-
acetyl glucosamines could be interpreted.
During viral replication, HA is synthesized as a single chain
precursor (HA0) and cleaved by specific host proteases into the
infectious HA1/HA2 form. In baculovirus expression systems,
highly pathogenic HAs, with a polybasic cleavage site, are
expressed as an HA1/HA2 form [24], whereas HAs with
monobasic cleavage sites (single Arg) from low pathogenic viruses
are expressed as the HA0 form [25]. NY107 is regarded as a low
pathogenic virus, and as expected, was produced in the HA0
form (Figure S2). However, subsequent digestion with thrombin
protease to remove the His-tag resulted in cleavage to a profile on
SDS-PAGE comparable to that of an HA1/HA2 form (Figure S2).
A comparison of the NY107 cleavage site with the consensus
cleavage pattern in the MEROPS database (http://merops.
sanger.ac.uk) suggests it to be a possible thrombin cleavage site.
Based on their molecular phylogenies, HAs are divided into two
groups and five clades: group 1 includes H8, H9, and H12; H1,
H2, H5, and H6; H11, H13 and H16; group 2 includes H3, H4,
and H14; H7, H10 and H15 [26]. Among all available HA
structures, we selected ten representative HAs from both avian and
human subtypes for structural analysis. As expected, NY107 HA is
structurally very similar to the Avian-H7 in all comparisons and
closely related to H3, the other group 2 members used in the
analyses (Tables S1 and S2).
The receptor binding site
The RBS is at the membrane distal end of each HA monomer
and its specificity for sialic acid and the nature of its linkage to a
vicinal galactose residue is a major determinant of host range-
restriction. The consensus RBS for all current HAs is composed of
three major structural elements: a 190-helix (residues 188–194), a
220-loop (residues 221–228), and a 130-loop (residues 134–138).
In addition, highly conserved residues (Tyr98, Trp153, His183,
and Tyr195) form the base of the pocket.
Although the NY107 RBS is similar to other subtypes (H1, H2,
H3, H5, and H9), a previously observed specific feature of H7 HAs,
is also observed in the NY107 150-loop region: two residues inserted
at position 158 result in this loop protruding more than 6A˚ towards
the binding site compared to other subtype HAs (Figure 1B and
Table S2) [27]. More interestingly, the eight amino acid deletion,
only found in the North American lineage H7s, from position 221 to
228 (Figure S1), resulted in a complete loss of the 220-loop
(Figure 1B). Sequence alignment shows that Arg220 and Arg229 are
conserved in all influenza A HA subtypes (Figure S1), but structural
alignment of NY107 HA shows Arg220 occupying the Gly228
position, and the much shorter loop turns at residue Pro217
(Figure 1C). The Ca distance between NY107 Arg220 and its
homolog in the Av-H7 structure (PDB: 1TI8) [27] is 5.8A˚ , and they
point in opposite directions (Figure 1C). The side chain direction of
Av-H7 Arg220 is almost parallel with the beta sheet after Arg229,
whereas the NY107 Arg220 points downward to the binding pocket.
The Ca position of Arg229 in both H7 structures remains the same,
except the side chain in the NY107 swings away by about 5.9A˚
(Figure 1C) and could help to stabilize this region by forming a
hydrogen bond to the mainchain carbonyl of Gln210 in the
neighboring monomer. In the absence of the 220-loop in NY107
HA, upon glycan binding the long side chain of Arg220 compensates
for its loss and is displaced 4A˚ upward to form hydrogen bonds with
receptor analogs inside the binding pocket (Figure 1D).
Effect of loop truncation on the receptor binding
specificity of NY107
Previously, mutations in the HA receptor binding domains of
H1N1 (Glu190Asp/Gly225Asp) and H2N2/H3N2 (Gln226Leu
and Gly228Ser) subtypes were responsible for adaptation of these
viruses to pandemic strains [24,28,29,30]. Due to missing residues
221–228 in the NY107 HA RBS, neither mechanism for
adaptation is possible. Thus, in order to look more closely at the
role of the missing loop and its effect on receptor specificity, we
first subjected the recombinant HA (recHA) to glycan microarray
analyses and compared it to a reverse genetics-derived NY107
virus, and a co-circulating Eurasian virus and recHA, A/
Author Summary
Influenza virus adaptation to different hosts usually results
in a switch in receptor specificity of the viral surface coat
protein, hemagglutinin. Indeed, the hemagglutinin sub-
types from the last two human influenza pandemics of the
20th Century (H2 in 1957 and H3 1968) both adapted
successfully to human-type receptor specificity through
only two amino acid mutations in the receptor binding
pocket (Glutamine226RLeucine and Glycine228RSerine).
The recent human infections reported with other avian
subtypes such as H5, H7 and H9 have raised public health
concerns and focused efforts on identifying potential
subtypes from which a future pandemic strain may
emerge. Since 1996, H7 viruses of the North American
lineage have been circulating in regional live bird markets,
containing an eight amino acid deletion in the receptor-
binding site of HA. Here we report a detailed structural
analysis of the receptor binding site of a hemagglutinin
from the North American lineage of H7N2 viruses, in
complex with avian and human receptor analogs, to
understand how these viruses have adapted to such a
dramatic structural change in the binding site while
remaining one of the predominant circulating viral strains.
Structure of a North American H7 Hemagglutinin
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Netherlands/219/2003 (NL219), that has the consensus avian
sequence in the 220-loop and it also infected a human [15].
Glycan microarray analysis of recombinant NY107 (Figure 2A
and Table 2) revealed a highly restricted binding profile with
strong binding to only a2-3 sulfated (#4–8), a2-3 branched (#9–
11) and mixed a2-3/a2-6 branched sialosides (#60–64) as well as
to the long linear sialyl di- and tri-lactosamines (#22, 24). Weak
binding was also observed (above background) to other a2-3
glycans on the array. The recombinant NY107 also revealed a
strict glycan binding preference to only one a2-6 glycan, the
internal structure, Galb1-3(Neu5Aca2-6)GlcNAcb1-3Galb1-4Glc
(#58; LSTb) (Figure 2A), a glycan highlighted in a previous study
[22]. The virus with higher valency and avidity revealed stronger
binding to all a2-3 groups, in addition to the branched di-sialyl a2-
6 biantennary structures (#46–48) as well the LSTb (#58)
(Figure 2B and Table 2). In contrast, the NL219 recHA (Figure 2C
and Table 2) bound well to only the avian a2-3 containing sialyl-
glycans (sulfated, branched, linear and fucosylated). Its corre-
sponding virus also reflected this specificity although it also
revealed strong binding to a2-3 N-glycolylneuraminic acid
(Neu5Gc) containing glycans (#66–70) (Figure 2D and Table 2).
To further assess the effect of the missing 220 loop on HA
structural stability and receptor specificity it was essential to
evaluate these functions on the ancestral HA containing the full
length 220-loop. To this end, we engineered an HA with an avian
H7 consensus (PQVNGQSG) 220-loop re-introduced (NY107-
220ins) into the NY107 HA and recovered this virus by reverse
genetics. Compared to the NY107 virus (Figure 2A) glycan
microarray analyses of the resulting NY107-220ins virus (Figure 3A
and Table 2) revealed a decrease in binding to branched (#9–11)
and linear (#12–27) a2-3 sialosides and a loss of binding to the
branched di-sialyl a2-6 biantennary structures (#46–48), LSTb
(#58) as well as the mixed a2-3/a2-6 branched sialosides (#60–
64). In addition, sequence analysis of the NY107-220ins HA
revealed the presence of quasispecies in the second position of the
inserted loop, P(Q/K)VNGQSG, suggesting that re-introduction
of the loop alone is not tolerated and does not create an avian-type
binding profile. Thus other amino acid substitutions in the HA
might have co-evolved with the deletion of the 220 loop to help
stabilize the RBS/HA to maintain functionality.
When viruses containing this 220-loop deletion emerged in North
America in the mid 90’s, four additional amino acid substitutions,
Gly114Arg, Asp119Gly, Gly186Glu and Gly205Arg, in the HA1 as
well as an Asp19Asn in the HA2 chain were also introduced to most
of the circulating isolates. Of these, Gly186Glu and Gly205Arg in
the HA1 are close to the RBS, at the monomer interface, and could
potentially modulate its structure and/or function. NY107 viruses
with a restored consensus 220-loop and a single Glu186Gly (NY107-
ins-186) or Arg205Gly (NY107-ins-205) substitution as well as the
Glu186Gly/Arg205Gly double substitution (NY107-ins-186/205)
were derived by reverse genetics and evaluated. Glycan microarray
analysis for the three resulting viruses revealed similar glycan
binding profiles with increased binding to a2-3 sialosides, including
mixed a2-3/a2-6 branched sialosides (#60–64), a2-3 Neu5Gc
(#66–70), but limited binding to the a2,6 sialosides (Figures 3B, 3C,
3D), resulting in a binding profile virtually identical to that of the
NL219 virus and other avian influenza viruses (Figure 2D) [30].
Sequence analysis of the three reverse genetics derived viruses
revealed no mutations/quasispecies in the HAs of either the NY107-
ins-186 or the NY107-ins-186/205 virus stocks, indicative of
replication fitness. For the NY107-ins-205 virus however, a
Glu186Gly substitution emerged in the HA after only two passages
in eggs following recovery from DNA transfection, indicating the
importance of the co-variant position 186 with respect to HA
functionality/glycan specificity. Altogether, the data indicates that
the H7 subtype avian influenza viruses that were circulating in
Table 1. Data collection and refinement statistics.
NY107
NY107+39SLN
NY107+69SLN
NY107+LSTb
Data collection
Space group
P212121
P212121
P212121
P212121
Cell dimensions (A˚)
66.96, 115.92, 251.61
67.80, 116.70, 249.84
66.60, 116.58, 250.68
67.08, 116.52, 251.95
Resolution (A˚)
50-2.6 (2.69-2.60)a
30-2.7 (2.80-2.70)
50-3.0 (3.11-3.0)
50-2.6 (2.69-2.60)
Rsym or Rmerge
10.6 (41.3)
14.6 (48.6)
14.3 (35.4)
12.2 (31.5)
I/s
39.6 (2.0)
24.3 (1.7)
34.2 (8.2)
40.5 (9.9)
Completeness (%)
99.2 (99.0)
99.3 (94.6)
92.3 (75.6)
91.3 (86.2)
Redundancy
7.2 (6.2)
5.8 (5.5)
4.9 (4.4)
10.9 (11.2)
Refinement
Resolution (A˚)
50-2.6 (2.67-2.60)
30-2.7 (2.77-2.70)
50-3.0 (3.08-3.00)
50-2.6 (2.67-2.60)
No. of reflections (total)
57285
51770
33421
53603
No. of reflections (test)
3053
2769
1779
2842
Rwork/Rfree
21.7/25.6
21.4/26.4
20.5/26.0
20.4/24.7
No. of atoms
11795
11878
11648
12108
r.m.s.d.- bond length (A˚)
0.006
0.006
0.008
0.006
r.m.s.d.- bond angle (u)
0.905
0.974
1.085
0.859
MolProbityb scores
Favored (%)
96.9
96.5
94.3
97.1
Outliers (%) (No. of residues)
0.1 (1/1434)
0.0 (0/1429)
0.1 (2/1433)
0.1 (2/1435)
aNumbers in parentheses refer to the highest resolution shell.
bReference [51].
doi:10.1371/journal.ppat.1001081.t001
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aquatic birds and poultry in North America before 1996 exhibited a
classic avian a2-3 sialoside binding preference. In order for the 220-
loop deletion to be tolerated, concurrent Gly186Glu and Gly205Arg
substitutions in the vicinity of RBS of HA emerged to achieve a
restricted a2-3 binding profile and only a moderate/limited increase
in binding to branched di-sialyl a2-6 biantennary structures (#46–
48) as well the a2,6 internal sialoside, LSTb (#58).
NY107 avian receptor complex
To understand from a structural perspective how NY107
interacts with host receptors, we solved the structure of NY107
in complex with an avian and two human receptor analogs.
For the avian receptor analog, 39SLN, the electron density
maps revealed well-ordered features for the Sia-1, Gal-2, and
GlcNAc-3 in the NY107 HA complex structure (Figure 4A).
Structural comparison of NY107 HA binding to other, H1, H2,
H3, H5, and H9 subtypes (Figure S2A) revealed that 39SLN
binding to NY107 resembled binding of the other published
HAs. Indeed, the terminal Sia-1 moiety is positioned almost
identically in all structures, and forms the majority of hydrogen
bonds and contacts with residues in the RBS (Figure 4A and
Table S3).
Figure 1. NY107 HA monomer and comparison of its RBS to other HA structures. (A) One monomer is shown with the HA1 chain colored in
green and the HA2 chain in cyan. The location of the receptor binding site and the HA1/HA2 cleavage site are circled. (B) The superposition of
receptor binding domains of NY107 (green), Av-H7 (marine), 1918-Hu-H1 (magenta), Hu-H5 (yellow), Hu-H3 (orange), and Sw-H9 (grey). The proximity
of Arg220 and Gln226 are highlighted. Three structural elements comprising this binding site are labeled. The two major differences are the extended
150-loop and the deletion of 220-loop of NY107. (C) Overlap of NY107 (green) and Av-H7 (marine) (PDB: 1TI8) illustrates the compensatory effect of
R220 bringing it close to the position occupied by G228 in the avian HA. (D) Overlap of the NY107 (green), NY107- 39SLN (orange), NY107-69SLN (red),
and NY107-LSTb (magenta) structures. All the figures were generated and rendered with the use of MacPyMOL [56].
doi:10.1371/journal.ppat.1001081.g001
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Published avian HA structures with an intact 220-loop form
very close interactions with Gal-2 of 39SLN via residue Gln226
which is important in receptor specificity and host adaptation. For
example, in the avian H7/39SLN HA structure it interacts with
Gal-2 O4 [31]. In the NY107 HA structure, although Gln226 is
absent and no other residue occupies the same space as Gln226
(Figure 1B), Arg220 does forms a hydrogen bond between Arg220
NH2 and Gal-2 O4 (Figure 4A). Interestingly, although there was
interpretable density for the GlcNAc-3 (Figure 4A and Figure
S4B), no hydrogen bonding was apparent between the HA and the
GlcNAc-3, which is consistent with other reported structures [32].
Thus, for binding to avian receptors, the trans conformation of
a2-3 linkages is essential and perhaps only the first two saccharides
are required. Indeed, due to the absence of 220-loop in the NY107
HA structure, the ‘‘aperture’’ of the RBS formed by 220-loop and
130-loop in regular HAs is increased by ,10 A˚ , so that the
branched, internal, and perhaps more complicated glycans might
be accommodated more efficiently.
NY107 human receptor complexes
In the NY107/69SLN complex, only Sia-1 and Gal-2 are
ordered (Figure 4B). The Sia-1 remains in the same position as
previously analyzed glycan/HA complexes from H1, H2, H3, H5,
and H9 (Figure S3B), whereas the Av-H7 complex structure with
Sialyllacto-N-tetraose c (LSTc) did not reveal any density for the
Sia-1 in the receptor binding site [31]. The Gal-2 position varies
significantly among different subtypes. Compared to the human-
adapted H1 HA [32], Gal-2 in the NY107 HA is 3A˚ higher, and
thus is further from the protein (Figure S3B). In NY107, the Gal-2
only forms an intramolecular, saccharide-saccharide interaction
with Sia-1. The poor electron density map and fewer interactions
with protein residues suggest that the cis conformation of a2-6
Figure 2. Receptor specificity of NY107 recHA and virus. Glycan microarray analysis of recombinant NY107 HA (A) and NY107 virus (B)
compared to the recHA (C) and virus (D) from a Eurasian lineage A/Netherlands/219/2003 H7 influenza virus that was circulating in the same year and
also infected a human. Colored bars highlight glycans that contain a2-3 SA (blue) and a2-6 SA (red), a2-6/a2-3 mixed SA (purple), N-glycolyl SA
(green), a2-8 SA (brown), b2-6 and 9-O-acetyl SA, and non-SA (grey). Error bars reflect the standard error in the signal for six independent replicates
on the array. Structures of each of the numbered glycans are found in Table S4.
doi:10.1371/journal.ppat.1001081.g002
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linkages in 69SLN trisaccharides show a reduced binding affinity
with NY107.
Glycan array results with NY107 revealed a strong binding
signal for the internal a2-6 sialoside, LSTb. To further investigate
this interaction, we solved the structure of the NY107/LSTb
complex. The final model contained Sia-1, NAG-2, Gal-3, and
Gal-5 in the RBS. Although glycan microarray data indicated
NY107 to have a specific affinity for LSTb, few interactions were
apparent from the crystal structure. Sia-1 still forms multiple
hydrogen bonds with residues in the RBS (Table S3 & Figure 4C).
The branched Gal-5 interacts with Ser137, to help stabilize the
LSTb binding. However, Arg220 and Lys193, the two residues
showing close binding with 39SLN, did not form any hydrogen
bonds with LSTb. In the structure, Gal-5 also interacts with a
crystal packing symmetry mate and thus the flexibility of whole
LSTb may be restricted. In solution, with more freedom, the
LSTb should be able to tilt closer to the RBS, and thus Glc-4 may
have more interactions with the 190-helix than seen in the crystal
structure.
Discussion
Human infections by avian influenza viruses, including H7
subtypes, continue to pose a major public health threat. Although
the
species
barrier
prevents
avian
influenza
viruses
from
widespread infection of the human population, the molecular
determinants of efficient interspecies transmission and pathoge-
nicity are still poorly understood. The viral coat protein HA
however, is perhaps a critical molecule since previous pandemic
viruses modified their receptor specificity and overcame the
interspecies barrier to spread in the human population. Although
HA structures alone and in complex with receptor analogs provide
considerable insight into receptor binding, it is clear that HAs from
different species and subtypes have significant structural variation.
Indeed, low-pathogenic H7N2 avian influenza viruses with an 8
amino acid deletion within its RBS started to circulate in live-bird
markets in the northeast United States in 1996. Despite what one
would consider a debilitating mutation, these viruses have been
reported as the predominant isolate [33]. Whether such a deletion
contributed to their evolutionary success and how are an
important questions, especially in light of NY107’s ability to
produce respiratory illness in humans [16], as well as its reported
increased affinity for human-type receptors and ability for contact
transmission in ferrets [21]. To try to help answer these questions,
we have analyzed the molecular structures of NY107 and its
complexes with receptor analogs to explain receptor specificity at
the molecular level.
The crystal structures of NY107 and its complexes with both
avian and human receptor analogs describe a mechanism as to
how an influenza virus might adapt by dramatically altering its
RBS, and still be functional. Arg220 of the HA1 chain of NY107
compensates for the loss of the 220-loop, by forming hydrogen
bonds with Gal-2 from the avian analog (binding was not observed
in either of the structures complexes with the human analogs).
However, in the LSTb complex, branched Gal-5 forms extra
interactions with the 130-loop, thus improving the binding
preference for this particular glycan. Consistent with the structural
evidence, glycan microarray analyses of NY107 revealed a strong
binding preference for the branched a2-6 sialoside, LSTb. Except
for the absence of the 220-loop, other key residues within the RBS
are conserved in NY107 and thus, direct interactions with sialic
acid are maintained.
The 220-loop is recognized as one of the three crucial structural
elements in the RBS. Aside from the North American lineage
H7N2 viruses, which have been circulating with a deletion (221–
228) in this loop, there has been one other report describing a
seven amino acid deletion (224–230) in a laboratory generated
H3N2 escape mutant which was reported to have a slightly
Table 2. Comparison of the sialoside receptor specificity of the HAs from H7 influenza viruses.
Glycan Group
Graph
Numbera
NY107
RecHA
NY107
Virus
NY107-ins
Virus
NY107-ins
E186G Virus
NY107-ins
R205G Virus
NY107-ins E186G/
R205G Virus
NL219
RecHA
NL219
Virus
a2-3
Sulfated
4–8
+++b
+++
+++
+++
+++
+++
+++
+++
Branched
9–11
+++
+++
+
+++
+++
+++
+++
+++
Linear
12–27
+
+++
+
+++
+++
+++
+++
+++
Fucosylated
28–34
2
+++
+++
+++
+++
+++
+++
+++
a2-6
Sulfated
41
2
2
2
2
2
2
2
2
Branched mono-sialyl
42–45, 49
2
2
2
2
2
2
2
2
Branched di-sialyl
46–48
2
+++
2
2
2
2
2
2
Linear
50–56
2
2
2
2
2
2
2
2
Internal
58–59
+++
+++
2
2
2
2
2
2
Other
Sialic acid
1–2
2
+++
+
2
2
2
2
2
a2-3/a2-6 Branched
60–64
2
2
2
+++
+++
+++
+++
+++
Neu5Gcc
65–72
2
2
2
+++
+++
+++
2
+++
aMembers of each group are identified according to the graph number used in the microarray data in Figures 2 and 3 and correspond to numbers in the complete
glycan list (Table S4).
bBinding of samples to glycan subclasses are qualitatively estimated based on relative strength of the signal for the data shown in Figures 2 and 3: strong (+++), weak
(+), absent (2).
cN-glycolylneuraminic acid.
doi:10.1371/journal.ppat.1001081.t002
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6
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increased affinity for a2-3-linked glycans by hemagglutination
assay [34]. Meanwhile, the equivalent region in the hemaggluti-
nin-esterase-fusion (HEF) protein of influenza C virus reveals a
rearrangement resulting in a truncated 260-loop in its RBS (Figure
S5) [35]. However, without structural data with appropriate
receptor analogs, it is not possible to compare the role of these loop
variants in receptor binding to the H7 HA structure described
here.
When compared to NL219, another co-circulating H7 avian
virus HA (Figure 2C and D), overall binding to a2-3-linked
glycans was markedly reduced, while increased binding to a2-6-
linked receptors was only marginal. However, these results focus
attention on only 2 sub-classes of human-type receptors that may
be important for infection (and transmission in ferrets). The
NY107 virus interaction with biantennary glycans (Figure 2B),
although weak (not seen in Figure 2A with recHA), is a possible
route for virus entry as biantennary structures are common on
tissues, i.e. glycan profiling data from human lung tissue on the
Consortium for Functional Glycomics (CFG) web site. In addition,
the internal sialoside, LSTb, was observed in both virus and
recHA microarray data, suggesting this type of glycan has good
affinity for this HA. The significance of this is unknown since
LSTb has only been described in human milk [36].
Interestingly, NY107 and NL219 virus receptor binding and
specificity has been addressed previously using glycan microarray
analysis that reported a significantly increased preference for a2-6
and decreased preference for a2-3-linked sialosides [22]. In
addition, the same viruses were also included in a recent study
from Gambaryan et al. using a competitive solid-phase binding
assay [23]. Our findings confirm and extend the receptor binding
specificity reported by these authors in that they reported both
viruses binding to sulfated sialylglycans with a lactosamine (Galb1-
4GlcNAc core and reported only a moderate binding affinity for
a2-6-sialyllactosamine, the human-type receptor analog used in
their assay.
The 220-loop is an integral feature of the receptor binding site,
and thus one would predict that such a deletion might have
compromised this strain to be deleted from the population of
circulating viruses. However, this was not the case [33] and its
existence appears to be in part due to the additional mutations at
Figure 3. Effect of 220-loop deletion and additional RBS mutations on NY107 receptor specificity. NY107 was engineered to restore the
220-loop to a consensus full-length HA from 1996 (A) and additional co-variant amino acid substitutions, Glu186Gly (B), Arg205Gly (C) and the double
mutant Glu186Gly/Arg205Gly (D) to restore, on the NY107 framework, an HA RBS found in viruses prior to the introduction of the deletion in North
American viruses. Colored bars group glycans as described in Figure 3. Error bars reflect the standard error in the signal for six independent replicates
on the array. Structures of each of the numbered glycans are found in Table S4.
doi:10.1371/journal.ppat.1001081.g003
Structure of a North American H7 Hemagglutinin
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7
September 2010 | Volume 6 | Issue 9 | e1001081
positions 186 and 205. Restoration of the loop with either or both
residues mutated back to the pre-1994 consensus sequence
resulted in a classic avian influenza virus binding profile. The
emergence of the Glu186Gly mutation in the HA of the NY107-
ins-205 mutant after only two passages of the rescued virus in
eggs, also indicates the importance of these positions for HA
functionality/glycan specificity. Analysis of the structural data
reveals that positions 186 and 205 are on opposite sides of a
monomer but are both close to the 220-loop deletion region in
the trimeric form. The Glu at position 186 is close to Arg220 and
may interact with Arg220 when binding avian receptors. Position
205 in the neighboring monomer may be important in trimer
stability and maintaining RBS functionality. If one models the pre-
1996 220-loop restored into the NY107 structure, Arg205, Glu186
and the loop all clash, thus explaining the Glu186Gly mutation
that emerged in the NY107-ins-205 virus HA after limited egg
passage.
The NY107 RBS with its more restricted a2-3 glycan binding
preference and weak/moderate increase in a2-6 binding may have
given the virus a selective advantage to be maintained in poultry at
live bird markets and supplying farms. Certain terrestrial birds,
such as quails and chickens, have recently been shown to present
both human and avian types of receptors in the trachea and
intestine [37,38,39]. Although it is not known what specific glycans
are presented in these animals, it is conceivable that a virus with
mixed specificity might have a distinct advantage over avian
viruses that have specific avian receptor requirements, particularly
in bird markets where multiple species coalesce. Previous results
with H7N2, H9N2 and H5N1 viruses all highlight the fact that an
increase in a2-6-binding preference is not sufficient for efficient
transmission of avian influenza viruses to humans [22,40,41].
Although it remains to be seen whether prolonged circulation of
viruses in terrestrial birds, such as domestic chickens, can provide a
possible route for viruses to adapt for efficient human infection
[11], continued surveillance of influenza viruses from avian and
other animal reservoirs is urgently needed to define their zoonotic
potential.
Materials and Methods
Cloning
Based on H3 numbering [42], cDNA corresponding to residues
11–329 (HA1) and 1–176 (HA2) of the ectodomain of the
hemagglutinin
(HA)
from
A/New
York/107/2003
(H7N2;
Genbank:ACC55270)
and
A/Netherlands/219/2003
(H7N7;
Genebank: AAR02640) was cloned into the baculovirus transfer
vector, pAcGP67-A (BD Biosciences), incorporating a C-terminal
thrombin cleavage site, a ‘‘foldon’’ sequence [43] and a His-tag at
the extreme C-terminus of the construct to enable protein
purification [25,44]. Transfection and virus amplification were
carried out according to the baculovirus expression system manual
(BD Biosciences Pharmingen).
Figure 4. Glycan interactions within the NY107 RBS. The top panel shows the interactions of NY107 with (A) 39SLN, (B) 69SLN and (C) LSTb.
NY107 is shown in orange/red/magenta cartoon respectively. The interacting HA residues are shown as green sticks. The bottom panel shows the
electron density map of the ligands. The NY107 is shown in the same colors as above, and the ligands are shown as green sticks, the 2fo-fc electron
density maps (contoured at 1s) are shown in grey. Simulated annealing omit maps are shown in supplementary Figure S4.
doi:10.1371/journal.ppat.1001081.g004
Structure of a North American H7 Hemagglutinin
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Protein expression and purification
Soluble NY107 was recovered from the cell supernatant by metal
affinity chromatography using Ni-NTA resin (Qiagen Inc.). Frac-
tions containing NY107 were pooled and dialyzed against 10 mM
Tris-HCl, 50 mM NaCl, pH 8.0, then subjected to ion-exchange
chromatography (IEX) using a Mono-Q HR 10/10 column (GE
Healthcare). IEX purified NY107 was subjected to thrombin digest
(3 units/mg protein; overnight at 4uC) and purified by gel filtra-
tion chromatography using a Superdex-200 16/60 column (GE
Healthcare) and 50 mM Tris-HCl, 100 mM NaCl, pH 8.0 as
running buffer. Protein eluting as a trimer was buffer exchanged
into 10 mM Tris-HCl, 50 mM NaCl, pH 8.0 and concentrated to
14.5 mg/ml for crystallization trials. At this stage, the protein
sample still contained the additional plasmid-encoded residues at
both the N (ADPG) and C terminus (SGRLVPR).
Crystallization, ligand soaking and data collection
Initial crystallization trials were set up using a Topaz Free
Interface Diffusion (FID) Crystallizer system (Fluidigm Corpora-
tion, San Francisco, CA). Crystals were observed in several
conditions containing PEG 3350 or PEG 4000. Following opti-
mization, diffraction quality crystals for NY107 were obtained at
room temperature using a modified method for microbath under
oil [45], by mixing the protein with reservoir solution containing
20% PEG 3350, 0.2 M magnesium chloride at pH 7.2. For
receptor analog complexes, crystals were soaked for 3 hours in the
crystallization buffer containing 10 mM 39SLN or 69SLN (V-labs
Inc., Covington, LA), or overnight in 10mM LSTb (Sigma, St.
Louis, MO). All crystals were flash-cooled at 100K using 20%
glycerol as the cryo-protectant. Datasets were collected at
Advanced Photon Source (APS) beamlines 22 ID and BM at
100K. Data were processed with the DENZO-SACLEPACK suite
[46]. Statistics for data collection are presented in Table 1.
Structure determination and refinement
The
structure
of
NY107
was
determined
by
molecular
replacement with Phaser [47] using the structure of the avian
H7 (Av-H7) from A/turkey/Italy/2002, pdb:1TI8 (HA1, 78%
identity; HA2, 90% identity) as the searching model. One HA
trimer occupies the asymmetric unit with an estimated solvent
content of 58% based on a Matthews’ coefficient (Vm) of 2.9 A˚ 3/
Da. Rigid body refinement of the trimer led to an overall R/Rfree
of 28.6%/37.4%. The model was then ‘‘mutated’’ to the correct
sequence and rebuilt by Coot [48], then the protein structures
were refined with REFMAC [49] using TLS refinement [50]. The
final models were assessed using MolProbity [51]. The three
complex structures were refined and evaluated using the same
strategy. All statistics for data processing and refinement are
presented in Table 1. Electron density maps (2fo-fc) were
generated in Refmac [49] while simulated annealing omit maps
were generated by sa-omit-map, a part of the Crystallography and
NMR System (CNS) software [52].
Virus generation
Wild type and mutant viruses of NY107 (H7N2) and A/
Netherland/219/2003 (H7N7) were generated from plasmids by a
reverse genetics approach [53]. To generate viruses with amino
acid insertion or substitution in the HA, mutations were
introduced into plasmid DNA with an overlap extension PCR
approach [54]. Viruses derived by plasmid transfection of HK293
cells were propagated in eggs. The genomes of resulting virus
stocks were sequenced to detect the emergence of possible variants
during amplification.
Glycan binding analyses
Glycan microarray printing and recHA analyses have been
described previously [24,30,44,55] (see Table 2 for glycans used
for analyses in these experiments). Virus were analyzed on the
microarray as described previously [30].
PDB accession codes
The atomic coordinates and structure factors of NY107 are
available from the RCSB PDB under accession codes 3M5G for
the unliganded NY107, 3M5H for the NY107 with 39-SLN and
3M5I and 3M5J for NY107 with 69SLN and LSTb, respectively.
Accession/ID numbers for genes/proteins used in this
work
A/New York/107/03 (H7N2), Genbank: ACC55270; A/
Netherlands/219/03 (H7N7), Genbank: AAR02640; A/Hong
Kong/1-9/68 (H3N2), 2HMG; A/Duck/Ukraine/1/63 (H3N8),
PDB: 1MQL; A/South Carolina/1/18 (H1N1), PDB: 1RD8; A/
Puerto Rico/8/34 (H1N1), PDB: 1RU7; A/Swine/Iowa/15/30
(H1N1), PDB: 1RUY; A/Singapore/1/1957
(H2N2), PDB:
2WRC; A/Viet Nam/1203/04 (H5N1), PDB: 2FK0; A/Duck/
Singapore/3/97 (H5N3), PDB: 1JSM; A/Swine/Hong Kong/9/
98 (H9N2), PDB: 1JSD; A/Turkey/Italy/8000/02 (H7N3), PDB:
1TI8; C/Johannesburg/1/66, 1FLC.
Supporting Information
Figure S1
Sequence alignment of selected structurally available
HAs. Human H3 (PDB: 2HMG), Avian H3 (PDB: 1MQL), 1918-
Human H1 (PDB: 1RD8), 1934-Human H1 (PDB: 1RU7), Swine
H1 (PDB: 1RUY), 1957-Huamn H2 (PDB: 2WRC), Human H5
(PDB: 2FK0), Avian H5 (PDB: 1JSM), Swine H9 (PDB: 1JSD),
and Avian H7 (PDB: 1TI8) were used in the alignments. The
fusion domain of HA1 is highlighted in magenta, the vestigial
esterase domain is highlighted in green, the receptor binding
domain is highlighted in blue, and the fusion domain of HA2 is
highlighted in red. Residue numbering is based on the H3 HA
sequence.
Found at: doi:10.1371/journal.ppat.1001081.s001 (2.84 MB TIF)
Figure S2
Expression and purification of NY107. SDS-PAGE
reveals that NY107 was expressed as the HA0 form with a mass
approximately 60kDa (middle lane). Thrombin cleavage resulted in
an unexpected reduction in band size to a HA1/HA2 profile (right
lane) with possible multiple glycoforms for the HA2 clearly present.
Found at: doi:10.1371/journal.ppat.1001081.s002 (0.23 MB TIF)
Figure S3
Comparison of glycan binding to NY107 with other
HAs. A. Overlap of a2-3 ligands binding in the receptor binding
site from NY-H7 (green), Av-H3 (orange), 1930-Hu-H1 (magenta),
1957-Hu-H2 (cyan), Av-H5 (yellow), and Sw-H9 (grey). B.
Overlap of a2-6 linkage ligands binding in the receptor binding
site from NY-H7 (green), Av-H3 (orange), 1930-Hu-H1 (magenta),
1957-Hu-H2 (cyan), Av-H5 (yellow), and Sw-H9 (grey).
Found at: doi:10.1371/journal.ppat.1001081.s003 (2.55 MB TIF)
Figure S4
Simulated annealing omit maps of the receptor
binding site (contoured at 1s). A. NY107 (blue), B. NY107-39SLN
(orange), C. NY107-69SLN (red), and D. NY107-LSTb (magenta).
The protein model is shown in cartoon, and the residues involved
in the binding to receptor analogs were shown in sticks. Maps were
generated using version 1.2 of the Crystallography and NMR
System (CNS) software.
Found
at:
doi:10.1371/journal.ppat.1001081.s004
(1.93
MB
TIF)
Structure of a North American H7 Hemagglutinin
PLoS Pathogens | www.plospathogens.org
9
September 2010 | Volume 6 | Issue 9 | e1001081
Figure S5
Comparison of NY107 RBS to HEF. Overlap of RBS
from NY107 (green), Av-H7 (marine) and HEF (magenta).
Found at: doi:10.1371/journal.ppat.1001081.s005 (1.12 MB TIF)
Table S1
Comparison of r.m.s.d. (A˚ ) for different HA domains.
For analyzing differences in the overall structure, r.m.s.d. values
were calculated between monomers or domains of different HA’s,
after the Ca atoms of the HA2 domains were superposed by
sequence and structural alignment onto the equivalent domains of
NY107.
Found
at:
doi:10.1371/journal.ppat.1001081.s006
(0.04
MB
DOC)
Table S2
Comparison of r.m.s.d. (A˚ ) for individual domains.
Each domain was superimposed separately to determine how the
individual NY107 domains compared to equivalent domains in the
other structures.
Found
at:
doi:10.1371/journal.ppat.1001081.s007
(0.04
MB
DOC)
Table S3
Molecular interactions between NY107 and receptor
analogs. The hydrogen bond cutoff is 3.8 A˚
for the listing
interactions.
Found
at:
doi:10.1371/journal.ppat.1001081.s008
(0.07
MB
DOC)
Table S4
Glycan array differences between NY107, the fully
restored NY107-ins, and NL219 (virus and rHA). The color
coding in the left hand column reflects the same coloring scheme
used in Figures 2 and 3. Significant binding of samples to glycans
are qualitatively estimated based on relative strength of the signal
for the data shown in Figures 2 and 3 Strong (+++), weak (+).
Found
at:
doi:10.1371/journal.ppat.1001081.s009
(0.19
MB
DOC)
Acknowledgments
The authors would like to thank the staff of SER-CAT sector 22 for their
help in data collection. We also thank WHO Global Influenza Surveillance
Network for providing NY107 and NL219 viruses from which the reverse
genetics viruses were generated. Glycan microarray data presented here
will be made available on-line through the CFG web site upon publication
(www.functionalglycomics.org). The findings and conclusions in this report
are those of the authors and do not necessarily represent the views of the
Centers for Disease Control and Prevention or the Agency for Toxic
Substances and Disease Registry.
Author Contributions
Conceived and designed the experiments: HY LMC PJC ROD JS.
Performed the experiments: HY LMC PJC JS. Analyzed the data: HY
LMC PJC ROD JS. Wrote the paper: HY LMC PJC ROD JS.
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3M5L
|
Crystal structure of HCV NS3/4A protease in complex with ITMN-191
|
Drug resistance against HCV NS3/4A inhibitors is
defined by the balance of substrate recognition
versus inhibitor binding
Keith P. Romano1, Akbar Ali1, William E. Royer, and Celia A. Schiffer2
Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, MA 01605
Edited by John M. Coffin, Tufts University School of Medicine, Boston, MA, and approved September 14, 2010 (received for review May 13, 2010)
Hepatitis C virus infects an estimated 180 million people world-
wide, prompting enormous efforts to develop inhibitors targeting
the essential NS3/4A protease. Resistance against the most promis-
ing protease inhibitors, telaprevir, boceprevir, and ITMN-191, has
emerged in clinical trials. In this study, crystal structures of the
NS3/4A protease domain reveal that viral substrates bind to the
protease active site in a conserved manner defining a consensus
volume, or substrate envelope. Mutations that confer the most
severe resistance in the clinic occur where the inhibitors protrude
from the substrate envelope, as these changes selectively weaken
inhibitor binding without compromising the binding of substrates.
These findings suggest a general model for predicting the suscept-
ibility of protease inhibitors to resistance: drugs designed to fit
within the substrate envelope will be less susceptible to resistance,
as mutations affecting inhibitor binding would simultaneously
interfere with the recognition of viral substrates.
drug design ∣hepatitis C ∣substrate envelope
D
rug resistance is a major obstacle in the treatment of quickly
evolving diseases. Hepatitis C virus (HCV) is a genetically
diverse Hepacivirus of the Flaviviridae family infecting an esti-
mated 180 million people worldwide (1). The viral RNA genome
is translated as a single polyprotein and subsequently processed
by host-cell and viral proteases into structural (C, E1, E2, and p7)
and nonstructural (NS2, NS3, NS4A, NS4B, NS5A, and NS5B)
proteins (2). The viral RNA-dependent RNA polymerase, NS5B,
is inherently inaccurate and misincorporation of bases accounts
for a very high mutation rate (3). While some mutations are neu-
tral, others will alter the viability of the virus and propagate with
varying efficiencies in each patient. Thus HCV infected indivi-
duals will develop a heterogeneous population of virus variants
known as quasispecies (4). As patients begin treatment, the selec-
tive pressures of antiviral drugs will favor drug resistant variants
(5). Therefore, an inhibitor must not only recognize one protein
variant, but an ensemble of related enzymes. A detailed under-
standing of the atomic mechanisms of resistance is essential to
effectively combat drug resistance against HCV antivirals.
The essential HCV NS3/4A protease is an attractive therapeu-
tic target responsible for cleaving at least four sites along the viral
polyprotein. These sites share little sequence homology except
for an acid at position P6, Cys or Thr at P1, and Ser or Ala at
P1′ (Table S1). The first cleavage event at the 3-4A junction
occurs in cis as a unimolecular process, while the remaining sub-
strates are processed bimolecularly in trans. The NS3/4A protease
also cleaves the human cellular targets TRIF and MAVS, which
confounds the innate immune response to viral infection (6–8).
Early drug design efforts were hampered by the relatively shallow,
featureless architecture of the protease active site. The eventual
observation of N-terminal product inhibition served as a stepping
stone for the discovery of more potent peptidomimetic inhibitors
(9, 10). Over the past decade, pharmaceutical companies have
further developed these lead compounds. Many structure-activ-
ity-relationship (SAR) studies have been performed to evaluate
the effect of different functional moieties on protease inhibition
at positions P4-P1′ (11–17). Crystal structures have been deter-
mined of the NS3/4A protease domain bound to a variety of
inhibitors as well as of several drug resistant protease variants,
such as R155K and V36M (18, 19). These data elucidate the mo-
lecular interactions of NS3/4A with inhibitors and the effect of
specific drug resistance mutations on binding. These efforts, con-
ducted in parallel by several pharmaceutical companies, led to
the discovery of many protease inhibitors. Proof-of-concept for
the successful clinical activity of this drug class was first demon-
strated by the macrocyclic inhibitor BILN-2061 (Boehringer
Ingelheim) (20, 21), which was later dropped from clinical trials
in 2006 due to cardiotoxicity (22). Many other NS3/4A protease
inhibitors are currently in development, and telaprevir (Vertex),
boceprevir (Schering-Plough), and ITMN-191 (Intermune) lead
the way in advanced phases of human clinical trials (Fig. 1A).
Despite these successes, the rapid acquisition of drug resis-
tance has limited the efficacy of the most potent NS3/4A protease
inhibitors in both replicon studies and human clinical trials
(Fig. 1B and Table 1). In this study, we show that mutations con-
ferring the most severe resistance occur where the protease
extensively contacts the inhibitors but not the natural viral sub-
strates. Four crystal structures of the NS3/4A protease domain in
complex with the N-terminal products of viral substrates reveal a
conserved mode of substrate binding, with the consensus volume
defining the substrate envelope. The protease inhibitors ITMN-
191 (3M5L), TMC435 (3KEE) (23), and boceprevir (2OC8) (24)
protrude extensively from the substrate envelope in regions that
correlate with known sites of resistance mutations. Most notably,
the P2 moieties of all three drugs protrude to contact A156 and
R155, which mutate to confer high-level resistance against nearly
all drugs reported in the literature (25–30). These findings sug-
gest that drug resistance results from a change in molecular
recognition and imply that drugs designed to fit within the sub-
strate envelope will be less susceptible to resistance, as mutations
altering inhibitor binding will simultaneously interfere with the
binding of substrates.
Results
Synthesis of ITMN-191. We synthesized the macrocyclic inhibitor
ITMN-191 using a convergent reaction sequence described in
SI Text. Briefly, the P2 and P1-P1′ fragments were preassembled
Author contributions: K.P.R., A.A., and C.A.S. designed research; K.P.R. and A.A. performed
research; A.A. and W.E.R. contributed new reagents/analytic tools; K.P.R., W.E.R., and
C.A.S. analyzed data; and K.P.R., A.A., W.E.R., and C.A.S. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The atomic coordinates and structure factors have been deposited in the
Protein Data Bank, www.pdb.org (PDB ID codes 3M5L, 3M5M, 3M5N, and 3M5O).
1K.P.R. and A.A. contributed equally to this work.
2To whom correspondence should be addressed. E-mail: Celia.Schiffer@umassmed.edu.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1006370107/-/DCSupplemental.
20986–20991 ∣PNAS ∣December 7, 2010 ∣vol. 107 ∣no. 49
www.pnas.org/cgi/doi/10.1073/pnas.1006370107
and the macrocyclic drug compound was generated by a four-
step reaction sequence, including P2-P3 amide coupling, ester
hydrolysis, coupling with the P1-P1′ fragment, and ring-closing
metathesis. The P2-P3 fragment was assembled by coupling the
commercially available Boc-protected amino acid (S)-2-(tert-
butoxycarbonylamino)non-8-enoic acid (Acme Biosciences, Inc)
with the preassembled P2 fragment, (3R, 5S)-5-(methoxy-
carbonyl)pyrrolidin-3-yl 4-fluoroisoindoline-2-carboxylate (31),
using O-(7-azabenzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexa-
fluorophosphate (HATU)/diisopropylethylamine (DIPEA). Hy-
drolysis of the P2-P3 methyl ester with LiOH:H2O in a mixture
of THF-MeOH-H2O followed by coupling of the resulting acid
under HATU/DIPEA conditions with the preassembled P1-P1′
fragment, (1R, 2S)-1-amino-N-(cyclopropylsulfonyl)-2-vinylcyclo-
propanecarboxamide (32), provided the bis-olefin precursor for
ring-closing metathesis. Cyclization of the bis-olefin intermediate
was accomplished using a highly efficient ring-closing metathesis
catalyst Zhan 1B and provided the protease inhibitor ITMN-191.
Structure Determination of Inhibitor and Substrate Complexes.
Although NS3/4A cleaves the viral polyprotein of over 3,000
residues at four specific sites in vivo, we focused on the local
interactions of the protease domain with short peptide sequences
corresponding to the immediate cleavage sites. All structural
studies were carried out with the highly soluble, single-chain con-
struct of the NS3/4A protease domain described previously (33),
which contains a fragment of the essential cofactor NS4A cova-
lently linked at the N terminus by a flexible linker. A similar pro-
tease construct was shown to retain comparable catalytic activity
to the authentic protein complex (34). Crystallization trials were
initially carried out using the inactive (S139A) protease variant
in complex with substrate peptides spanning P7-P5′. The 4A4B
substrate complex revealed cleavage of the scissile bond and no
ordered regions for the C-terminal fragment of the substrate.
Similar observations were previously described for two other
serine proteases where catalytic activity was observed, presum-
ably facilitated by water, despite Ala substitutions of the catalytic
Ser (35, 36). Thus all subsequent crystallization trials with the
NS3/4A protease were performed using N-terminal cleavage
products of the viral substrates spanning P7-P1.
NS3/4A crystal structures in complex with ITMN-191 and
peptide products 4A4B, 4B5A, and 5A5B were determined and
refined at 1.25 Å, 1.70 Å, 1.90 Å, and 1.60 Å resolution, respec-
tively (Table S2). The complexes crystallized in the space groups
P212121 and P21 with one, two, or four molecules in the asym-
metric unit. The average B factors range from 16.8–29.7 Å2
and there are no outliers in the Ramachandran plots. These
structures represent the highest resolution crystal structures of
NS3/4A protease reported to date.
Overall Structure Analysis. The NS3/4A protease domain adopts a
tertiary fold characteristic of serine proteases of the chymotrypsin
family (37, 38). A total of nine protease molecules were modeled
in the four crystal structures solved in this study with an overall
rms deviation (rmsd) of 0.28 Å. The rmsds reveal the five most
variable regions of the protease to be (Fig. S1): (i) the linker con-
necting cofactor 4A at the N terminus, (ii) the loop containing
residues 65–70, (iii) the zinc-binding site containing residues
95–105, (iv) the 310 helix region spanning residues 128–136,
and (v) the active site antiparallel β-sheet containing residues
156–168. These structural differences likely indicate inherent
flexibility in the protease and do not appear to correlate with
ligand type or active site occupancy.
Analysis of Product Complexes. Product complexes 4A4B, 4B5A,
and 5A5B were further analyzed with the C terminus of the
full-length NS3/4A structure (1CU1), which contains the N-term-
inal cleavage product of viral substrate 3-4A (39). All four
products bind to the protease active site in a conserved manner
(Fig. 2), forming an antiparallel β-sheet with residues 154–160
Fig. 1.
NS3/4A protease inhibitors and reported sites of drug resistance.
(A) The leading protease inhibitors in development mimic the N-terminal side
of the viral substrates. (B) The majority of reported drug resistance mutations
cluster around the protease active site with the catalytic triad depicted in
yellow.
Table 1. Drug resistance mutations reported in replicon studies and
clinical trials*
Residue
Mutation
Drug
V36
A, M, L, G
Boceprevir, telaprevir
Q41
R
Boceprevir, ITMN-191
F43
S, C, V, I
Boceprevir, telaprevir, ITMN-191,
TMC435
V55
A
Boceprevir
T54
A, S
Boceprevir, telaprevir
Q80
K, R, H, G, L
TMC435
S138
T
ITMN-191, TMC435†
R155
K, T, I, M, G, L, S, Q
Boceprevir, telaprevir, ITMN-191,
BILN-2061, TMC435
A156
V, T, S, I, G
Boceprevir, telaprevir, ITMN-191,
BILN-2061, TMC435
V158
I
Boceprevir
D168
A, V, E, G, N, T, Y, H, I
ITMN-191, BILN-2061, TMC435
V170
A
Boceprevir, telaprevir
M175
L
Boceprevir
*References (18, 25, 26, 28, 30–37).
†TMC435 displays reduced activity against S138T, but the mutation was not
observed in selection experiments.
Romano et al.
PNAS
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vol. 107
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20987
BIOPHYSICS AND
COMPUTATIONAL BIOLOGY
CHEMISTRY
and burying 500–600 Å2 of solvent accessible surface area as cal-
culated by PISA (40). The peptide backbone torsions are very
similar, being most conserved at position P1 and deviating slightly
toward position P4. Eight hydrogen bonds between backbone
amide and carbonyl groups are completely conserved, involving
protease residues S159 (C159 in product 3-4A), A157, R155,
S139A, S138, and G137. S159 (C159 in product 3-4A), and
A157 each contribute two hydrogen bonds with the P5 and P3
peptide residues, respectively. All P1 terminal carboxyl groups
sit in the oxyanion hole, hydrogen bonding with the Nϵ atom
of H57 and the amide nitrogens of residues 137–139. Although
only product 4B5A contains a proline at P2, the other substrate
sequences still adopt constrained P2 φ torsion angles. Thus
products bind similarly despite their high sequence diversity.
The P1 and P6 residues are most conserved among the
substrate sequences, as are most of their interactions with the
protease. The P1 side chains interact with the aromatic ring of
F154. In all structures but product complex 4B5A, K165 forms
salt-bridges with the P6 acids, while residues R123, D168,
R155, and the catalytic D81 form an ionic network along one
surface of the bound products (Fig. 2). In complex 4B5A,
R123 interacts directly with the P6 acid, while D168 reorients
and no longer contacts R155. Other molecular interactions in
the product complexes are more diverse. Notably, K136 interacts
differently with the cleavage products, forming: (i) a hydrogen
bond with the P2 backbone carbonyl oxygen of 3-4A; (ii) a
salt-bridge with the P3 glutamate of product 4A4B; and (iii) non-
specific van der Waals interactions with the P2 and P3 side chains
of products 4B5A and 5A5B. Also, in product complex 4A4B, an
intramolecular hydrogen bond forms between the P3 and P5 glu-
tamate residues, while the unique P4 acid of product 3-4A forms
salt-bridges with the guanidinium groups of R123 and R155. Thus
distinct patterns of side chain interactions underlie the set of con-
served features involved in NS3/4A cleavage product binding.
The Substrate Envelope. To further analyze the structural similari-
ties of the four NS3/4A product complexes, the active sites were
superposed on the Cα atoms of residues 137–139 and 154–160,
revealing that both the active site residues and substrate products
spanning P6-P1 align closely with an average Cα rmsd of 0.24 Å
and 0.35 Å, respectively. The consensus van der Waals volume
shared by any three of the four cleavage products was then cal-
culated to generate the NS3/4A substrate envelope (Fig. 3A).
This shape could not be predicted by the primary sequences alone
and highlights the conserved mode of viral substrate recognition
despite their high sequence diversity.
Analysis of Inhibitor Complexes. ITMN-191, TMC435, and bocepre-
vir are all peptidomimetic NS3/4A protease inhibitors. Active site
superpositions of these drug complexes reveal that the inhibitors
interact with many of the same protease residues as the cleavage
products. Despite the P3-P1 cyclization of ITMN-191 and
TMC435, the functional groups are positioned similarly in all
three inhibitor complexes. The P1 cysteine surrogates interact
with the aromatic ring of F154, while the P2 and P3 moieties over-
lap closely. Although TMC435 does not contain a P4 substituent,
the P4 tert-butyl groups of ITMN-191 and boceprevir also align
closely. In addition, the P1 and P3 backbone atoms of all inhibi-
tors hydrogen bond with the carbonyl oxygens of R155 and A157,
respectively. These observations verify the peptidomimetic nat-
ure of these drugs and support their observed mechanism as
competitive active site inhibitors.
The largest variation between these three protease inhibitors
occurs at P2 where the aromatic rings of ITMN-191 and TMC435
stack against the guanidinium group of R155 (Fig. 3). This
molecular interaction alters the electrostatic network involving
R123, D168, R155, and D81. R155 rotates nearly 180° around
Cδ relative to its conformation observed in product complexes,
losing its hydrogen bond with D81 but maintaining interaction
with D168. Mutations at R155 or D168 would disrupt the elec-
trostatic network and destabilize this packing thereby lowering
the affinity of these macrocyclic drugs. This observation provides
a structural rationale for the drug resistance mutations R155K,
as previously proposed (19), and D168A/V, which both confer
a selective advantage in vitro in the presence of ITMN-191 or
TMC435 (26, 30). In addition, the TMC435 complex reveals
that R155 is stabilized by a hydrogen bond with Q80, which also
mutates to confer resistance to TMC435 (30). Thus many of the
primary drug resistance mutations can be explained by the disrup-
tion of atomic interactions involving the P2 functional groups of
the drugs.
Insights into Drug Resistance. To determine the locations where the
inhibitors protrude from the substrate envelope, the inhibitor and
product complexes were also superposed using residues 137–139
and 154–160. The van der Waals volumes of inhibitor protrusion
from the substrate envelope (V out) (41, 42) were calculated for
each drug and compared with published EC50 fold-change data
for drug resistance variants (30). The magnitudes of the EC50
fold-change data determined for each NS3/4A mutant generally
trend with the V out values for the three drugs. The P2 moieties of
boceprevir, ITMN-191 and TMC435 protrude most extensively
from the substrate envelope with V out values of 105, 294, and
Fig. 2.
Stereo view of N-terminal cleavage product binding to NS3/4A pro-
tease. N-terminal protease cleavage products (A) 3-4A, (B) 4A4B, (C) 4B5A,
and (D) 5A5B are depicted as they bind to the protease active site. All
conserved interactions are indicated by black dashes, while red lines depict
interactions that are not present in all structures. The electrostatic network
involving residues R123, D168, R155, and D81 is indicated by blue dashes.
20988
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www.pnas.org/cgi/doi/10.1073/pnas.1006370107
Romano et al.
496 Å3, respectively (Table 2). However, the precise level of drug
resistance observed is also determined by the particular change
in molecular interaction occurring for a given mutation. For ex-
ample, A156 and R155 pack with the P2 moieties of these three
inhibitors where they protrude beyond the substrate envelope.
Mutations of A156 to bulkier side chains would result in a steric
clash with the P2 drug moieties. Indeed, the rigid dimethylcyclo-
propane group of boceprevir protrudes from the substrate envel-
ope at the P2 subsite, and A156V or A156T confer 65 and 75
fold-changes in EC50, respectively (Table 2). Similarly, molecular
changes at R155 and D168 would result in a substantial loss of
interactions with P2. The most extensive protrusions of ITMN-
191 and TMC435 at P2 trend with their greatest fold-change
in potency of nearly 450 and 600, respectively, from mutations
in this subsite. Thus the extent by which an inhibitor protrudes
from the substrate envelope in a given subsite is indicative of
its vulnerability to resistance.
Further structural analyses with the substrate envelope provide
insights into other NS3/4A drug resistance mutations. The P1′
sulfonamide groups of ITMN-191 and TMC435, as well as the
P1′ ketoamide of boceprevir, protrude from the substrate envel-
ope near residues Q41 and F43, which both mutate to confer
low-level resistance to these drugs (25, 30, 43). The keto group
of boceprevir also projects outside the substrate envelope near
T54 and V55. T54A/S confers low-level resistance to boceprevir,
while V55A was recently identified in patient isolates after treat-
ment with boceprevir (44). The analogous carbonyl groups of
ITMN-191 and TMC435, however, are orientated in the opposite
direction and protrude toward S138. In fact, in vitro studies reveal
reduced activity for ITMN-191 and TMC435 against S138T
variants, while boceprevir remains fully active (30, 43). The bulky
P4 tert-butyl group of boceprevir extends outside the substrate
envelope contacting V158; the V158I variant has lower affinity
for this drug, likely due to a steric clash (45). This variant may
also impact the affinity of ITMN-191, as its P4 tert-butyl also pro-
trudes at the same location. These findings demonstrate that in
regions outside the P2 subsite, positions where ITMN-191,
TMC435, and boceprevir protrude from the substrate envelope
also correlate with many other known sites of drug resistance
mutations.
Discussion
The emergence of drug resistance is a major obstacle in modern
medicine that limits the long-term usefulness of the most promis-
ing therapeutics. By considering how HCV NS3/4A protease in-
hibitors bind relative to natural viral substrates, we discovered
that primary sites of resistance occur in regions of the protease
where drugs protrude from the substrate envelope. In particular,
R155 and A156, which mutate to confer severe resistance against
ITMN-191, TMC435, and boceprevir, interact closely with the P2
drug moieties where they protrude most extensively from the
substrate envelope. Molecular changes at these residues confer
resistance by selectively weakening inhibitor binding without
compromising the binding of viral substrates. We further specu-
late that these mutations will not considerably affect the binding
of the host cellular substrates TRIF and MAVS, which likely fit
well within the substrate envelope as they share many features
with the viral substrates. However, TRIF contains a track of
eight proline residues instead of an acidic residue at position
P6, which may modulate its binding. Further structural studies
are warranted to better ascertain the molecular details of how
these cellular substrates are recognized by the NS3/4A protease.
Although this study focuses on ITMN-191, TMC435, and
boceprevir, other NS3/4A protease inhibitors in clinical trials, in-
cluding telaprevir, narlaprevir, and vaniprevir (Fig. 1A), contain
similar functional groups that likely protrude from the substrate
envelope. Most notably, all these drug candidates contain bulky
P2 moieties and are therefore susceptible to cross-resistance
against mutations at R155 and A156. R155 and A156 mutations
have been shown to confer telaprevir resistance in treated
patients (46). Cross-resistance studies have also shown that nar-
Table 2. EC50 fold-change (FC) data * for several NS3/4A drug
resistant variants tabulated with Vout, the van der Waals volume of
protrusion from the substrate envelope, at each subsite of the
enzyme
Boceprevir
ITMN-191
TMC435
Subsite
Resistance
mutation
Vout
(Å3)
EC50
FC
Vout
(Å3)
EC50
FC
Vout
(Å3)
EC50
FC
Total
292
500
649
P1
76
67
64
P2
105
294
496
Q80R
0.5
3.5
6.9
Q80K
0.8
2.3
7.7
R155K
4.7
447
30
A156V
75
63
177
A156T
65
41
44
D168A
0.7
153
594
D168E
0.8
75
40
P3
34
70
67
P4
76
69
0
V158I
3.3†
ND
ND
*Antiviral activity was reported previously by Lenz et al., 2010 (30).
†Fold-change in EC50 reported in replicon assay by Qiu et al., 2009 (45).
Fig. 3.
Stereo view of the NS3/4A substrate envelope and protease
inhibitors. (A) After active site superpositions, the overlapping van der Waals
volume shared by any three of the four cleavage products defines the
substrate envelope, depicted in blue. NS3/4A protease residues which mutate
to confer drug resistance are shown in brown. (B) ITMN-191, (C) boceprevir
and (D) TMC435 protrude from the substrate envelope at several locations,
which correlate with known sites of drug-resistant mutations to each inhibi-
tor, shown in red.
Romano et al.
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laprevir displays similar fold losses in activity against most of the
known drug resistance mutations for telaprevir and boceprevir
(47). Ultimately, to slow the emergence of multidrug resistant
viral strains, inhibitors should be confined within the substrate
envelope, particularly at the P2 position. To compensate for the
loss of binding affinity that will likely accompany these changes,
additional interactions could potentially be optimized spanning
the S4-S6 subsites of the protease and the catalytic triad.
Our findings further suggest a general model for using the sub-
strate envelope to predict patterns of drug resistance in other
quickly evolving diseases. For drug resistance to occur, mutations
must selectively weaken a target’s affinity for an inhibitor without
significantly altering its natural biological function. Mutations
occurring outside the substrate envelope are better able to
achieve this effect, as these molecular changes can selectively
alter inhibitor binding without compromising the binding of
natural substrates. Whenever the interaction of a drug target with
its biological substrates can be structurally characterized, we pre-
dict that drugs designed to fit within the substrate envelope will
be less susceptible to resistance. Structure-based design strategies
can utilize this model as an added constraint to develop inhibitors
that fit within the substrate envelope. In fact, previous work in
our laboratory provides proof-of-concept for the successful incor-
poration of the substrate envelope in the design of unique HIV
protease inhibitors, which maintain high affinities against a panel
of multidrug resistant variants of HIV-1 protease (42, 48–53). As
a general paradigm, design efforts incorporating the substrate
envelope would facilitate a more rationale evaluation of drug
candidates and lead to the development of more robust inhibitors
that are less susceptible to resistance.
Materials and Methods
Protein Crystallization. The NS3/4A protease construct was expressed and
purified as reported previously (33, 54), detailed in SI Text. Purified protein
was concentrated to ∼3 mg∕mL and loaded on a HiLoad Superdex75 16/60
column equilibrated with 25 mM 2-(N-morpholino)ethanesulfonic acid (MES)
at pH 6.5, 500 mM NaCl, 10% glycerol, 30 μM zinc chloride, and 2 mM DTT.
The protease fractions were pooled and concentrated to 20–25 mg∕mL with
an Amicon Ultra-15 device (10 kD; Millipore). The concentrated samples were
then incubated at 4 °C for 1 h with 2–20 M excess of viral substrate 4A4B,
peptide products 4B5A or 5A5B, or ITMN-191. Information about the synth-
esis of viral peptides and ITMN-191 is provided in SI Text. Diffraction-quality
crystals were obtained overnight for all ligands by mixing equal volume of
concentrated protein solution with precipitant solution (20–26% PEG-3350,
0.1 M sodium MES buffer at pH 6.5, and 4% ammonium sulfate) in 24-well
VDX hanging drop trays.
Data Collection and Structure Solution. Crystals were flash-frozen in liquid
nitrogen and mounted under constant cryostream. X-ray diffraction data
were collected at Advanced Photon Source BioCARS 14-IDB, 14-BMC, and
LS-CAT 21-ID-F. Diffraction intensities of the complexes were indexed,
integrated, and scaled using the programs HKL2000 (55) and XDS (56). 5%
of the data was used to calculate R-free (57). All structure solutions were
generated using isomorphous molecular replacement with PHASER (58) or
AMORE. The NS3/4A protease domain (PDB code 2A4G) (59) was used for
molecular replacement in solving the product 4A4B structure, and this struc-
ture was subsequently used for solving the other complexes. In all cases,
initial refinement was carried out in the absence of modeled ligand, which
was subsequently built in. Phases were improved using ARP/wARP (60). Itera-
tive rounds of translation, libration, and screw (TLS) and restrained refine-
ment with CCP4 (61) and graphical model building with COOT (62) until
convergence was achieved. The final structures were evaluated with
MolProbity (63) prior to deposition in the protein data bank.
Structural Analysis. Double-difference plots (64) were used to determine the
structurally invariant regions of the protease, consisting of residues 32–36,
42–47, 50–54, 84–86, and 140–143. Structures were superposed with
PyMOL (65) using the Cα atoms of these residues for all protease molecules
from the solved structures (nine total). The B chain of product complex
4A4B was used as the reference structure in all alignments. Fit of individual
inhibitors into the substrate envelope was quantified by mapping the
substrate envelope and the van der Waals volume of each inhibitor on a
three-dimensional grid with spacing of 0.5 Å. Vout for each drug moiety
was computed by counting the grid cells, which were occupied by any inhi-
bitor atom of that site but not the substrate envelope, and multiplied by
the grid cell size, 0.125 Å3 (41, 42).
Substrate Envelope and Inhibitor Analyses. NS3/4A substrate envelope was
computed using product complexes 4A4B (B chain), 4B5A (D chain), and
5A5B (A chain). In structures with multiple protease molecules in the asym-
metric unit, the one containing the most ordered peptide product was used
for the alignment. The protease domain of the full-length NS3/4A structure
(A chain; PDB code 1CU1) (39), including the C-terminal six amino acids,
was included as a product complex 3-4A. All active site alignments were
performed in PyMOL using Cα atoms of protease residues 137–139 and
154–160. After superposition, Gaussian object maps were generated in Py-
MOL for each cleavage product. Four consensus Gaussian maps were then
calculated, representing the intersecting volume of a group of three object
maps. Finally, the summation of these four consensus maps was generated to
construct the substrate envelope, depicting the van der Waals volume shared
by any three of the four products. The previously determined boceprevir
complex (PDB code 2OC8) (24) and TMC435 complex (PDB code 3KEE) (23)
were used in this study (66).
ACKNOWLEDGMENTS. We thank H. Klei for helpful discussions. We also thank
Z. Wawrzak, M. Bolbat, and K. Brister of the LS-CAT beamline at Argonne
National Laboratory for data collection of the ITMN-191 complex; M. Nalam
and R. Bandaranayake for assistance with structural refinement; A. Ozen for
providing V out calculations; and S. Shandilya and Y. Cai for computational
support. The National Institute of Health (NIH) Grants R01-GM65347 and
R01-AI085051 supported this work. Use of Advanced Photon Source (APS)
was supported by the Department of Energy (DOE), Basic Energy Sciences,
Office of Science, under Contract No. DE-AC02-06CH11357. Use of the Bio-
CARS Sector 14 was supported by NIH-NCRR RR007707. Use of the LS-CAT
Sector 21 was supported by the Michigan Economic Development Corpora-
tion and the Michigan Technology TriCorridor under Grant 085P1000817.
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3M5M
|
Avoiding drug resistance against HCV NS3/4A protease inhibitors
|
Drug resistance against HCV NS3/4A inhibitors is
defined by the balance of substrate recognition
versus inhibitor binding
Keith P. Romano1, Akbar Ali1, William E. Royer, and Celia A. Schiffer2
Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, MA 01605
Edited by John M. Coffin, Tufts University School of Medicine, Boston, MA, and approved September 14, 2010 (received for review May 13, 2010)
Hepatitis C virus infects an estimated 180 million people world-
wide, prompting enormous efforts to develop inhibitors targeting
the essential NS3/4A protease. Resistance against the most promis-
ing protease inhibitors, telaprevir, boceprevir, and ITMN-191, has
emerged in clinical trials. In this study, crystal structures of the
NS3/4A protease domain reveal that viral substrates bind to the
protease active site in a conserved manner defining a consensus
volume, or substrate envelope. Mutations that confer the most
severe resistance in the clinic occur where the inhibitors protrude
from the substrate envelope, as these changes selectively weaken
inhibitor binding without compromising the binding of substrates.
These findings suggest a general model for predicting the suscept-
ibility of protease inhibitors to resistance: drugs designed to fit
within the substrate envelope will be less susceptible to resistance,
as mutations affecting inhibitor binding would simultaneously
interfere with the recognition of viral substrates.
drug design ∣hepatitis C ∣substrate envelope
D
rug resistance is a major obstacle in the treatment of quickly
evolving diseases. Hepatitis C virus (HCV) is a genetically
diverse Hepacivirus of the Flaviviridae family infecting an esti-
mated 180 million people worldwide (1). The viral RNA genome
is translated as a single polyprotein and subsequently processed
by host-cell and viral proteases into structural (C, E1, E2, and p7)
and nonstructural (NS2, NS3, NS4A, NS4B, NS5A, and NS5B)
proteins (2). The viral RNA-dependent RNA polymerase, NS5B,
is inherently inaccurate and misincorporation of bases accounts
for a very high mutation rate (3). While some mutations are neu-
tral, others will alter the viability of the virus and propagate with
varying efficiencies in each patient. Thus HCV infected indivi-
duals will develop a heterogeneous population of virus variants
known as quasispecies (4). As patients begin treatment, the selec-
tive pressures of antiviral drugs will favor drug resistant variants
(5). Therefore, an inhibitor must not only recognize one protein
variant, but an ensemble of related enzymes. A detailed under-
standing of the atomic mechanisms of resistance is essential to
effectively combat drug resistance against HCV antivirals.
The essential HCV NS3/4A protease is an attractive therapeu-
tic target responsible for cleaving at least four sites along the viral
polyprotein. These sites share little sequence homology except
for an acid at position P6, Cys or Thr at P1, and Ser or Ala at
P1′ (Table S1). The first cleavage event at the 3-4A junction
occurs in cis as a unimolecular process, while the remaining sub-
strates are processed bimolecularly in trans. The NS3/4A protease
also cleaves the human cellular targets TRIF and MAVS, which
confounds the innate immune response to viral infection (6–8).
Early drug design efforts were hampered by the relatively shallow,
featureless architecture of the protease active site. The eventual
observation of N-terminal product inhibition served as a stepping
stone for the discovery of more potent peptidomimetic inhibitors
(9, 10). Over the past decade, pharmaceutical companies have
further developed these lead compounds. Many structure-activ-
ity-relationship (SAR) studies have been performed to evaluate
the effect of different functional moieties on protease inhibition
at positions P4-P1′ (11–17). Crystal structures have been deter-
mined of the NS3/4A protease domain bound to a variety of
inhibitors as well as of several drug resistant protease variants,
such as R155K and V36M (18, 19). These data elucidate the mo-
lecular interactions of NS3/4A with inhibitors and the effect of
specific drug resistance mutations on binding. These efforts, con-
ducted in parallel by several pharmaceutical companies, led to
the discovery of many protease inhibitors. Proof-of-concept for
the successful clinical activity of this drug class was first demon-
strated by the macrocyclic inhibitor BILN-2061 (Boehringer
Ingelheim) (20, 21), which was later dropped from clinical trials
in 2006 due to cardiotoxicity (22). Many other NS3/4A protease
inhibitors are currently in development, and telaprevir (Vertex),
boceprevir (Schering-Plough), and ITMN-191 (Intermune) lead
the way in advanced phases of human clinical trials (Fig. 1A).
Despite these successes, the rapid acquisition of drug resis-
tance has limited the efficacy of the most potent NS3/4A protease
inhibitors in both replicon studies and human clinical trials
(Fig. 1B and Table 1). In this study, we show that mutations con-
ferring the most severe resistance occur where the protease
extensively contacts the inhibitors but not the natural viral sub-
strates. Four crystal structures of the NS3/4A protease domain in
complex with the N-terminal products of viral substrates reveal a
conserved mode of substrate binding, with the consensus volume
defining the substrate envelope. The protease inhibitors ITMN-
191 (3M5L), TMC435 (3KEE) (23), and boceprevir (2OC8) (24)
protrude extensively from the substrate envelope in regions that
correlate with known sites of resistance mutations. Most notably,
the P2 moieties of all three drugs protrude to contact A156 and
R155, which mutate to confer high-level resistance against nearly
all drugs reported in the literature (25–30). These findings sug-
gest that drug resistance results from a change in molecular
recognition and imply that drugs designed to fit within the sub-
strate envelope will be less susceptible to resistance, as mutations
altering inhibitor binding will simultaneously interfere with the
binding of substrates.
Results
Synthesis of ITMN-191. We synthesized the macrocyclic inhibitor
ITMN-191 using a convergent reaction sequence described in
SI Text. Briefly, the P2 and P1-P1′ fragments were preassembled
Author contributions: K.P.R., A.A., and C.A.S. designed research; K.P.R. and A.A. performed
research; A.A. and W.E.R. contributed new reagents/analytic tools; K.P.R., W.E.R., and
C.A.S. analyzed data; and K.P.R., A.A., W.E.R., and C.A.S. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The atomic coordinates and structure factors have been deposited in the
Protein Data Bank, www.pdb.org (PDB ID codes 3M5L, 3M5M, 3M5N, and 3M5O).
1K.P.R. and A.A. contributed equally to this work.
2To whom correspondence should be addressed. E-mail: Celia.Schiffer@umassmed.edu.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1006370107/-/DCSupplemental.
20986–20991 ∣PNAS ∣December 7, 2010 ∣vol. 107 ∣no. 49
www.pnas.org/cgi/doi/10.1073/pnas.1006370107
and the macrocyclic drug compound was generated by a four-
step reaction sequence, including P2-P3 amide coupling, ester
hydrolysis, coupling with the P1-P1′ fragment, and ring-closing
metathesis. The P2-P3 fragment was assembled by coupling the
commercially available Boc-protected amino acid (S)-2-(tert-
butoxycarbonylamino)non-8-enoic acid (Acme Biosciences, Inc)
with the preassembled P2 fragment, (3R, 5S)-5-(methoxy-
carbonyl)pyrrolidin-3-yl 4-fluoroisoindoline-2-carboxylate (31),
using O-(7-azabenzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexa-
fluorophosphate (HATU)/diisopropylethylamine (DIPEA). Hy-
drolysis of the P2-P3 methyl ester with LiOH:H2O in a mixture
of THF-MeOH-H2O followed by coupling of the resulting acid
under HATU/DIPEA conditions with the preassembled P1-P1′
fragment, (1R, 2S)-1-amino-N-(cyclopropylsulfonyl)-2-vinylcyclo-
propanecarboxamide (32), provided the bis-olefin precursor for
ring-closing metathesis. Cyclization of the bis-olefin intermediate
was accomplished using a highly efficient ring-closing metathesis
catalyst Zhan 1B and provided the protease inhibitor ITMN-191.
Structure Determination of Inhibitor and Substrate Complexes.
Although NS3/4A cleaves the viral polyprotein of over 3,000
residues at four specific sites in vivo, we focused on the local
interactions of the protease domain with short peptide sequences
corresponding to the immediate cleavage sites. All structural
studies were carried out with the highly soluble, single-chain con-
struct of the NS3/4A protease domain described previously (33),
which contains a fragment of the essential cofactor NS4A cova-
lently linked at the N terminus by a flexible linker. A similar pro-
tease construct was shown to retain comparable catalytic activity
to the authentic protein complex (34). Crystallization trials were
initially carried out using the inactive (S139A) protease variant
in complex with substrate peptides spanning P7-P5′. The 4A4B
substrate complex revealed cleavage of the scissile bond and no
ordered regions for the C-terminal fragment of the substrate.
Similar observations were previously described for two other
serine proteases where catalytic activity was observed, presum-
ably facilitated by water, despite Ala substitutions of the catalytic
Ser (35, 36). Thus all subsequent crystallization trials with the
NS3/4A protease were performed using N-terminal cleavage
products of the viral substrates spanning P7-P1.
NS3/4A crystal structures in complex with ITMN-191 and
peptide products 4A4B, 4B5A, and 5A5B were determined and
refined at 1.25 Å, 1.70 Å, 1.90 Å, and 1.60 Å resolution, respec-
tively (Table S2). The complexes crystallized in the space groups
P212121 and P21 with one, two, or four molecules in the asym-
metric unit. The average B factors range from 16.8–29.7 Å2
and there are no outliers in the Ramachandran plots. These
structures represent the highest resolution crystal structures of
NS3/4A protease reported to date.
Overall Structure Analysis. The NS3/4A protease domain adopts a
tertiary fold characteristic of serine proteases of the chymotrypsin
family (37, 38). A total of nine protease molecules were modeled
in the four crystal structures solved in this study with an overall
rms deviation (rmsd) of 0.28 Å. The rmsds reveal the five most
variable regions of the protease to be (Fig. S1): (i) the linker con-
necting cofactor 4A at the N terminus, (ii) the loop containing
residues 65–70, (iii) the zinc-binding site containing residues
95–105, (iv) the 310 helix region spanning residues 128–136,
and (v) the active site antiparallel β-sheet containing residues
156–168. These structural differences likely indicate inherent
flexibility in the protease and do not appear to correlate with
ligand type or active site occupancy.
Analysis of Product Complexes. Product complexes 4A4B, 4B5A,
and 5A5B were further analyzed with the C terminus of the
full-length NS3/4A structure (1CU1), which contains the N-term-
inal cleavage product of viral substrate 3-4A (39). All four
products bind to the protease active site in a conserved manner
(Fig. 2), forming an antiparallel β-sheet with residues 154–160
Fig. 1.
NS3/4A protease inhibitors and reported sites of drug resistance.
(A) The leading protease inhibitors in development mimic the N-terminal side
of the viral substrates. (B) The majority of reported drug resistance mutations
cluster around the protease active site with the catalytic triad depicted in
yellow.
Table 1. Drug resistance mutations reported in replicon studies and
clinical trials*
Residue
Mutation
Drug
V36
A, M, L, G
Boceprevir, telaprevir
Q41
R
Boceprevir, ITMN-191
F43
S, C, V, I
Boceprevir, telaprevir, ITMN-191,
TMC435
V55
A
Boceprevir
T54
A, S
Boceprevir, telaprevir
Q80
K, R, H, G, L
TMC435
S138
T
ITMN-191, TMC435†
R155
K, T, I, M, G, L, S, Q
Boceprevir, telaprevir, ITMN-191,
BILN-2061, TMC435
A156
V, T, S, I, G
Boceprevir, telaprevir, ITMN-191,
BILN-2061, TMC435
V158
I
Boceprevir
D168
A, V, E, G, N, T, Y, H, I
ITMN-191, BILN-2061, TMC435
V170
A
Boceprevir, telaprevir
M175
L
Boceprevir
*References (18, 25, 26, 28, 30–37).
†TMC435 displays reduced activity against S138T, but the mutation was not
observed in selection experiments.
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and burying 500–600 Å2 of solvent accessible surface area as cal-
culated by PISA (40). The peptide backbone torsions are very
similar, being most conserved at position P1 and deviating slightly
toward position P4. Eight hydrogen bonds between backbone
amide and carbonyl groups are completely conserved, involving
protease residues S159 (C159 in product 3-4A), A157, R155,
S139A, S138, and G137. S159 (C159 in product 3-4A), and
A157 each contribute two hydrogen bonds with the P5 and P3
peptide residues, respectively. All P1 terminal carboxyl groups
sit in the oxyanion hole, hydrogen bonding with the Nϵ atom
of H57 and the amide nitrogens of residues 137–139. Although
only product 4B5A contains a proline at P2, the other substrate
sequences still adopt constrained P2 φ torsion angles. Thus
products bind similarly despite their high sequence diversity.
The P1 and P6 residues are most conserved among the
substrate sequences, as are most of their interactions with the
protease. The P1 side chains interact with the aromatic ring of
F154. In all structures but product complex 4B5A, K165 forms
salt-bridges with the P6 acids, while residues R123, D168,
R155, and the catalytic D81 form an ionic network along one
surface of the bound products (Fig. 2). In complex 4B5A,
R123 interacts directly with the P6 acid, while D168 reorients
and no longer contacts R155. Other molecular interactions in
the product complexes are more diverse. Notably, K136 interacts
differently with the cleavage products, forming: (i) a hydrogen
bond with the P2 backbone carbonyl oxygen of 3-4A; (ii) a
salt-bridge with the P3 glutamate of product 4A4B; and (iii) non-
specific van der Waals interactions with the P2 and P3 side chains
of products 4B5A and 5A5B. Also, in product complex 4A4B, an
intramolecular hydrogen bond forms between the P3 and P5 glu-
tamate residues, while the unique P4 acid of product 3-4A forms
salt-bridges with the guanidinium groups of R123 and R155. Thus
distinct patterns of side chain interactions underlie the set of con-
served features involved in NS3/4A cleavage product binding.
The Substrate Envelope. To further analyze the structural similari-
ties of the four NS3/4A product complexes, the active sites were
superposed on the Cα atoms of residues 137–139 and 154–160,
revealing that both the active site residues and substrate products
spanning P6-P1 align closely with an average Cα rmsd of 0.24 Å
and 0.35 Å, respectively. The consensus van der Waals volume
shared by any three of the four cleavage products was then cal-
culated to generate the NS3/4A substrate envelope (Fig. 3A).
This shape could not be predicted by the primary sequences alone
and highlights the conserved mode of viral substrate recognition
despite their high sequence diversity.
Analysis of Inhibitor Complexes. ITMN-191, TMC435, and bocepre-
vir are all peptidomimetic NS3/4A protease inhibitors. Active site
superpositions of these drug complexes reveal that the inhibitors
interact with many of the same protease residues as the cleavage
products. Despite the P3-P1 cyclization of ITMN-191 and
TMC435, the functional groups are positioned similarly in all
three inhibitor complexes. The P1 cysteine surrogates interact
with the aromatic ring of F154, while the P2 and P3 moieties over-
lap closely. Although TMC435 does not contain a P4 substituent,
the P4 tert-butyl groups of ITMN-191 and boceprevir also align
closely. In addition, the P1 and P3 backbone atoms of all inhibi-
tors hydrogen bond with the carbonyl oxygens of R155 and A157,
respectively. These observations verify the peptidomimetic nat-
ure of these drugs and support their observed mechanism as
competitive active site inhibitors.
The largest variation between these three protease inhibitors
occurs at P2 where the aromatic rings of ITMN-191 and TMC435
stack against the guanidinium group of R155 (Fig. 3). This
molecular interaction alters the electrostatic network involving
R123, D168, R155, and D81. R155 rotates nearly 180° around
Cδ relative to its conformation observed in product complexes,
losing its hydrogen bond with D81 but maintaining interaction
with D168. Mutations at R155 or D168 would disrupt the elec-
trostatic network and destabilize this packing thereby lowering
the affinity of these macrocyclic drugs. This observation provides
a structural rationale for the drug resistance mutations R155K,
as previously proposed (19), and D168A/V, which both confer
a selective advantage in vitro in the presence of ITMN-191 or
TMC435 (26, 30). In addition, the TMC435 complex reveals
that R155 is stabilized by a hydrogen bond with Q80, which also
mutates to confer resistance to TMC435 (30). Thus many of the
primary drug resistance mutations can be explained by the disrup-
tion of atomic interactions involving the P2 functional groups of
the drugs.
Insights into Drug Resistance. To determine the locations where the
inhibitors protrude from the substrate envelope, the inhibitor and
product complexes were also superposed using residues 137–139
and 154–160. The van der Waals volumes of inhibitor protrusion
from the substrate envelope (V out) (41, 42) were calculated for
each drug and compared with published EC50 fold-change data
for drug resistance variants (30). The magnitudes of the EC50
fold-change data determined for each NS3/4A mutant generally
trend with the V out values for the three drugs. The P2 moieties of
boceprevir, ITMN-191 and TMC435 protrude most extensively
from the substrate envelope with V out values of 105, 294, and
Fig. 2.
Stereo view of N-terminal cleavage product binding to NS3/4A pro-
tease. N-terminal protease cleavage products (A) 3-4A, (B) 4A4B, (C) 4B5A,
and (D) 5A5B are depicted as they bind to the protease active site. All
conserved interactions are indicated by black dashes, while red lines depict
interactions that are not present in all structures. The electrostatic network
involving residues R123, D168, R155, and D81 is indicated by blue dashes.
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496 Å3, respectively (Table 2). However, the precise level of drug
resistance observed is also determined by the particular change
in molecular interaction occurring for a given mutation. For ex-
ample, A156 and R155 pack with the P2 moieties of these three
inhibitors where they protrude beyond the substrate envelope.
Mutations of A156 to bulkier side chains would result in a steric
clash with the P2 drug moieties. Indeed, the rigid dimethylcyclo-
propane group of boceprevir protrudes from the substrate envel-
ope at the P2 subsite, and A156V or A156T confer 65 and 75
fold-changes in EC50, respectively (Table 2). Similarly, molecular
changes at R155 and D168 would result in a substantial loss of
interactions with P2. The most extensive protrusions of ITMN-
191 and TMC435 at P2 trend with their greatest fold-change
in potency of nearly 450 and 600, respectively, from mutations
in this subsite. Thus the extent by which an inhibitor protrudes
from the substrate envelope in a given subsite is indicative of
its vulnerability to resistance.
Further structural analyses with the substrate envelope provide
insights into other NS3/4A drug resistance mutations. The P1′
sulfonamide groups of ITMN-191 and TMC435, as well as the
P1′ ketoamide of boceprevir, protrude from the substrate envel-
ope near residues Q41 and F43, which both mutate to confer
low-level resistance to these drugs (25, 30, 43). The keto group
of boceprevir also projects outside the substrate envelope near
T54 and V55. T54A/S confers low-level resistance to boceprevir,
while V55A was recently identified in patient isolates after treat-
ment with boceprevir (44). The analogous carbonyl groups of
ITMN-191 and TMC435, however, are orientated in the opposite
direction and protrude toward S138. In fact, in vitro studies reveal
reduced activity for ITMN-191 and TMC435 against S138T
variants, while boceprevir remains fully active (30, 43). The bulky
P4 tert-butyl group of boceprevir extends outside the substrate
envelope contacting V158; the V158I variant has lower affinity
for this drug, likely due to a steric clash (45). This variant may
also impact the affinity of ITMN-191, as its P4 tert-butyl also pro-
trudes at the same location. These findings demonstrate that in
regions outside the P2 subsite, positions where ITMN-191,
TMC435, and boceprevir protrude from the substrate envelope
also correlate with many other known sites of drug resistance
mutations.
Discussion
The emergence of drug resistance is a major obstacle in modern
medicine that limits the long-term usefulness of the most promis-
ing therapeutics. By considering how HCV NS3/4A protease in-
hibitors bind relative to natural viral substrates, we discovered
that primary sites of resistance occur in regions of the protease
where drugs protrude from the substrate envelope. In particular,
R155 and A156, which mutate to confer severe resistance against
ITMN-191, TMC435, and boceprevir, interact closely with the P2
drug moieties where they protrude most extensively from the
substrate envelope. Molecular changes at these residues confer
resistance by selectively weakening inhibitor binding without
compromising the binding of viral substrates. We further specu-
late that these mutations will not considerably affect the binding
of the host cellular substrates TRIF and MAVS, which likely fit
well within the substrate envelope as they share many features
with the viral substrates. However, TRIF contains a track of
eight proline residues instead of an acidic residue at position
P6, which may modulate its binding. Further structural studies
are warranted to better ascertain the molecular details of how
these cellular substrates are recognized by the NS3/4A protease.
Although this study focuses on ITMN-191, TMC435, and
boceprevir, other NS3/4A protease inhibitors in clinical trials, in-
cluding telaprevir, narlaprevir, and vaniprevir (Fig. 1A), contain
similar functional groups that likely protrude from the substrate
envelope. Most notably, all these drug candidates contain bulky
P2 moieties and are therefore susceptible to cross-resistance
against mutations at R155 and A156. R155 and A156 mutations
have been shown to confer telaprevir resistance in treated
patients (46). Cross-resistance studies have also shown that nar-
Table 2. EC50 fold-change (FC) data * for several NS3/4A drug
resistant variants tabulated with Vout, the van der Waals volume of
protrusion from the substrate envelope, at each subsite of the
enzyme
Boceprevir
ITMN-191
TMC435
Subsite
Resistance
mutation
Vout
(Å3)
EC50
FC
Vout
(Å3)
EC50
FC
Vout
(Å3)
EC50
FC
Total
292
500
649
P1
76
67
64
P2
105
294
496
Q80R
0.5
3.5
6.9
Q80K
0.8
2.3
7.7
R155K
4.7
447
30
A156V
75
63
177
A156T
65
41
44
D168A
0.7
153
594
D168E
0.8
75
40
P3
34
70
67
P4
76
69
0
V158I
3.3†
ND
ND
*Antiviral activity was reported previously by Lenz et al., 2010 (30).
†Fold-change in EC50 reported in replicon assay by Qiu et al., 2009 (45).
Fig. 3.
Stereo view of the NS3/4A substrate envelope and protease
inhibitors. (A) After active site superpositions, the overlapping van der Waals
volume shared by any three of the four cleavage products defines the
substrate envelope, depicted in blue. NS3/4A protease residues which mutate
to confer drug resistance are shown in brown. (B) ITMN-191, (C) boceprevir
and (D) TMC435 protrude from the substrate envelope at several locations,
which correlate with known sites of drug-resistant mutations to each inhibi-
tor, shown in red.
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laprevir displays similar fold losses in activity against most of the
known drug resistance mutations for telaprevir and boceprevir
(47). Ultimately, to slow the emergence of multidrug resistant
viral strains, inhibitors should be confined within the substrate
envelope, particularly at the P2 position. To compensate for the
loss of binding affinity that will likely accompany these changes,
additional interactions could potentially be optimized spanning
the S4-S6 subsites of the protease and the catalytic triad.
Our findings further suggest a general model for using the sub-
strate envelope to predict patterns of drug resistance in other
quickly evolving diseases. For drug resistance to occur, mutations
must selectively weaken a target’s affinity for an inhibitor without
significantly altering its natural biological function. Mutations
occurring outside the substrate envelope are better able to
achieve this effect, as these molecular changes can selectively
alter inhibitor binding without compromising the binding of
natural substrates. Whenever the interaction of a drug target with
its biological substrates can be structurally characterized, we pre-
dict that drugs designed to fit within the substrate envelope will
be less susceptible to resistance. Structure-based design strategies
can utilize this model as an added constraint to develop inhibitors
that fit within the substrate envelope. In fact, previous work in
our laboratory provides proof-of-concept for the successful incor-
poration of the substrate envelope in the design of unique HIV
protease inhibitors, which maintain high affinities against a panel
of multidrug resistant variants of HIV-1 protease (42, 48–53). As
a general paradigm, design efforts incorporating the substrate
envelope would facilitate a more rationale evaluation of drug
candidates and lead to the development of more robust inhibitors
that are less susceptible to resistance.
Materials and Methods
Protein Crystallization. The NS3/4A protease construct was expressed and
purified as reported previously (33, 54), detailed in SI Text. Purified protein
was concentrated to ∼3 mg∕mL and loaded on a HiLoad Superdex75 16/60
column equilibrated with 25 mM 2-(N-morpholino)ethanesulfonic acid (MES)
at pH 6.5, 500 mM NaCl, 10% glycerol, 30 μM zinc chloride, and 2 mM DTT.
The protease fractions were pooled and concentrated to 20–25 mg∕mL with
an Amicon Ultra-15 device (10 kD; Millipore). The concentrated samples were
then incubated at 4 °C for 1 h with 2–20 M excess of viral substrate 4A4B,
peptide products 4B5A or 5A5B, or ITMN-191. Information about the synth-
esis of viral peptides and ITMN-191 is provided in SI Text. Diffraction-quality
crystals were obtained overnight for all ligands by mixing equal volume of
concentrated protein solution with precipitant solution (20–26% PEG-3350,
0.1 M sodium MES buffer at pH 6.5, and 4% ammonium sulfate) in 24-well
VDX hanging drop trays.
Data Collection and Structure Solution. Crystals were flash-frozen in liquid
nitrogen and mounted under constant cryostream. X-ray diffraction data
were collected at Advanced Photon Source BioCARS 14-IDB, 14-BMC, and
LS-CAT 21-ID-F. Diffraction intensities of the complexes were indexed,
integrated, and scaled using the programs HKL2000 (55) and XDS (56). 5%
of the data was used to calculate R-free (57). All structure solutions were
generated using isomorphous molecular replacement with PHASER (58) or
AMORE. The NS3/4A protease domain (PDB code 2A4G) (59) was used for
molecular replacement in solving the product 4A4B structure, and this struc-
ture was subsequently used for solving the other complexes. In all cases,
initial refinement was carried out in the absence of modeled ligand, which
was subsequently built in. Phases were improved using ARP/wARP (60). Itera-
tive rounds of translation, libration, and screw (TLS) and restrained refine-
ment with CCP4 (61) and graphical model building with COOT (62) until
convergence was achieved. The final structures were evaluated with
MolProbity (63) prior to deposition in the protein data bank.
Structural Analysis. Double-difference plots (64) were used to determine the
structurally invariant regions of the protease, consisting of residues 32–36,
42–47, 50–54, 84–86, and 140–143. Structures were superposed with
PyMOL (65) using the Cα atoms of these residues for all protease molecules
from the solved structures (nine total). The B chain of product complex
4A4B was used as the reference structure in all alignments. Fit of individual
inhibitors into the substrate envelope was quantified by mapping the
substrate envelope and the van der Waals volume of each inhibitor on a
three-dimensional grid with spacing of 0.5 Å. Vout for each drug moiety
was computed by counting the grid cells, which were occupied by any inhi-
bitor atom of that site but not the substrate envelope, and multiplied by
the grid cell size, 0.125 Å3 (41, 42).
Substrate Envelope and Inhibitor Analyses. NS3/4A substrate envelope was
computed using product complexes 4A4B (B chain), 4B5A (D chain), and
5A5B (A chain). In structures with multiple protease molecules in the asym-
metric unit, the one containing the most ordered peptide product was used
for the alignment. The protease domain of the full-length NS3/4A structure
(A chain; PDB code 1CU1) (39), including the C-terminal six amino acids,
was included as a product complex 3-4A. All active site alignments were
performed in PyMOL using Cα atoms of protease residues 137–139 and
154–160. After superposition, Gaussian object maps were generated in Py-
MOL for each cleavage product. Four consensus Gaussian maps were then
calculated, representing the intersecting volume of a group of three object
maps. Finally, the summation of these four consensus maps was generated to
construct the substrate envelope, depicting the van der Waals volume shared
by any three of the four products. The previously determined boceprevir
complex (PDB code 2OC8) (24) and TMC435 complex (PDB code 3KEE) (23)
were used in this study (66).
ACKNOWLEDGMENTS. We thank H. Klei for helpful discussions. We also thank
Z. Wawrzak, M. Bolbat, and K. Brister of the LS-CAT beamline at Argonne
National Laboratory for data collection of the ITMN-191 complex; M. Nalam
and R. Bandaranayake for assistance with structural refinement; A. Ozen for
providing V out calculations; and S. Shandilya and Y. Cai for computational
support. The National Institute of Health (NIH) Grants R01-GM65347 and
R01-AI085051 supported this work. Use of Advanced Photon Source (APS)
was supported by the Department of Energy (DOE), Basic Energy Sciences,
Office of Science, under Contract No. DE-AC02-06CH11357. Use of the Bio-
CARS Sector 14 was supported by NIH-NCRR RR007707. Use of the LS-CAT
Sector 21 was supported by the Michigan Economic Development Corpora-
tion and the Michigan Technology TriCorridor under Grant 085P1000817.
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|
3M5N
|
Crystal structure of HCV NS3/4A protease in complex with N-terminal product 4B5A
|
Drug resistance against HCV NS3/4A inhibitors is
defined by the balance of substrate recognition
versus inhibitor binding
Keith P. Romano1, Akbar Ali1, William E. Royer, and Celia A. Schiffer2
Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, MA 01605
Edited by John M. Coffin, Tufts University School of Medicine, Boston, MA, and approved September 14, 2010 (received for review May 13, 2010)
Hepatitis C virus infects an estimated 180 million people world-
wide, prompting enormous efforts to develop inhibitors targeting
the essential NS3/4A protease. Resistance against the most promis-
ing protease inhibitors, telaprevir, boceprevir, and ITMN-191, has
emerged in clinical trials. In this study, crystal structures of the
NS3/4A protease domain reveal that viral substrates bind to the
protease active site in a conserved manner defining a consensus
volume, or substrate envelope. Mutations that confer the most
severe resistance in the clinic occur where the inhibitors protrude
from the substrate envelope, as these changes selectively weaken
inhibitor binding without compromising the binding of substrates.
These findings suggest a general model for predicting the suscept-
ibility of protease inhibitors to resistance: drugs designed to fit
within the substrate envelope will be less susceptible to resistance,
as mutations affecting inhibitor binding would simultaneously
interfere with the recognition of viral substrates.
drug design ∣hepatitis C ∣substrate envelope
D
rug resistance is a major obstacle in the treatment of quickly
evolving diseases. Hepatitis C virus (HCV) is a genetically
diverse Hepacivirus of the Flaviviridae family infecting an esti-
mated 180 million people worldwide (1). The viral RNA genome
is translated as a single polyprotein and subsequently processed
by host-cell and viral proteases into structural (C, E1, E2, and p7)
and nonstructural (NS2, NS3, NS4A, NS4B, NS5A, and NS5B)
proteins (2). The viral RNA-dependent RNA polymerase, NS5B,
is inherently inaccurate and misincorporation of bases accounts
for a very high mutation rate (3). While some mutations are neu-
tral, others will alter the viability of the virus and propagate with
varying efficiencies in each patient. Thus HCV infected indivi-
duals will develop a heterogeneous population of virus variants
known as quasispecies (4). As patients begin treatment, the selec-
tive pressures of antiviral drugs will favor drug resistant variants
(5). Therefore, an inhibitor must not only recognize one protein
variant, but an ensemble of related enzymes. A detailed under-
standing of the atomic mechanisms of resistance is essential to
effectively combat drug resistance against HCV antivirals.
The essential HCV NS3/4A protease is an attractive therapeu-
tic target responsible for cleaving at least four sites along the viral
polyprotein. These sites share little sequence homology except
for an acid at position P6, Cys or Thr at P1, and Ser or Ala at
P1′ (Table S1). The first cleavage event at the 3-4A junction
occurs in cis as a unimolecular process, while the remaining sub-
strates are processed bimolecularly in trans. The NS3/4A protease
also cleaves the human cellular targets TRIF and MAVS, which
confounds the innate immune response to viral infection (6–8).
Early drug design efforts were hampered by the relatively shallow,
featureless architecture of the protease active site. The eventual
observation of N-terminal product inhibition served as a stepping
stone for the discovery of more potent peptidomimetic inhibitors
(9, 10). Over the past decade, pharmaceutical companies have
further developed these lead compounds. Many structure-activ-
ity-relationship (SAR) studies have been performed to evaluate
the effect of different functional moieties on protease inhibition
at positions P4-P1′ (11–17). Crystal structures have been deter-
mined of the NS3/4A protease domain bound to a variety of
inhibitors as well as of several drug resistant protease variants,
such as R155K and V36M (18, 19). These data elucidate the mo-
lecular interactions of NS3/4A with inhibitors and the effect of
specific drug resistance mutations on binding. These efforts, con-
ducted in parallel by several pharmaceutical companies, led to
the discovery of many protease inhibitors. Proof-of-concept for
the successful clinical activity of this drug class was first demon-
strated by the macrocyclic inhibitor BILN-2061 (Boehringer
Ingelheim) (20, 21), which was later dropped from clinical trials
in 2006 due to cardiotoxicity (22). Many other NS3/4A protease
inhibitors are currently in development, and telaprevir (Vertex),
boceprevir (Schering-Plough), and ITMN-191 (Intermune) lead
the way in advanced phases of human clinical trials (Fig. 1A).
Despite these successes, the rapid acquisition of drug resis-
tance has limited the efficacy of the most potent NS3/4A protease
inhibitors in both replicon studies and human clinical trials
(Fig. 1B and Table 1). In this study, we show that mutations con-
ferring the most severe resistance occur where the protease
extensively contacts the inhibitors but not the natural viral sub-
strates. Four crystal structures of the NS3/4A protease domain in
complex with the N-terminal products of viral substrates reveal a
conserved mode of substrate binding, with the consensus volume
defining the substrate envelope. The protease inhibitors ITMN-
191 (3M5L), TMC435 (3KEE) (23), and boceprevir (2OC8) (24)
protrude extensively from the substrate envelope in regions that
correlate with known sites of resistance mutations. Most notably,
the P2 moieties of all three drugs protrude to contact A156 and
R155, which mutate to confer high-level resistance against nearly
all drugs reported in the literature (25–30). These findings sug-
gest that drug resistance results from a change in molecular
recognition and imply that drugs designed to fit within the sub-
strate envelope will be less susceptible to resistance, as mutations
altering inhibitor binding will simultaneously interfere with the
binding of substrates.
Results
Synthesis of ITMN-191. We synthesized the macrocyclic inhibitor
ITMN-191 using a convergent reaction sequence described in
SI Text. Briefly, the P2 and P1-P1′ fragments were preassembled
Author contributions: K.P.R., A.A., and C.A.S. designed research; K.P.R. and A.A. performed
research; A.A. and W.E.R. contributed new reagents/analytic tools; K.P.R., W.E.R., and
C.A.S. analyzed data; and K.P.R., A.A., W.E.R., and C.A.S. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The atomic coordinates and structure factors have been deposited in the
Protein Data Bank, www.pdb.org (PDB ID codes 3M5L, 3M5M, 3M5N, and 3M5O).
1K.P.R. and A.A. contributed equally to this work.
2To whom correspondence should be addressed. E-mail: Celia.Schiffer@umassmed.edu.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1006370107/-/DCSupplemental.
20986–20991 ∣PNAS ∣December 7, 2010 ∣vol. 107 ∣no. 49
www.pnas.org/cgi/doi/10.1073/pnas.1006370107
and the macrocyclic drug compound was generated by a four-
step reaction sequence, including P2-P3 amide coupling, ester
hydrolysis, coupling with the P1-P1′ fragment, and ring-closing
metathesis. The P2-P3 fragment was assembled by coupling the
commercially available Boc-protected amino acid (S)-2-(tert-
butoxycarbonylamino)non-8-enoic acid (Acme Biosciences, Inc)
with the preassembled P2 fragment, (3R, 5S)-5-(methoxy-
carbonyl)pyrrolidin-3-yl 4-fluoroisoindoline-2-carboxylate (31),
using O-(7-azabenzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexa-
fluorophosphate (HATU)/diisopropylethylamine (DIPEA). Hy-
drolysis of the P2-P3 methyl ester with LiOH:H2O in a mixture
of THF-MeOH-H2O followed by coupling of the resulting acid
under HATU/DIPEA conditions with the preassembled P1-P1′
fragment, (1R, 2S)-1-amino-N-(cyclopropylsulfonyl)-2-vinylcyclo-
propanecarboxamide (32), provided the bis-olefin precursor for
ring-closing metathesis. Cyclization of the bis-olefin intermediate
was accomplished using a highly efficient ring-closing metathesis
catalyst Zhan 1B and provided the protease inhibitor ITMN-191.
Structure Determination of Inhibitor and Substrate Complexes.
Although NS3/4A cleaves the viral polyprotein of over 3,000
residues at four specific sites in vivo, we focused on the local
interactions of the protease domain with short peptide sequences
corresponding to the immediate cleavage sites. All structural
studies were carried out with the highly soluble, single-chain con-
struct of the NS3/4A protease domain described previously (33),
which contains a fragment of the essential cofactor NS4A cova-
lently linked at the N terminus by a flexible linker. A similar pro-
tease construct was shown to retain comparable catalytic activity
to the authentic protein complex (34). Crystallization trials were
initially carried out using the inactive (S139A) protease variant
in complex with substrate peptides spanning P7-P5′. The 4A4B
substrate complex revealed cleavage of the scissile bond and no
ordered regions for the C-terminal fragment of the substrate.
Similar observations were previously described for two other
serine proteases where catalytic activity was observed, presum-
ably facilitated by water, despite Ala substitutions of the catalytic
Ser (35, 36). Thus all subsequent crystallization trials with the
NS3/4A protease were performed using N-terminal cleavage
products of the viral substrates spanning P7-P1.
NS3/4A crystal structures in complex with ITMN-191 and
peptide products 4A4B, 4B5A, and 5A5B were determined and
refined at 1.25 Å, 1.70 Å, 1.90 Å, and 1.60 Å resolution, respec-
tively (Table S2). The complexes crystallized in the space groups
P212121 and P21 with one, two, or four molecules in the asym-
metric unit. The average B factors range from 16.8–29.7 Å2
and there are no outliers in the Ramachandran plots. These
structures represent the highest resolution crystal structures of
NS3/4A protease reported to date.
Overall Structure Analysis. The NS3/4A protease domain adopts a
tertiary fold characteristic of serine proteases of the chymotrypsin
family (37, 38). A total of nine protease molecules were modeled
in the four crystal structures solved in this study with an overall
rms deviation (rmsd) of 0.28 Å. The rmsds reveal the five most
variable regions of the protease to be (Fig. S1): (i) the linker con-
necting cofactor 4A at the N terminus, (ii) the loop containing
residues 65–70, (iii) the zinc-binding site containing residues
95–105, (iv) the 310 helix region spanning residues 128–136,
and (v) the active site antiparallel β-sheet containing residues
156–168. These structural differences likely indicate inherent
flexibility in the protease and do not appear to correlate with
ligand type or active site occupancy.
Analysis of Product Complexes. Product complexes 4A4B, 4B5A,
and 5A5B were further analyzed with the C terminus of the
full-length NS3/4A structure (1CU1), which contains the N-term-
inal cleavage product of viral substrate 3-4A (39). All four
products bind to the protease active site in a conserved manner
(Fig. 2), forming an antiparallel β-sheet with residues 154–160
Fig. 1.
NS3/4A protease inhibitors and reported sites of drug resistance.
(A) The leading protease inhibitors in development mimic the N-terminal side
of the viral substrates. (B) The majority of reported drug resistance mutations
cluster around the protease active site with the catalytic triad depicted in
yellow.
Table 1. Drug resistance mutations reported in replicon studies and
clinical trials*
Residue
Mutation
Drug
V36
A, M, L, G
Boceprevir, telaprevir
Q41
R
Boceprevir, ITMN-191
F43
S, C, V, I
Boceprevir, telaprevir, ITMN-191,
TMC435
V55
A
Boceprevir
T54
A, S
Boceprevir, telaprevir
Q80
K, R, H, G, L
TMC435
S138
T
ITMN-191, TMC435†
R155
K, T, I, M, G, L, S, Q
Boceprevir, telaprevir, ITMN-191,
BILN-2061, TMC435
A156
V, T, S, I, G
Boceprevir, telaprevir, ITMN-191,
BILN-2061, TMC435
V158
I
Boceprevir
D168
A, V, E, G, N, T, Y, H, I
ITMN-191, BILN-2061, TMC435
V170
A
Boceprevir, telaprevir
M175
L
Boceprevir
*References (18, 25, 26, 28, 30–37).
†TMC435 displays reduced activity against S138T, but the mutation was not
observed in selection experiments.
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and burying 500–600 Å2 of solvent accessible surface area as cal-
culated by PISA (40). The peptide backbone torsions are very
similar, being most conserved at position P1 and deviating slightly
toward position P4. Eight hydrogen bonds between backbone
amide and carbonyl groups are completely conserved, involving
protease residues S159 (C159 in product 3-4A), A157, R155,
S139A, S138, and G137. S159 (C159 in product 3-4A), and
A157 each contribute two hydrogen bonds with the P5 and P3
peptide residues, respectively. All P1 terminal carboxyl groups
sit in the oxyanion hole, hydrogen bonding with the Nϵ atom
of H57 and the amide nitrogens of residues 137–139. Although
only product 4B5A contains a proline at P2, the other substrate
sequences still adopt constrained P2 φ torsion angles. Thus
products bind similarly despite their high sequence diversity.
The P1 and P6 residues are most conserved among the
substrate sequences, as are most of their interactions with the
protease. The P1 side chains interact with the aromatic ring of
F154. In all structures but product complex 4B5A, K165 forms
salt-bridges with the P6 acids, while residues R123, D168,
R155, and the catalytic D81 form an ionic network along one
surface of the bound products (Fig. 2). In complex 4B5A,
R123 interacts directly with the P6 acid, while D168 reorients
and no longer contacts R155. Other molecular interactions in
the product complexes are more diverse. Notably, K136 interacts
differently with the cleavage products, forming: (i) a hydrogen
bond with the P2 backbone carbonyl oxygen of 3-4A; (ii) a
salt-bridge with the P3 glutamate of product 4A4B; and (iii) non-
specific van der Waals interactions with the P2 and P3 side chains
of products 4B5A and 5A5B. Also, in product complex 4A4B, an
intramolecular hydrogen bond forms between the P3 and P5 glu-
tamate residues, while the unique P4 acid of product 3-4A forms
salt-bridges with the guanidinium groups of R123 and R155. Thus
distinct patterns of side chain interactions underlie the set of con-
served features involved in NS3/4A cleavage product binding.
The Substrate Envelope. To further analyze the structural similari-
ties of the four NS3/4A product complexes, the active sites were
superposed on the Cα atoms of residues 137–139 and 154–160,
revealing that both the active site residues and substrate products
spanning P6-P1 align closely with an average Cα rmsd of 0.24 Å
and 0.35 Å, respectively. The consensus van der Waals volume
shared by any three of the four cleavage products was then cal-
culated to generate the NS3/4A substrate envelope (Fig. 3A).
This shape could not be predicted by the primary sequences alone
and highlights the conserved mode of viral substrate recognition
despite their high sequence diversity.
Analysis of Inhibitor Complexes. ITMN-191, TMC435, and bocepre-
vir are all peptidomimetic NS3/4A protease inhibitors. Active site
superpositions of these drug complexes reveal that the inhibitors
interact with many of the same protease residues as the cleavage
products. Despite the P3-P1 cyclization of ITMN-191 and
TMC435, the functional groups are positioned similarly in all
three inhibitor complexes. The P1 cysteine surrogates interact
with the aromatic ring of F154, while the P2 and P3 moieties over-
lap closely. Although TMC435 does not contain a P4 substituent,
the P4 tert-butyl groups of ITMN-191 and boceprevir also align
closely. In addition, the P1 and P3 backbone atoms of all inhibi-
tors hydrogen bond with the carbonyl oxygens of R155 and A157,
respectively. These observations verify the peptidomimetic nat-
ure of these drugs and support their observed mechanism as
competitive active site inhibitors.
The largest variation between these three protease inhibitors
occurs at P2 where the aromatic rings of ITMN-191 and TMC435
stack against the guanidinium group of R155 (Fig. 3). This
molecular interaction alters the electrostatic network involving
R123, D168, R155, and D81. R155 rotates nearly 180° around
Cδ relative to its conformation observed in product complexes,
losing its hydrogen bond with D81 but maintaining interaction
with D168. Mutations at R155 or D168 would disrupt the elec-
trostatic network and destabilize this packing thereby lowering
the affinity of these macrocyclic drugs. This observation provides
a structural rationale for the drug resistance mutations R155K,
as previously proposed (19), and D168A/V, which both confer
a selective advantage in vitro in the presence of ITMN-191 or
TMC435 (26, 30). In addition, the TMC435 complex reveals
that R155 is stabilized by a hydrogen bond with Q80, which also
mutates to confer resistance to TMC435 (30). Thus many of the
primary drug resistance mutations can be explained by the disrup-
tion of atomic interactions involving the P2 functional groups of
the drugs.
Insights into Drug Resistance. To determine the locations where the
inhibitors protrude from the substrate envelope, the inhibitor and
product complexes were also superposed using residues 137–139
and 154–160. The van der Waals volumes of inhibitor protrusion
from the substrate envelope (V out) (41, 42) were calculated for
each drug and compared with published EC50 fold-change data
for drug resistance variants (30). The magnitudes of the EC50
fold-change data determined for each NS3/4A mutant generally
trend with the V out values for the three drugs. The P2 moieties of
boceprevir, ITMN-191 and TMC435 protrude most extensively
from the substrate envelope with V out values of 105, 294, and
Fig. 2.
Stereo view of N-terminal cleavage product binding to NS3/4A pro-
tease. N-terminal protease cleavage products (A) 3-4A, (B) 4A4B, (C) 4B5A,
and (D) 5A5B are depicted as they bind to the protease active site. All
conserved interactions are indicated by black dashes, while red lines depict
interactions that are not present in all structures. The electrostatic network
involving residues R123, D168, R155, and D81 is indicated by blue dashes.
20988
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Romano et al.
496 Å3, respectively (Table 2). However, the precise level of drug
resistance observed is also determined by the particular change
in molecular interaction occurring for a given mutation. For ex-
ample, A156 and R155 pack with the P2 moieties of these three
inhibitors where they protrude beyond the substrate envelope.
Mutations of A156 to bulkier side chains would result in a steric
clash with the P2 drug moieties. Indeed, the rigid dimethylcyclo-
propane group of boceprevir protrudes from the substrate envel-
ope at the P2 subsite, and A156V or A156T confer 65 and 75
fold-changes in EC50, respectively (Table 2). Similarly, molecular
changes at R155 and D168 would result in a substantial loss of
interactions with P2. The most extensive protrusions of ITMN-
191 and TMC435 at P2 trend with their greatest fold-change
in potency of nearly 450 and 600, respectively, from mutations
in this subsite. Thus the extent by which an inhibitor protrudes
from the substrate envelope in a given subsite is indicative of
its vulnerability to resistance.
Further structural analyses with the substrate envelope provide
insights into other NS3/4A drug resistance mutations. The P1′
sulfonamide groups of ITMN-191 and TMC435, as well as the
P1′ ketoamide of boceprevir, protrude from the substrate envel-
ope near residues Q41 and F43, which both mutate to confer
low-level resistance to these drugs (25, 30, 43). The keto group
of boceprevir also projects outside the substrate envelope near
T54 and V55. T54A/S confers low-level resistance to boceprevir,
while V55A was recently identified in patient isolates after treat-
ment with boceprevir (44). The analogous carbonyl groups of
ITMN-191 and TMC435, however, are orientated in the opposite
direction and protrude toward S138. In fact, in vitro studies reveal
reduced activity for ITMN-191 and TMC435 against S138T
variants, while boceprevir remains fully active (30, 43). The bulky
P4 tert-butyl group of boceprevir extends outside the substrate
envelope contacting V158; the V158I variant has lower affinity
for this drug, likely due to a steric clash (45). This variant may
also impact the affinity of ITMN-191, as its P4 tert-butyl also pro-
trudes at the same location. These findings demonstrate that in
regions outside the P2 subsite, positions where ITMN-191,
TMC435, and boceprevir protrude from the substrate envelope
also correlate with many other known sites of drug resistance
mutations.
Discussion
The emergence of drug resistance is a major obstacle in modern
medicine that limits the long-term usefulness of the most promis-
ing therapeutics. By considering how HCV NS3/4A protease in-
hibitors bind relative to natural viral substrates, we discovered
that primary sites of resistance occur in regions of the protease
where drugs protrude from the substrate envelope. In particular,
R155 and A156, which mutate to confer severe resistance against
ITMN-191, TMC435, and boceprevir, interact closely with the P2
drug moieties where they protrude most extensively from the
substrate envelope. Molecular changes at these residues confer
resistance by selectively weakening inhibitor binding without
compromising the binding of viral substrates. We further specu-
late that these mutations will not considerably affect the binding
of the host cellular substrates TRIF and MAVS, which likely fit
well within the substrate envelope as they share many features
with the viral substrates. However, TRIF contains a track of
eight proline residues instead of an acidic residue at position
P6, which may modulate its binding. Further structural studies
are warranted to better ascertain the molecular details of how
these cellular substrates are recognized by the NS3/4A protease.
Although this study focuses on ITMN-191, TMC435, and
boceprevir, other NS3/4A protease inhibitors in clinical trials, in-
cluding telaprevir, narlaprevir, and vaniprevir (Fig. 1A), contain
similar functional groups that likely protrude from the substrate
envelope. Most notably, all these drug candidates contain bulky
P2 moieties and are therefore susceptible to cross-resistance
against mutations at R155 and A156. R155 and A156 mutations
have been shown to confer telaprevir resistance in treated
patients (46). Cross-resistance studies have also shown that nar-
Table 2. EC50 fold-change (FC) data * for several NS3/4A drug
resistant variants tabulated with Vout, the van der Waals volume of
protrusion from the substrate envelope, at each subsite of the
enzyme
Boceprevir
ITMN-191
TMC435
Subsite
Resistance
mutation
Vout
(Å3)
EC50
FC
Vout
(Å3)
EC50
FC
Vout
(Å3)
EC50
FC
Total
292
500
649
P1
76
67
64
P2
105
294
496
Q80R
0.5
3.5
6.9
Q80K
0.8
2.3
7.7
R155K
4.7
447
30
A156V
75
63
177
A156T
65
41
44
D168A
0.7
153
594
D168E
0.8
75
40
P3
34
70
67
P4
76
69
0
V158I
3.3†
ND
ND
*Antiviral activity was reported previously by Lenz et al., 2010 (30).
†Fold-change in EC50 reported in replicon assay by Qiu et al., 2009 (45).
Fig. 3.
Stereo view of the NS3/4A substrate envelope and protease
inhibitors. (A) After active site superpositions, the overlapping van der Waals
volume shared by any three of the four cleavage products defines the
substrate envelope, depicted in blue. NS3/4A protease residues which mutate
to confer drug resistance are shown in brown. (B) ITMN-191, (C) boceprevir
and (D) TMC435 protrude from the substrate envelope at several locations,
which correlate with known sites of drug-resistant mutations to each inhibi-
tor, shown in red.
Romano et al.
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laprevir displays similar fold losses in activity against most of the
known drug resistance mutations for telaprevir and boceprevir
(47). Ultimately, to slow the emergence of multidrug resistant
viral strains, inhibitors should be confined within the substrate
envelope, particularly at the P2 position. To compensate for the
loss of binding affinity that will likely accompany these changes,
additional interactions could potentially be optimized spanning
the S4-S6 subsites of the protease and the catalytic triad.
Our findings further suggest a general model for using the sub-
strate envelope to predict patterns of drug resistance in other
quickly evolving diseases. For drug resistance to occur, mutations
must selectively weaken a target’s affinity for an inhibitor without
significantly altering its natural biological function. Mutations
occurring outside the substrate envelope are better able to
achieve this effect, as these molecular changes can selectively
alter inhibitor binding without compromising the binding of
natural substrates. Whenever the interaction of a drug target with
its biological substrates can be structurally characterized, we pre-
dict that drugs designed to fit within the substrate envelope will
be less susceptible to resistance. Structure-based design strategies
can utilize this model as an added constraint to develop inhibitors
that fit within the substrate envelope. In fact, previous work in
our laboratory provides proof-of-concept for the successful incor-
poration of the substrate envelope in the design of unique HIV
protease inhibitors, which maintain high affinities against a panel
of multidrug resistant variants of HIV-1 protease (42, 48–53). As
a general paradigm, design efforts incorporating the substrate
envelope would facilitate a more rationale evaluation of drug
candidates and lead to the development of more robust inhibitors
that are less susceptible to resistance.
Materials and Methods
Protein Crystallization. The NS3/4A protease construct was expressed and
purified as reported previously (33, 54), detailed in SI Text. Purified protein
was concentrated to ∼3 mg∕mL and loaded on a HiLoad Superdex75 16/60
column equilibrated with 25 mM 2-(N-morpholino)ethanesulfonic acid (MES)
at pH 6.5, 500 mM NaCl, 10% glycerol, 30 μM zinc chloride, and 2 mM DTT.
The protease fractions were pooled and concentrated to 20–25 mg∕mL with
an Amicon Ultra-15 device (10 kD; Millipore). The concentrated samples were
then incubated at 4 °C for 1 h with 2–20 M excess of viral substrate 4A4B,
peptide products 4B5A or 5A5B, or ITMN-191. Information about the synth-
esis of viral peptides and ITMN-191 is provided in SI Text. Diffraction-quality
crystals were obtained overnight for all ligands by mixing equal volume of
concentrated protein solution with precipitant solution (20–26% PEG-3350,
0.1 M sodium MES buffer at pH 6.5, and 4% ammonium sulfate) in 24-well
VDX hanging drop trays.
Data Collection and Structure Solution. Crystals were flash-frozen in liquid
nitrogen and mounted under constant cryostream. X-ray diffraction data
were collected at Advanced Photon Source BioCARS 14-IDB, 14-BMC, and
LS-CAT 21-ID-F. Diffraction intensities of the complexes were indexed,
integrated, and scaled using the programs HKL2000 (55) and XDS (56). 5%
of the data was used to calculate R-free (57). All structure solutions were
generated using isomorphous molecular replacement with PHASER (58) or
AMORE. The NS3/4A protease domain (PDB code 2A4G) (59) was used for
molecular replacement in solving the product 4A4B structure, and this struc-
ture was subsequently used for solving the other complexes. In all cases,
initial refinement was carried out in the absence of modeled ligand, which
was subsequently built in. Phases were improved using ARP/wARP (60). Itera-
tive rounds of translation, libration, and screw (TLS) and restrained refine-
ment with CCP4 (61) and graphical model building with COOT (62) until
convergence was achieved. The final structures were evaluated with
MolProbity (63) prior to deposition in the protein data bank.
Structural Analysis. Double-difference plots (64) were used to determine the
structurally invariant regions of the protease, consisting of residues 32–36,
42–47, 50–54, 84–86, and 140–143. Structures were superposed with
PyMOL (65) using the Cα atoms of these residues for all protease molecules
from the solved structures (nine total). The B chain of product complex
4A4B was used as the reference structure in all alignments. Fit of individual
inhibitors into the substrate envelope was quantified by mapping the
substrate envelope and the van der Waals volume of each inhibitor on a
three-dimensional grid with spacing of 0.5 Å. Vout for each drug moiety
was computed by counting the grid cells, which were occupied by any inhi-
bitor atom of that site but not the substrate envelope, and multiplied by
the grid cell size, 0.125 Å3 (41, 42).
Substrate Envelope and Inhibitor Analyses. NS3/4A substrate envelope was
computed using product complexes 4A4B (B chain), 4B5A (D chain), and
5A5B (A chain). In structures with multiple protease molecules in the asym-
metric unit, the one containing the most ordered peptide product was used
for the alignment. The protease domain of the full-length NS3/4A structure
(A chain; PDB code 1CU1) (39), including the C-terminal six amino acids,
was included as a product complex 3-4A. All active site alignments were
performed in PyMOL using Cα atoms of protease residues 137–139 and
154–160. After superposition, Gaussian object maps were generated in Py-
MOL for each cleavage product. Four consensus Gaussian maps were then
calculated, representing the intersecting volume of a group of three object
maps. Finally, the summation of these four consensus maps was generated to
construct the substrate envelope, depicting the van der Waals volume shared
by any three of the four products. The previously determined boceprevir
complex (PDB code 2OC8) (24) and TMC435 complex (PDB code 3KEE) (23)
were used in this study (66).
ACKNOWLEDGMENTS. We thank H. Klei for helpful discussions. We also thank
Z. Wawrzak, M. Bolbat, and K. Brister of the LS-CAT beamline at Argonne
National Laboratory for data collection of the ITMN-191 complex; M. Nalam
and R. Bandaranayake for assistance with structural refinement; A. Ozen for
providing V out calculations; and S. Shandilya and Y. Cai for computational
support. The National Institute of Health (NIH) Grants R01-GM65347 and
R01-AI085051 supported this work. Use of Advanced Photon Source (APS)
was supported by the Department of Energy (DOE), Basic Energy Sciences,
Office of Science, under Contract No. DE-AC02-06CH11357. Use of the Bio-
CARS Sector 14 was supported by NIH-NCRR RR007707. Use of the LS-CAT
Sector 21 was supported by the Michigan Economic Development Corpora-
tion and the Michigan Technology TriCorridor under Grant 085P1000817.
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|
3M5O
|
Crystal structure of HCV NS3/4A protease in complex with N-terminal product 5A5B
|
Drug resistance against HCV NS3/4A inhibitors is
defined by the balance of substrate recognition
versus inhibitor binding
Keith P. Romano1, Akbar Ali1, William E. Royer, and Celia A. Schiffer2
Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, MA 01605
Edited by John M. Coffin, Tufts University School of Medicine, Boston, MA, and approved September 14, 2010 (received for review May 13, 2010)
Hepatitis C virus infects an estimated 180 million people world-
wide, prompting enormous efforts to develop inhibitors targeting
the essential NS3/4A protease. Resistance against the most promis-
ing protease inhibitors, telaprevir, boceprevir, and ITMN-191, has
emerged in clinical trials. In this study, crystal structures of the
NS3/4A protease domain reveal that viral substrates bind to the
protease active site in a conserved manner defining a consensus
volume, or substrate envelope. Mutations that confer the most
severe resistance in the clinic occur where the inhibitors protrude
from the substrate envelope, as these changes selectively weaken
inhibitor binding without compromising the binding of substrates.
These findings suggest a general model for predicting the suscept-
ibility of protease inhibitors to resistance: drugs designed to fit
within the substrate envelope will be less susceptible to resistance,
as mutations affecting inhibitor binding would simultaneously
interfere with the recognition of viral substrates.
drug design ∣hepatitis C ∣substrate envelope
D
rug resistance is a major obstacle in the treatment of quickly
evolving diseases. Hepatitis C virus (HCV) is a genetically
diverse Hepacivirus of the Flaviviridae family infecting an esti-
mated 180 million people worldwide (1). The viral RNA genome
is translated as a single polyprotein and subsequently processed
by host-cell and viral proteases into structural (C, E1, E2, and p7)
and nonstructural (NS2, NS3, NS4A, NS4B, NS5A, and NS5B)
proteins (2). The viral RNA-dependent RNA polymerase, NS5B,
is inherently inaccurate and misincorporation of bases accounts
for a very high mutation rate (3). While some mutations are neu-
tral, others will alter the viability of the virus and propagate with
varying efficiencies in each patient. Thus HCV infected indivi-
duals will develop a heterogeneous population of virus variants
known as quasispecies (4). As patients begin treatment, the selec-
tive pressures of antiviral drugs will favor drug resistant variants
(5). Therefore, an inhibitor must not only recognize one protein
variant, but an ensemble of related enzymes. A detailed under-
standing of the atomic mechanisms of resistance is essential to
effectively combat drug resistance against HCV antivirals.
The essential HCV NS3/4A protease is an attractive therapeu-
tic target responsible for cleaving at least four sites along the viral
polyprotein. These sites share little sequence homology except
for an acid at position P6, Cys or Thr at P1, and Ser or Ala at
P1′ (Table S1). The first cleavage event at the 3-4A junction
occurs in cis as a unimolecular process, while the remaining sub-
strates are processed bimolecularly in trans. The NS3/4A protease
also cleaves the human cellular targets TRIF and MAVS, which
confounds the innate immune response to viral infection (6–8).
Early drug design efforts were hampered by the relatively shallow,
featureless architecture of the protease active site. The eventual
observation of N-terminal product inhibition served as a stepping
stone for the discovery of more potent peptidomimetic inhibitors
(9, 10). Over the past decade, pharmaceutical companies have
further developed these lead compounds. Many structure-activ-
ity-relationship (SAR) studies have been performed to evaluate
the effect of different functional moieties on protease inhibition
at positions P4-P1′ (11–17). Crystal structures have been deter-
mined of the NS3/4A protease domain bound to a variety of
inhibitors as well as of several drug resistant protease variants,
such as R155K and V36M (18, 19). These data elucidate the mo-
lecular interactions of NS3/4A with inhibitors and the effect of
specific drug resistance mutations on binding. These efforts, con-
ducted in parallel by several pharmaceutical companies, led to
the discovery of many protease inhibitors. Proof-of-concept for
the successful clinical activity of this drug class was first demon-
strated by the macrocyclic inhibitor BILN-2061 (Boehringer
Ingelheim) (20, 21), which was later dropped from clinical trials
in 2006 due to cardiotoxicity (22). Many other NS3/4A protease
inhibitors are currently in development, and telaprevir (Vertex),
boceprevir (Schering-Plough), and ITMN-191 (Intermune) lead
the way in advanced phases of human clinical trials (Fig. 1A).
Despite these successes, the rapid acquisition of drug resis-
tance has limited the efficacy of the most potent NS3/4A protease
inhibitors in both replicon studies and human clinical trials
(Fig. 1B and Table 1). In this study, we show that mutations con-
ferring the most severe resistance occur where the protease
extensively contacts the inhibitors but not the natural viral sub-
strates. Four crystal structures of the NS3/4A protease domain in
complex with the N-terminal products of viral substrates reveal a
conserved mode of substrate binding, with the consensus volume
defining the substrate envelope. The protease inhibitors ITMN-
191 (3M5L), TMC435 (3KEE) (23), and boceprevir (2OC8) (24)
protrude extensively from the substrate envelope in regions that
correlate with known sites of resistance mutations. Most notably,
the P2 moieties of all three drugs protrude to contact A156 and
R155, which mutate to confer high-level resistance against nearly
all drugs reported in the literature (25–30). These findings sug-
gest that drug resistance results from a change in molecular
recognition and imply that drugs designed to fit within the sub-
strate envelope will be less susceptible to resistance, as mutations
altering inhibitor binding will simultaneously interfere with the
binding of substrates.
Results
Synthesis of ITMN-191. We synthesized the macrocyclic inhibitor
ITMN-191 using a convergent reaction sequence described in
SI Text. Briefly, the P2 and P1-P1′ fragments were preassembled
Author contributions: K.P.R., A.A., and C.A.S. designed research; K.P.R. and A.A. performed
research; A.A. and W.E.R. contributed new reagents/analytic tools; K.P.R., W.E.R., and
C.A.S. analyzed data; and K.P.R., A.A., W.E.R., and C.A.S. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The atomic coordinates and structure factors have been deposited in the
Protein Data Bank, www.pdb.org (PDB ID codes 3M5L, 3M5M, 3M5N, and 3M5O).
1K.P.R. and A.A. contributed equally to this work.
2To whom correspondence should be addressed. E-mail: Celia.Schiffer@umassmed.edu.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1006370107/-/DCSupplemental.
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and the macrocyclic drug compound was generated by a four-
step reaction sequence, including P2-P3 amide coupling, ester
hydrolysis, coupling with the P1-P1′ fragment, and ring-closing
metathesis. The P2-P3 fragment was assembled by coupling the
commercially available Boc-protected amino acid (S)-2-(tert-
butoxycarbonylamino)non-8-enoic acid (Acme Biosciences, Inc)
with the preassembled P2 fragment, (3R, 5S)-5-(methoxy-
carbonyl)pyrrolidin-3-yl 4-fluoroisoindoline-2-carboxylate (31),
using O-(7-azabenzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexa-
fluorophosphate (HATU)/diisopropylethylamine (DIPEA). Hy-
drolysis of the P2-P3 methyl ester with LiOH:H2O in a mixture
of THF-MeOH-H2O followed by coupling of the resulting acid
under HATU/DIPEA conditions with the preassembled P1-P1′
fragment, (1R, 2S)-1-amino-N-(cyclopropylsulfonyl)-2-vinylcyclo-
propanecarboxamide (32), provided the bis-olefin precursor for
ring-closing metathesis. Cyclization of the bis-olefin intermediate
was accomplished using a highly efficient ring-closing metathesis
catalyst Zhan 1B and provided the protease inhibitor ITMN-191.
Structure Determination of Inhibitor and Substrate Complexes.
Although NS3/4A cleaves the viral polyprotein of over 3,000
residues at four specific sites in vivo, we focused on the local
interactions of the protease domain with short peptide sequences
corresponding to the immediate cleavage sites. All structural
studies were carried out with the highly soluble, single-chain con-
struct of the NS3/4A protease domain described previously (33),
which contains a fragment of the essential cofactor NS4A cova-
lently linked at the N terminus by a flexible linker. A similar pro-
tease construct was shown to retain comparable catalytic activity
to the authentic protein complex (34). Crystallization trials were
initially carried out using the inactive (S139A) protease variant
in complex with substrate peptides spanning P7-P5′. The 4A4B
substrate complex revealed cleavage of the scissile bond and no
ordered regions for the C-terminal fragment of the substrate.
Similar observations were previously described for two other
serine proteases where catalytic activity was observed, presum-
ably facilitated by water, despite Ala substitutions of the catalytic
Ser (35, 36). Thus all subsequent crystallization trials with the
NS3/4A protease were performed using N-terminal cleavage
products of the viral substrates spanning P7-P1.
NS3/4A crystal structures in complex with ITMN-191 and
peptide products 4A4B, 4B5A, and 5A5B were determined and
refined at 1.25 Å, 1.70 Å, 1.90 Å, and 1.60 Å resolution, respec-
tively (Table S2). The complexes crystallized in the space groups
P212121 and P21 with one, two, or four molecules in the asym-
metric unit. The average B factors range from 16.8–29.7 Å2
and there are no outliers in the Ramachandran plots. These
structures represent the highest resolution crystal structures of
NS3/4A protease reported to date.
Overall Structure Analysis. The NS3/4A protease domain adopts a
tertiary fold characteristic of serine proteases of the chymotrypsin
family (37, 38). A total of nine protease molecules were modeled
in the four crystal structures solved in this study with an overall
rms deviation (rmsd) of 0.28 Å. The rmsds reveal the five most
variable regions of the protease to be (Fig. S1): (i) the linker con-
necting cofactor 4A at the N terminus, (ii) the loop containing
residues 65–70, (iii) the zinc-binding site containing residues
95–105, (iv) the 310 helix region spanning residues 128–136,
and (v) the active site antiparallel β-sheet containing residues
156–168. These structural differences likely indicate inherent
flexibility in the protease and do not appear to correlate with
ligand type or active site occupancy.
Analysis of Product Complexes. Product complexes 4A4B, 4B5A,
and 5A5B were further analyzed with the C terminus of the
full-length NS3/4A structure (1CU1), which contains the N-term-
inal cleavage product of viral substrate 3-4A (39). All four
products bind to the protease active site in a conserved manner
(Fig. 2), forming an antiparallel β-sheet with residues 154–160
Fig. 1.
NS3/4A protease inhibitors and reported sites of drug resistance.
(A) The leading protease inhibitors in development mimic the N-terminal side
of the viral substrates. (B) The majority of reported drug resistance mutations
cluster around the protease active site with the catalytic triad depicted in
yellow.
Table 1. Drug resistance mutations reported in replicon studies and
clinical trials*
Residue
Mutation
Drug
V36
A, M, L, G
Boceprevir, telaprevir
Q41
R
Boceprevir, ITMN-191
F43
S, C, V, I
Boceprevir, telaprevir, ITMN-191,
TMC435
V55
A
Boceprevir
T54
A, S
Boceprevir, telaprevir
Q80
K, R, H, G, L
TMC435
S138
T
ITMN-191, TMC435†
R155
K, T, I, M, G, L, S, Q
Boceprevir, telaprevir, ITMN-191,
BILN-2061, TMC435
A156
V, T, S, I, G
Boceprevir, telaprevir, ITMN-191,
BILN-2061, TMC435
V158
I
Boceprevir
D168
A, V, E, G, N, T, Y, H, I
ITMN-191, BILN-2061, TMC435
V170
A
Boceprevir, telaprevir
M175
L
Boceprevir
*References (18, 25, 26, 28, 30–37).
†TMC435 displays reduced activity against S138T, but the mutation was not
observed in selection experiments.
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and burying 500–600 Å2 of solvent accessible surface area as cal-
culated by PISA (40). The peptide backbone torsions are very
similar, being most conserved at position P1 and deviating slightly
toward position P4. Eight hydrogen bonds between backbone
amide and carbonyl groups are completely conserved, involving
protease residues S159 (C159 in product 3-4A), A157, R155,
S139A, S138, and G137. S159 (C159 in product 3-4A), and
A157 each contribute two hydrogen bonds with the P5 and P3
peptide residues, respectively. All P1 terminal carboxyl groups
sit in the oxyanion hole, hydrogen bonding with the Nϵ atom
of H57 and the amide nitrogens of residues 137–139. Although
only product 4B5A contains a proline at P2, the other substrate
sequences still adopt constrained P2 φ torsion angles. Thus
products bind similarly despite their high sequence diversity.
The P1 and P6 residues are most conserved among the
substrate sequences, as are most of their interactions with the
protease. The P1 side chains interact with the aromatic ring of
F154. In all structures but product complex 4B5A, K165 forms
salt-bridges with the P6 acids, while residues R123, D168,
R155, and the catalytic D81 form an ionic network along one
surface of the bound products (Fig. 2). In complex 4B5A,
R123 interacts directly with the P6 acid, while D168 reorients
and no longer contacts R155. Other molecular interactions in
the product complexes are more diverse. Notably, K136 interacts
differently with the cleavage products, forming: (i) a hydrogen
bond with the P2 backbone carbonyl oxygen of 3-4A; (ii) a
salt-bridge with the P3 glutamate of product 4A4B; and (iii) non-
specific van der Waals interactions with the P2 and P3 side chains
of products 4B5A and 5A5B. Also, in product complex 4A4B, an
intramolecular hydrogen bond forms between the P3 and P5 glu-
tamate residues, while the unique P4 acid of product 3-4A forms
salt-bridges with the guanidinium groups of R123 and R155. Thus
distinct patterns of side chain interactions underlie the set of con-
served features involved in NS3/4A cleavage product binding.
The Substrate Envelope. To further analyze the structural similari-
ties of the four NS3/4A product complexes, the active sites were
superposed on the Cα atoms of residues 137–139 and 154–160,
revealing that both the active site residues and substrate products
spanning P6-P1 align closely with an average Cα rmsd of 0.24 Å
and 0.35 Å, respectively. The consensus van der Waals volume
shared by any three of the four cleavage products was then cal-
culated to generate the NS3/4A substrate envelope (Fig. 3A).
This shape could not be predicted by the primary sequences alone
and highlights the conserved mode of viral substrate recognition
despite their high sequence diversity.
Analysis of Inhibitor Complexes. ITMN-191, TMC435, and bocepre-
vir are all peptidomimetic NS3/4A protease inhibitors. Active site
superpositions of these drug complexes reveal that the inhibitors
interact with many of the same protease residues as the cleavage
products. Despite the P3-P1 cyclization of ITMN-191 and
TMC435, the functional groups are positioned similarly in all
three inhibitor complexes. The P1 cysteine surrogates interact
with the aromatic ring of F154, while the P2 and P3 moieties over-
lap closely. Although TMC435 does not contain a P4 substituent,
the P4 tert-butyl groups of ITMN-191 and boceprevir also align
closely. In addition, the P1 and P3 backbone atoms of all inhibi-
tors hydrogen bond with the carbonyl oxygens of R155 and A157,
respectively. These observations verify the peptidomimetic nat-
ure of these drugs and support their observed mechanism as
competitive active site inhibitors.
The largest variation between these three protease inhibitors
occurs at P2 where the aromatic rings of ITMN-191 and TMC435
stack against the guanidinium group of R155 (Fig. 3). This
molecular interaction alters the electrostatic network involving
R123, D168, R155, and D81. R155 rotates nearly 180° around
Cδ relative to its conformation observed in product complexes,
losing its hydrogen bond with D81 but maintaining interaction
with D168. Mutations at R155 or D168 would disrupt the elec-
trostatic network and destabilize this packing thereby lowering
the affinity of these macrocyclic drugs. This observation provides
a structural rationale for the drug resistance mutations R155K,
as previously proposed (19), and D168A/V, which both confer
a selective advantage in vitro in the presence of ITMN-191 or
TMC435 (26, 30). In addition, the TMC435 complex reveals
that R155 is stabilized by a hydrogen bond with Q80, which also
mutates to confer resistance to TMC435 (30). Thus many of the
primary drug resistance mutations can be explained by the disrup-
tion of atomic interactions involving the P2 functional groups of
the drugs.
Insights into Drug Resistance. To determine the locations where the
inhibitors protrude from the substrate envelope, the inhibitor and
product complexes were also superposed using residues 137–139
and 154–160. The van der Waals volumes of inhibitor protrusion
from the substrate envelope (V out) (41, 42) were calculated for
each drug and compared with published EC50 fold-change data
for drug resistance variants (30). The magnitudes of the EC50
fold-change data determined for each NS3/4A mutant generally
trend with the V out values for the three drugs. The P2 moieties of
boceprevir, ITMN-191 and TMC435 protrude most extensively
from the substrate envelope with V out values of 105, 294, and
Fig. 2.
Stereo view of N-terminal cleavage product binding to NS3/4A pro-
tease. N-terminal protease cleavage products (A) 3-4A, (B) 4A4B, (C) 4B5A,
and (D) 5A5B are depicted as they bind to the protease active site. All
conserved interactions are indicated by black dashes, while red lines depict
interactions that are not present in all structures. The electrostatic network
involving residues R123, D168, R155, and D81 is indicated by blue dashes.
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Romano et al.
496 Å3, respectively (Table 2). However, the precise level of drug
resistance observed is also determined by the particular change
in molecular interaction occurring for a given mutation. For ex-
ample, A156 and R155 pack with the P2 moieties of these three
inhibitors where they protrude beyond the substrate envelope.
Mutations of A156 to bulkier side chains would result in a steric
clash with the P2 drug moieties. Indeed, the rigid dimethylcyclo-
propane group of boceprevir protrudes from the substrate envel-
ope at the P2 subsite, and A156V or A156T confer 65 and 75
fold-changes in EC50, respectively (Table 2). Similarly, molecular
changes at R155 and D168 would result in a substantial loss of
interactions with P2. The most extensive protrusions of ITMN-
191 and TMC435 at P2 trend with their greatest fold-change
in potency of nearly 450 and 600, respectively, from mutations
in this subsite. Thus the extent by which an inhibitor protrudes
from the substrate envelope in a given subsite is indicative of
its vulnerability to resistance.
Further structural analyses with the substrate envelope provide
insights into other NS3/4A drug resistance mutations. The P1′
sulfonamide groups of ITMN-191 and TMC435, as well as the
P1′ ketoamide of boceprevir, protrude from the substrate envel-
ope near residues Q41 and F43, which both mutate to confer
low-level resistance to these drugs (25, 30, 43). The keto group
of boceprevir also projects outside the substrate envelope near
T54 and V55. T54A/S confers low-level resistance to boceprevir,
while V55A was recently identified in patient isolates after treat-
ment with boceprevir (44). The analogous carbonyl groups of
ITMN-191 and TMC435, however, are orientated in the opposite
direction and protrude toward S138. In fact, in vitro studies reveal
reduced activity for ITMN-191 and TMC435 against S138T
variants, while boceprevir remains fully active (30, 43). The bulky
P4 tert-butyl group of boceprevir extends outside the substrate
envelope contacting V158; the V158I variant has lower affinity
for this drug, likely due to a steric clash (45). This variant may
also impact the affinity of ITMN-191, as its P4 tert-butyl also pro-
trudes at the same location. These findings demonstrate that in
regions outside the P2 subsite, positions where ITMN-191,
TMC435, and boceprevir protrude from the substrate envelope
also correlate with many other known sites of drug resistance
mutations.
Discussion
The emergence of drug resistance is a major obstacle in modern
medicine that limits the long-term usefulness of the most promis-
ing therapeutics. By considering how HCV NS3/4A protease in-
hibitors bind relative to natural viral substrates, we discovered
that primary sites of resistance occur in regions of the protease
where drugs protrude from the substrate envelope. In particular,
R155 and A156, which mutate to confer severe resistance against
ITMN-191, TMC435, and boceprevir, interact closely with the P2
drug moieties where they protrude most extensively from the
substrate envelope. Molecular changes at these residues confer
resistance by selectively weakening inhibitor binding without
compromising the binding of viral substrates. We further specu-
late that these mutations will not considerably affect the binding
of the host cellular substrates TRIF and MAVS, which likely fit
well within the substrate envelope as they share many features
with the viral substrates. However, TRIF contains a track of
eight proline residues instead of an acidic residue at position
P6, which may modulate its binding. Further structural studies
are warranted to better ascertain the molecular details of how
these cellular substrates are recognized by the NS3/4A protease.
Although this study focuses on ITMN-191, TMC435, and
boceprevir, other NS3/4A protease inhibitors in clinical trials, in-
cluding telaprevir, narlaprevir, and vaniprevir (Fig. 1A), contain
similar functional groups that likely protrude from the substrate
envelope. Most notably, all these drug candidates contain bulky
P2 moieties and are therefore susceptible to cross-resistance
against mutations at R155 and A156. R155 and A156 mutations
have been shown to confer telaprevir resistance in treated
patients (46). Cross-resistance studies have also shown that nar-
Table 2. EC50 fold-change (FC) data * for several NS3/4A drug
resistant variants tabulated with Vout, the van der Waals volume of
protrusion from the substrate envelope, at each subsite of the
enzyme
Boceprevir
ITMN-191
TMC435
Subsite
Resistance
mutation
Vout
(Å3)
EC50
FC
Vout
(Å3)
EC50
FC
Vout
(Å3)
EC50
FC
Total
292
500
649
P1
76
67
64
P2
105
294
496
Q80R
0.5
3.5
6.9
Q80K
0.8
2.3
7.7
R155K
4.7
447
30
A156V
75
63
177
A156T
65
41
44
D168A
0.7
153
594
D168E
0.8
75
40
P3
34
70
67
P4
76
69
0
V158I
3.3†
ND
ND
*Antiviral activity was reported previously by Lenz et al., 2010 (30).
†Fold-change in EC50 reported in replicon assay by Qiu et al., 2009 (45).
Fig. 3.
Stereo view of the NS3/4A substrate envelope and protease
inhibitors. (A) After active site superpositions, the overlapping van der Waals
volume shared by any three of the four cleavage products defines the
substrate envelope, depicted in blue. NS3/4A protease residues which mutate
to confer drug resistance are shown in brown. (B) ITMN-191, (C) boceprevir
and (D) TMC435 protrude from the substrate envelope at several locations,
which correlate with known sites of drug-resistant mutations to each inhibi-
tor, shown in red.
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laprevir displays similar fold losses in activity against most of the
known drug resistance mutations for telaprevir and boceprevir
(47). Ultimately, to slow the emergence of multidrug resistant
viral strains, inhibitors should be confined within the substrate
envelope, particularly at the P2 position. To compensate for the
loss of binding affinity that will likely accompany these changes,
additional interactions could potentially be optimized spanning
the S4-S6 subsites of the protease and the catalytic triad.
Our findings further suggest a general model for using the sub-
strate envelope to predict patterns of drug resistance in other
quickly evolving diseases. For drug resistance to occur, mutations
must selectively weaken a target’s affinity for an inhibitor without
significantly altering its natural biological function. Mutations
occurring outside the substrate envelope are better able to
achieve this effect, as these molecular changes can selectively
alter inhibitor binding without compromising the binding of
natural substrates. Whenever the interaction of a drug target with
its biological substrates can be structurally characterized, we pre-
dict that drugs designed to fit within the substrate envelope will
be less susceptible to resistance. Structure-based design strategies
can utilize this model as an added constraint to develop inhibitors
that fit within the substrate envelope. In fact, previous work in
our laboratory provides proof-of-concept for the successful incor-
poration of the substrate envelope in the design of unique HIV
protease inhibitors, which maintain high affinities against a panel
of multidrug resistant variants of HIV-1 protease (42, 48–53). As
a general paradigm, design efforts incorporating the substrate
envelope would facilitate a more rationale evaluation of drug
candidates and lead to the development of more robust inhibitors
that are less susceptible to resistance.
Materials and Methods
Protein Crystallization. The NS3/4A protease construct was expressed and
purified as reported previously (33, 54), detailed in SI Text. Purified protein
was concentrated to ∼3 mg∕mL and loaded on a HiLoad Superdex75 16/60
column equilibrated with 25 mM 2-(N-morpholino)ethanesulfonic acid (MES)
at pH 6.5, 500 mM NaCl, 10% glycerol, 30 μM zinc chloride, and 2 mM DTT.
The protease fractions were pooled and concentrated to 20–25 mg∕mL with
an Amicon Ultra-15 device (10 kD; Millipore). The concentrated samples were
then incubated at 4 °C for 1 h with 2–20 M excess of viral substrate 4A4B,
peptide products 4B5A or 5A5B, or ITMN-191. Information about the synth-
esis of viral peptides and ITMN-191 is provided in SI Text. Diffraction-quality
crystals were obtained overnight for all ligands by mixing equal volume of
concentrated protein solution with precipitant solution (20–26% PEG-3350,
0.1 M sodium MES buffer at pH 6.5, and 4% ammonium sulfate) in 24-well
VDX hanging drop trays.
Data Collection and Structure Solution. Crystals were flash-frozen in liquid
nitrogen and mounted under constant cryostream. X-ray diffraction data
were collected at Advanced Photon Source BioCARS 14-IDB, 14-BMC, and
LS-CAT 21-ID-F. Diffraction intensities of the complexes were indexed,
integrated, and scaled using the programs HKL2000 (55) and XDS (56). 5%
of the data was used to calculate R-free (57). All structure solutions were
generated using isomorphous molecular replacement with PHASER (58) or
AMORE. The NS3/4A protease domain (PDB code 2A4G) (59) was used for
molecular replacement in solving the product 4A4B structure, and this struc-
ture was subsequently used for solving the other complexes. In all cases,
initial refinement was carried out in the absence of modeled ligand, which
was subsequently built in. Phases were improved using ARP/wARP (60). Itera-
tive rounds of translation, libration, and screw (TLS) and restrained refine-
ment with CCP4 (61) and graphical model building with COOT (62) until
convergence was achieved. The final structures were evaluated with
MolProbity (63) prior to deposition in the protein data bank.
Structural Analysis. Double-difference plots (64) were used to determine the
structurally invariant regions of the protease, consisting of residues 32–36,
42–47, 50–54, 84–86, and 140–143. Structures were superposed with
PyMOL (65) using the Cα atoms of these residues for all protease molecules
from the solved structures (nine total). The B chain of product complex
4A4B was used as the reference structure in all alignments. Fit of individual
inhibitors into the substrate envelope was quantified by mapping the
substrate envelope and the van der Waals volume of each inhibitor on a
three-dimensional grid with spacing of 0.5 Å. Vout for each drug moiety
was computed by counting the grid cells, which were occupied by any inhi-
bitor atom of that site but not the substrate envelope, and multiplied by
the grid cell size, 0.125 Å3 (41, 42).
Substrate Envelope and Inhibitor Analyses. NS3/4A substrate envelope was
computed using product complexes 4A4B (B chain), 4B5A (D chain), and
5A5B (A chain). In structures with multiple protease molecules in the asym-
metric unit, the one containing the most ordered peptide product was used
for the alignment. The protease domain of the full-length NS3/4A structure
(A chain; PDB code 1CU1) (39), including the C-terminal six amino acids,
was included as a product complex 3-4A. All active site alignments were
performed in PyMOL using Cα atoms of protease residues 137–139 and
154–160. After superposition, Gaussian object maps were generated in Py-
MOL for each cleavage product. Four consensus Gaussian maps were then
calculated, representing the intersecting volume of a group of three object
maps. Finally, the summation of these four consensus maps was generated to
construct the substrate envelope, depicting the van der Waals volume shared
by any three of the four products. The previously determined boceprevir
complex (PDB code 2OC8) (24) and TMC435 complex (PDB code 3KEE) (23)
were used in this study (66).
ACKNOWLEDGMENTS. We thank H. Klei for helpful discussions. We also thank
Z. Wawrzak, M. Bolbat, and K. Brister of the LS-CAT beamline at Argonne
National Laboratory for data collection of the ITMN-191 complex; M. Nalam
and R. Bandaranayake for assistance with structural refinement; A. Ozen for
providing V out calculations; and S. Shandilya and Y. Cai for computational
support. The National Institute of Health (NIH) Grants R01-GM65347 and
R01-AI085051 supported this work. Use of Advanced Photon Source (APS)
was supported by the Department of Energy (DOE), Basic Energy Sciences,
Office of Science, under Contract No. DE-AC02-06CH11357. Use of the Bio-
CARS Sector 14 was supported by NIH-NCRR RR007707. Use of the LS-CAT
Sector 21 was supported by the Michigan Economic Development Corpora-
tion and the Michigan Technology TriCorridor under Grant 085P1000817.
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PNAS
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December 7, 2010
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vol. 107
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no. 49
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20991
BIOPHYSICS AND
COMPUTATIONAL BIOLOGY
CHEMISTRY
|
3M5Q
|
0.93 A Structure of Manganese-Bound Manganese Peroxidase
|
Ultrahigh (0.93 Å) Resolution Structure of Manganese Peroxidase
from Phanerochaete chrysosporium: Implications for the Catalytic
Mechanism†,£
Munirathinam Sundaramoorthy‡,*, Michael H. Gold∥, and Thomas L. Poulos⊥
‡ Department of Biochemistry, Vanderbilt University Medical Center, Nashville, TN 37232
∥ Department of Biochemistry and Molecular Biology, OGI School of Science and Engineering,
Oregon Health and Science University, Portland, OR 97291-1000
⊥ Departments of Molecular Biology & Biochemistry, Chemistry, and Pharmaceutical Sciences
University of California, Irvine, CA 92697-3900
Abstract
Manganese peroxidase (MnP) is an extracellular heme enzyme produced by the lignin-degrading
white-rot fungus Phanerochaete chrysosporium. MnP catalyzes the peroxide-dependent oxidation
of MnII to MnIII. The MnIII is released from the enzyme in complex with oxalate, enabling the oxalate-
MnIII complex to serve as a diffusible redox mediator capable of oxidizing lignin, especially under
the mediation of unsaturated fatty acids. One heme propionate and the side chains of Glu35, Glu39
and Asp179 have been identified as MnII ligands in our previous crystal structures of native MnP.
In our current work, new 0.93 Å and 1.05 Å crystal structures of MnP with and without bound
MnII, respectively, have been solved. This represents only the fourth structure of a protein of this
size at 0.93 Å resolution. In addition, this is the first structure of a heme peroxidase from a eukaryotic
organism at sub-Ångstrom resolution. These new structures reveal an ordering/disordering of the C-
terminal loop, which is likely required for Mn binding and release. In addition, the catalytic Arg42
residue at the active site, normally thought to function only in the peroxide activation process, also
undergoes ordering/disordering that is coupled to a transient H-bond with the Mn ligand, Glu39.
Finally, these high-resolution structures also reveal the exact H atoms in several parts of the structure
that are relevant to the catalytic mechanism.
Keywords
Manganese; peroxidase; crystallography; atomic resolution; refinement
£Atomic coordinates have been submitted to the Protein Data Bank as entries XXX and YYY.
© 2010 Elsevier Inc. All rights reserved.
*To whom correspondence should be addressed: Department of Biochemistry, Vanderbilt University School of Medicine, 23rd @ Pierce,
626 RRB, Nashville, TN 37232. m.sundaramoorthy@vanderbilt.edu. Phone: (615) 343-1373.
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Published in final edited form as:
J Inorg Biochem. 2010 June ; 104(6): 683–690. doi:10.1016/j.jinorgbio.2010.02.011.
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INTRODUCTION
White-rot basidiomycetous fungi are the only organisms capable of degrading the
phenylpropanoid, plant cell wall polymer, lignin [1-4]. The lignin-degrading system of these
fungi also can oxidize a variety of economically and environmentally important aromatic
pollutants [5-9] and will probably play an important role in processes for the conversion of
lignocellulosics materials to ethanol. Under ligninolytic conditions, the best-studied lignin-
degrading fungus, Phanerochaete chrysosporium, secretes two families of extracellular heme
peroxidases, lignin peroxidase (LiP) and manganese peroxidase (MnP) [2-4,10] and a hydrogen
peroxide generating system [4,6,11,12].
MnP from P. chrysosporium has been studied by a variety of biochemical and biophysical
methods [2,13-16]. The crystal structure of MnP illustrates that the heme environment of this
enzyme is similar to that of other plant and fungal peroxidases [17,18]. However, MnP is the
only heme peroxidase capable of the one-electron oxidation of MnII in a typical peroxidase
reaction cycle:
(1)
(2)
(3)
where MnPI and MnPII are the oxidized intermediates MnP compounds I and II, respectively.
Our earlier crystal structures of MnP show that the substrate, MnII, binds to one heme
propionate and the side chains of three amino acids, Glu35, Glu39, and Asp179, as well as two
solvent ligands [17,18]. This site was proven by kinetic and biophysical studies of wild-type
MnP and of proteins containing point mutations in the putative binding site. Alteration of the
proposed amino acid ligands in the Mn-binding site significantly affects Mn-binding and
oxidation [19-24] and crystals of both the single variant, D179N, and the double variant, E35Q-
D179N, lack electron density at the proposed Mn-binding site [25] suggesting that MnII is not
bound. Furthermore, competitive inhibitors such as CdII, bind at the identical site, although
with alternative geometry [18,24].
MnP is unique among enzymes, using manganese as a redox cofactor. Rather than permanently
sequestering MnII in an interior binding site, MnP selectively binds MnII near the surface of
the protein, oxidizes it and then releases the MnIII product in complex with organic acids such
as oxalate. The relatively stable MnIII-oxalate complex acts as a diffusible mediator to oxidize
the terminal substrate lignin, usually in the presence of a radical mediator such as an unsaturated
fatty acid [26,27]. Whether a MnII-chelator complex binds to the enzyme to form a ternary
complex or the chelator simply facilitates release of MnIII via ligand displacement has now
been resolved. Both NMR and crystal structure studies of the MnP-MnII complex in the absence
of chelators indicate that the later alternative occurs [16,18,28-30].
Crystals of MnP are very stable and are good candidates for high resolution structure
determination [18]. Although a good deal is known about the catalytic mechanism of MnP and
of other plant and fungal peroxidases, atomic level structures could help reveal details of heme
geometry and H-bonding critical to the catalytic cycle of this enzyme. Atomic-detail structures
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can also show the exact stereochemical properties of various prosthetic groups bound to
proteins. Protein crystallographers normally rely on stereochemical parameters from crystal
structures of small molecules to constrain prosthetic groups during refinement and hence, the
final results are prejudiced toward the small molecule model. This is an important problem
since it is a common axiom in structural biology that the protein modulates the reactivity of
the prosthetic group by altering both electronic and geometric properties. Heme proteins
provide a classic example of how the same heme, iron protoporphyrin IX, is used in a vast
majority of heme proteins including the globins, cytochromes, and many heme enzymes, yet
these heme proteins exhibit widely different functions. A detailed comparison of known heme
protein structures illustrates that the heme group deviates significantly from planarity and that
heme proteins, which are functionally related, exhibit similar types of non-planarity [31]. Thus,
the ability to detail the geometry of such prosthetic groups is another benefit of ultrahigh
resolution protein structures.
The use of a new generation of synchrotron radiation sources coupled with cryogenic
techniques has enabled the solution of an increasing number of protein structures at true atomic
resolution. This has provided an unprecedented level of detail in analyzing functionally
important features of protein structures, including the identification of individual H atoms. The
precise location of H atoms is important in understanding enzyme reactions, where H-bonds
and acid-base catalysis are often utilized. Recent examples include the serine protease
structures from Bacillus lentus [32] and Titiachium album limber [33] solved at 0.78Å and
0.98Å, respectively. For Bacillus lentus subtilisin an H atom was found between the essential
catalytic His and Asp which forms part of the well-documented serine protease Ser-His-Asp
catalytic triad. In addition, the His-Asp H-bond distance was found to be relatively short, 2.62Å
[32]. Such results have important implications for the details of enzyme catalyzed reactions
and, in the subtilisin example, suggest the role that “low-barrier” H-bonds [34] play in the
reaction.
In this study, the structure of MnP containing 357 amino acid residues was refined at 0.93 Å
resolution. Of the total of about 37,000 protein structures determined by x-ray crystallography
and deposited in the PDB, only five unique structures (a bacterial catalase, 1GWE [35];
PfluDING, a DING protein from Pseudomonas fluorescens, 3G63 [36]; cholesterol oxidase
from Streptomyces sp., 1N4W [37]; xylose isomerase, 1MUW [38]; and pentaerythritol
tetranitrate reducatase, 1VYR [39] are comparable to MnP in both chain length (> 350 residues)
and resolution (≥ 0.93 Å). Thus MnP is the first eukaryotic heme peroxidase to be analyzed at
sub-Angstrom resolution.
MATERIAL AND METHODS
Protein Purification and Crystallization
Wild-type MnP was purified from shaking cultures of Phanerochaete chrysosporium grown
on high carbon, low nitrogen medium, as previously described [13,14,24]. MnII free MnP was
prepared using a metal Chelax-100 column as described [24]. Crystals of MnII bound MnP
(Mn-MnP) were grown at room temperature using the hanging drop vapor diffusion method
with an excess of 5 mM MnCl2 as described [18,40]. The reservoir contained 30% (w/v) of
polyethylene glycol 8,000, 0.2 M ammonium sulfate, and 0.1 M sodium cacodylate buffer, pH
6.5. The crystallization drops were composed of 5 μl of the protein solution (10-15 mg/ml)
mixed with an equal volume of reservoir solution. The crystallization was initiated by a seeding
procedure, using serially diluted seed stocks prepared from old native MnP crystals. Crystals
of MnII free MnP were grown, using similar reservoir conditions except that the drops
contained 4 mM EDTA instead of MnCl2 and the crystallization temperature was 4 °C.
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Data Collection and Processing
Crystals were harvested in synthetic mother liquor identical to the reservoir solution and
transferred to the cryo-solution containing 10% (v/v) glycerol in the mother liquor. The cryo-
soaked crystals were flash-frozen in an N2 cryostream and all data sets were collected at -160
°C.
The MnII free MnP data set was collected at the Stanford Synchrotron Radiation Laboratory
(SSRL) beam line 7-1 at 1.08 Å wave length. The diffraction images were recorded in two
steps using a single crystal on a MAR imaging plate at different detector settings and exposure
times. The detector distance was 80 mm for the MAR300 plate to record high resolution
reflections in dose mode with 3600 unit (~45 s) per frame and moved to 120 mm for the
MAR180 plate to collect low resolution data with a 20 s exposure per frame. A total of 180
frames of 1° oscillation per frame were collected for each set. The images of each set were
processed separately to extract raw intensities using MOSFLM [41] and were scaled together
using SCALA to obtain a single data set at 1.05 Å (Table 1).
The data set for the MnII bound MnP (Mn-MnP) was collected at SSRL Beam Line 9-1 at 0.78
Å wave length, using a single crystal. High resolution frames were collected using the MAR345
imaging plate at a 110 mm detector distance in dose mode with 6,000 units per frame. A total
of 180 frames of 0.75° oscillation angle were collected to record high resolution data. Low
resolution frames were collected using the MAR240 at 120 mm distance with 10 s exposure
per frame. A total of 180 frames of 1° per frame were collected for each set. The intensities of
each set were integrated separately using DENZO [42] and merged using SCALEPACK to
obtain a single data set 0.93 Å (Table 1).
Refinement
Coordinates of the native MnP structure previously refined at 1.45 Å resolution (PDB
Accession Code: 1YYD) were used as the starting model for the refinement with the MnII free
MnP data set which was collected first. The reflections used for Rfree calculation in the 1.45
Å data set were flagged in the present data set and extended to full resolution at 1.05 Å using
XPLOR [43]. However, in order to keep the unused reflections to a minimum, only 2% of the
reflections were flagged for the test set (Table 2). The first round of refinement was carried
out using data in the 8.0-1.45 Å resolution range by conjugate gradient least squares (CGLS)
protocol in SHELXL [44]. The model was examined from amino- to carboxy-terminus guided
by σA-weighted 2Fo-Fc and Fo-Fc maps. The electron density for the previously disordered C-
terminal loop was more ordered for this data set and residues 342-352 were rebuilt. The
resolution was extended gradually in a few rounds and the automatic water picking option
(PLAN 50 2.4) was used to add new waters to the model. The refined model and new water
positions were examined and manually corrected at each stage. The iterative cycles of manual
model adjustment and refinement coupled with resolution extension was continued until the
highest resolution of 1.05 Å was reached. Water molecules refined with a B-factor > 50 Å2
(with occupancy set to unity) were removed from the refinement throughout this exercise. At
the end of 10 rounds of refinement the Rcryst (Fo > 4 σ) was 0.159 and the corresponding
Rfree was 0.177. At this stage, anisotropic B-factor or atomic displacement parameter (ADP)
refinement was turned on using ANIS keyword for all atoms and the model was refined in 20
CGLS cycles. The Rcryst and Rfree dropped to 0.122 and 0.154, respectively. Following this
step, a few disordered sides chains were rebuilt in multiple conformations and the model was
further refined. When the refinement converged, 20 cycles of CGLS refinement was run with
riding hydrogen atoms for the polypeptide, sugar residues, and heme group. Both Rcryst and
Rfree dropped to 0.110 and 0.123, respectively. Finally, 10 cycles of CGLS refinement was
carried out with all reflections (i.e. working set + test set) to the final R-factor of 0.111.
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For the 0.93 Å Mn-MnP data set, test reflections were flagged using the 1.05 Å data set as the
reference and extended to full resolution. The refined model of the MnII free structure was used
as the starting model for the refinement. However, the refinement was started at 1.5 Å with
isotropic B-factors. The substrate MnII ion and its amino acid ligands were modeled using
σA-weighted 2Fo-Fc and Fo-Fc maps. The resolution was increased to 0.93 Å in five rounds
(1.2 Å, 1.05 Å, 1.00 Å, 0.95 Å and 0.93 Å) of CGLS refinement and the STIR command was
used to extend the resolution in finer steps in several cycles (20-35) in each round. The maps
were examined at the end of each refinement and minor changes were made to the model. New
waters were added in each round but those refined with high B-factors (> 50 Å2) were removed
in the subsequent refinement. The refinement converged with an Rcryst of 0.167 and an Rfree
of 0.179 with isotropic B-factors. Subsequently anisotropic B-factor refinement was turned on
and after extensive refinement the Rcryst and Rfree values converged to 0.12 and 0.143,
respectively. At this stage, a few disordered side chains were identified and modeled in multiple
conformations and refined further. When the refinement converged, 20 cycles of CGLS
refinement was run with riding H atoms for the polypeptide, sugar residues and heme group
(Rcryst = 0.109 and Rfree = 0.128). However, riding H atoms were not included for the proximal
heme ligand, His173, the residue H-bonded to this histidine, Asp242, the distal base catalyst
His46, and the residue H-bonded to this histidine, Asn80. Finally, 10 cycles of CGLS
refinement was carried out with all reflections (i.e. working set + test set) to the final R-factor
of 0.109.
The refinement statistics are shown in Table 2. The metal-ligand distances for the heme
FeIII, the substrate MnII, and two structural CaII ions, were unrestrained throughout the
refinement. After the anisotropic B-factor refinement began, side chain disorders were
examined and several side chains were modeled in multiple conformations with appropriate
partial occupancies and total occupancy was restrained to unity using the FVAR instruction.
A few other residues were found to be highly disordered but their side chains were not truncated.
For the calculation of estimated standard deviations (ESDs) for the atomic positions, one cycle
of full-matrix least-squares refinement was performed (BLOC 1).
RESULTS AND DISCUSSION
Quality of the Maps and Models
Crystals of MnP from P. chrysosporium are very robust with low mosaicity ( < 0.2°) and can
tolerate prolonged exposure as well as freeze-thaw cycles as observed in our previous study
[18]. These well ordered crystals can diffract to 1.0 Å or better at a synchrotron source. The
maps calculated using ultrahigh resolution data show high clarity that is typical of such data
sets (Figure 1). The previously reported 1.45 Å structure showed an O-glycosylation site at
Ser336 with a β-mannose residue and extra density at the N-glycosylation site at Asn131 that
could be due to a potential β-mannose residue [18]. These features are observed in the ultrahigh
resolution maps as well, but the maps do not show additional sugar residues at these sites nor
do they reveal new glycosylation sites. The electron density for most of the bound waters is
very clear and all of them were refined with unit occupancy. There remain small pieces of
uninterpretable density in several places in the solvent regions.
The final model of the substrate-bound MnP (Mn-MnP) consists of 357 amino acid residues,
three sugar residues (GlcNac,GlcNac at Asn131 and a single mannose at Ser336) , a heme
prosthetic group, two structural calcium ions, a substrate MnII ion, and 478 solvent molecules,
including two glycerol molecules. The substrate-free MnP model differs only in lacking the
MnII ion in the Mn binding site and in the number of solvent molecules, 549, which includes
two glycerol molecules. The two models superimpose with an overall RMS deviation of 0.253
Å for all 357 Cα atoms and of 0.146 Å for all residues minus the C-terminal loop (residues
344-350). There is no significant difference between these structures and the previously
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reported 1.45 Å cryo structure or the 2.06 Å room temperature structure in overall topology
(Figure 2). About 91% of the non-glycine residues are in the most favored regions of the
Ramachandran plot and the remaining are in either allowed or generously allowed regions.
Disorder
The N-terminal disorder persists for the first 10 residues in the maps of both the 1.05 Å and
the 0.93 Å data sets. However, there is an improvement in the quality of density near the C-
terminal loop encompassing Gly344-Gly350 in the 1.05 Å map of the substrate-free MnP data
set. Interestingly, this region is still disordered in the 0.93 Å map of the Mn-MnP data set
(Figure 3). This difference in the quality of electron density between the substrate-free and
substrate-bound forms is probably not due to the difference in their resolution. Instead, this
dynamic disorder is likely due to the binding of the substrate with implications for the catalytic
mechanism of the enzyme. This C-terminal loop traverses near the Mn binding pocket and
must be sufficiently flexible to allow binding of the substrate MnII, the entry of dicarboxylic
acid chelators such as oxalate to bind the oxidized MnIII product [16,29], and the release of
the MnIII-chelator complex. When we revisited the maps of the previously reported data sets
of native MnP (1.45 Å), CdII-MnP complex (1.6 Å), SmIII-MnP complex (1.6 Å) and oxalate
soaked SmIII-MnP complex (1.4 Å) similar differences are observed [18]. Whereas all three
metal bound structures show disorder, the map of oxalate soaked SmIII-MnP complex, which
lacks a metal ion, shows a more ordered C-terminal loop (data not shown).
Besides the disorder at the N- and C-termini, there are small sections of surface loops and the
side chains of a few surface residues that are disordered. Some of the disordered side chains
could be modeled in two conformations with partial occupancies in each. Notable among them
are a few cysteine residues, (Cys3, Cys14 and Cys253) which are found to be partially reduced
due to radiation, and the active site residue Arg42. In addition, two of the MnII ligands, Glu35
and Glu39, are disordered and exhibit multiple conformations in the MnII free structure. Other
disordered residues show a complete lack of density which could not be modeled with
confidence. Nevertheless, the side chains were not truncated and were modeled according to
the known sequence information [45,46]. These residues are characterized by high isotropic
B-factors and exhibit unrealistic atomic displacement parameters (ADPs) as analyzed by
PARVATI [47].
Mn Binding Site
The manganese binding site in both the substrate free MnP and Mn-MnP structures is shown
in Figure 4 and Table 3 provides ligand coordination distances. As described in our earlier
study, treating MnP with EDTA did not completely remove the metal from the substrate binding
site but did lead to a substantial decrease in electron density [25]. In our current work, however,
we were able to crystallize metal free MnP protein and the structure confirms the dynamic
nature of the active site described in our previous studies [18,25]. Two metal ligands, Glu35
and Glu39, move from their original MnII binding conformations and this provides insights
into the mechanism of MnP. Once MnP oxidizes MnII to MnIII, the MnIII acts as a diffusible
oxidant of lignin and other oxidizable substrates only when complexed with a suitable chelator
[16,29]. The latter is required to stabilize the highly reactive MnIII. The question is how such
a complex forms.
In the structures of the single mutant (D179N) and the double mutant (E35Q, D179N) MnPs,
which are essentially inactive and, which do not bind MnII, the side chain of Glu39 moves
away (“open” conformation) from the metal binding site [25]. Similarly, the metal free structure
of oxalate soaked SmIII-MnP complex structure showed Glu39 in the “open” conformation
[18]. Another Mn ligand, Glu35, undergoes a less dramatic conformational change in these
structures as its movement is restricted by a salt bridge with the side chain of Arg177 [21].
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Whereas the mobility of Glu39 is effected by a rotation of the Cα–Cβ bond resulting in a major
movement, Glu35 side chain is rotated about the Cβ–Cγ bond resulting in variable, but minor
changes in the wild-type and mutant structures depending upon the electronic environment of
the site. Other ligands—the heme propionate and Asp179—do not move from their original
positions whether a metal ion is bound or not, strongly suggesting that precise geometry is
required for efficient MnII-binding and oxidation. This is confirmed by steady-state and
transient-state kinetic analyses of a MnP E39D single mutant and an E35D-E39D-D179E triple
mutant [23]. The single and triple mutant variants exhibit 20- and 40-fold increase in Km, and
103 and 104 decrease in catalytic efficiency, respectively. Although the overall charge is
retained, a decrease in the chain length of one ligand could not be compensated for by an
increase in the chain length of another ligand for either MnII binding or oxidation. Our previous
study using the non-reducible/oxidizable trivalent cation SmIII showed that it could bind at the
Mn binding site of MnP [18]. In addition, the release of SmIII by oxalate from the pregrown
SmIII-MnP crystals provides insights into how the Mn binding site can bind both divalent and
trivalent cations and how the trivalent cation can be released from the resting enzyme by organic
acids [18]. Taken together, our past studies and the present high resolution structures imply
that MnII binding is precise and the site is relatively rigid, except for the ability of Glu35 and
Glu39 to adopt two conformations—“closed” conformations in the metal bound state and
“open” conformations in the metal free state, possibly acting as a “gate”, enabling a small
carboxylic acid like oxalate or malonate to remove MnIII from the binding site. Without some
flexibility in the Mn ligands, it is difficult to envisage how oxalate could remove MnIII from
its coordination shell unless MnIII is first freely released.
Another residue, Arg42, is also disordered, and this has significance for peroxidase function.
This arginine residue is conserved in all peroxidases and is implicated in stabilizing the
compound I and II intermediates by forming a hydrogen bond with the oxyferryl oxygen [48,
49]. This active site arginine residue appears disordered or in multiple conformations in this
high resolution structures of MnP and in cytochrome c peroxidase (CcP) [48] but is in a well
defined single conformation in the compound I structure of CcP [48]. Arg42 is disordered in
both substrate-bound and -free structures of MnP and is modeled in two conformations—“in”
and “out”, in which Arg42 moves closer to and away from the oxyferryl center. However, the
main difference occurs in its interaction with Glu39. Besides providing a ligand to MnII, the
carboxylate group of Glu39 also forms a salt bridge with the guanidium group of Arg42 in the
“out” conformation in the substrate-bound structure. This is similar to the salt bridge between
Glu35 and Arg177 described earlier [17,18,21,50]. However, in the substrate-free structure,
movement of Glu39 to the “open” conformation breaks this salt bridge and a water molecule
occupies the position of the carboxylate oxygen of Glu39. Thus, Arg42 may also stabilize
MnII binding in the resting state through its interaction with Glu39, but its movement to the
“in” conformation to stabilize the oxyferryl group in compounds I and II may destabilize Glu39
enabling it to move out for the oxidized MnIII to be chelated by oxalate. In contrast, the other
two amino acid ligands for MnII, Glu35 and Asp179, are held in place by the rigid guanidium
group of the Arg177 side chain and backbone amide group of residue Ala187, respectively. In
support of this idea, disruption of the salt bridge between Glu35 and Arg177, through mutation
of the arginine residue, has been shown to significantly lower the efficiency of MnII binding
and oxidation in our previous work [21,50]. This new role for Arg42 is revealed for the first
time in this high resolution structure of MnP.
Heme Stereochemistry
The overall structure of Mn-MnP (Figure 2) at 0.93Å appears essentially the same as that in
our earlier lower resolution work [17,18]. Given the favorable ratio of data to parameters to be
refined, the least squares matrix could be inverted which provides an accurate estimate of bond
distances and angles. The electron density for the heme and proximal heme ligand together,
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shown in Figure 5 and Table 4, provides specific heme parameters. As expected from
spectroscopic studies [51], the heme Fe is pentacoordinate and high-spin. A functionally
important parameter of heme geometry is the displacement of the iron from the porphyrin core.
In MnP the Fe is 0.26Å out of plane compared to 0.48Å for typical model heme complexes
[52]. The ability of the Fe atom to move in and out of the porphyrin core as a function of
ligation, redox state, and spin state is a central feature of the hemoglobin allosteric mechanism
proposed by Perutz [53]. Compared to the globins, the Fe atom in MnP and CcP is closer to
the porphyrin core. We have attributed this difference [54] to the surrounding protein
environment which in the peroxidases places the proximal helix containing the heme His ligand
much closer to the protein. This prevents the spring-like up/down motion of the Fe-His-helix
as seen in hemoglobin. The functional relevance of this difference is that in peroxidases, the
iron is oxidized from FeIII to FeIV=O. The higher oxidation state, lower-spin state, and strong
FeIV=O bond all favor the FeIV closer to the heme plane. Hence, the resting FeIII state with
only partial displacement of the Fe from the porphyrin plane is poised for oxidation to FeIV,
which is reminiscent of the entatic state [55], where the kinetic and/or thermodynamic barriers
required for changing redox and spin-state are lowered owing to protein-metal interactions.
Mechanism and H atoms
As with many enzyme systems, H-bonds and proton transfer play a critical role in peroxidase
catalysis. The original stereochemical mechanism proposed for heme peroxidases [56]
indicates that the distal histidine residue (His 46 in MnP, Figure 2) acts as an acid-base catalyst
by removing a proton from the iron-linked peroxide O atom and delivering it to the leaving
OH moiety to produce water. As shown in Figure 5A, which of the N atoms of the imidazole
moiety in His46 carries the proton is clearly visible in Fo-Fc electron density maps. The
Nδ1atom is protonated and donates an H-bond to Asn80. This His-Asn H-bonding arrangement
is conserved in all peroxidases and is thought to play an important role in orientating the distal
His for catalysis as well as ensuring that the Nε2 is free to accept a proton from hydrogen
peroxide. Another level of detail at 0.93 Å is the unambiguous orientation of amide side chains
such as that for Asn80. At lower resolution, it is not possible to differentiate between the
Nδ2 and Oδ1 side chain atoms. However, as shown in Figure 5A the amide side chain Oδ1 atom
of Asn80 exhibits smeared electron density with the Cγ side chain atom while the Cδ-Nε2
density is much shaper. This indicates a Cγ-Oδ1 double bond and an H bond between the Nε1
of His 46 and the Oδ1 of Asn80.
The role of H-bonds involving the proximal His ligand is not well documented. In all known
heme peroxidase structures, the His ligand is within H-bonding distance of a buried Asp,
Asp242 in MnP (Figure 2). A number of important functional properties have been attributed
to this H-bond, one of which involves redox potential. A strong His-Asp H-bond favors a lower
redox potential by stabilizing the additional positive charge on FeIV compared to FeIII. In the
globins the histidine ligand forms an H-bond with a peptide carbonyl O atom, which
presumably is weaker than the His-Asp H-bond in peroxidases. This difference helps to explain
why the peroxidases exhibit lower redox potentials than globins. This view is supported by
both model heme systems [57-60], protein NMR [61], and site directed mutagenesis [62,63].
The present structure provides a somewhat complex picture of the His-Asp interaction. As
shown in Figure 5B there is a strong lobe of electron density in both Fo-Fc and 2Fo-Fc maps at
about 1.34Å from Oδ2 of Asp242. Initially we considered that this peak might be an H atom
which would mean that Asp242 carries the proton and not His173. However, the O-H distance,
1.34Å, is too long, compared to other H atoms in the structure, and the electron density peak
too large for an H atom. Two additional data sets to 1.15Å and 1.05Å have been collected and
maps generated from these data sets also exhibit additional electron density between Asp242
and His173 (data not shown). The prospects of a low-barrier H-bond seems unlikely since the
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O-N hydrogen bonding distance is quite normal at 2.83Å while low-barrier H-bonds are much
shorter.. The exact explanation for this extra electron density remains unclear.
Acknowledgments
This research was supported by a grant GM42614 (to T.L.P.) from the National Institutes of Health and grants
MCB-9808420 from the National Science Foundation and DE-03-96ER20235 from the Division of Energy
Biosciences, U.S. Department of Energy (to M.H.G). Portions of this research were carried out at the Stanford
Synchrotron Radiation Laboratory, a national user facility operated by Stanford University on behalf of the U.S.
Department of Energy, Office of Basic Energy Sciences. The SSRL Structural Molecular Biology Program is supported
by the Department of Energy, Office of Biological and Environmental Research, and by the National Institutes of
Health, National Center for Research Resources, Biomedical Technology Program, and the National Institute of
General Medical Sciences.
ABBREVIATIONS
MnP
manganese peroxidase
Mn-MnP
manganese-enzyme complex
L.S.
least squares
CGLS
conjugate gradient least squares
ESD
estimated standard deviation
ADP
atomic displacement parameter
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Figure 1.
(A) 2Fo-Fc map density for the refined heme group, calculated using the 0.93 Å native Mn-
MnP data set. The contours are drawn at 1.0 σ (green) and 4.0 σ (magenta). (B) Thermal
ellipsoid diagram for the heme group drawn using ORTEP. The structure, including anisotropic
B-factors, was refined using the SHELXL program [44].
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Figure 2.
(A) The overall structure of MnP. The red spheres are structural CaII ions conserved in
extracellular heme peroxidases. The location of the substrate, MnII, near the heme, is indicated.
(B) The active site structure of MnP. This architecture is highly conserved in heme peroxidase.
The main variations are the Phe residues which are Trp in the intercellular peroxidases,
cytochrome c and ascorbate peroxidase. The Asp242-His173 pair is conserved.
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Figure 3.
(A). C-terminal region of the MnP structure. The loop encompassing residues 342-352
traverses close to the MnII binding site. This loop is disordered when MnII is bound in the 0.93
Å structure of native Mn-MnP but is ordered in the substrate free MnP structure at 1.05 Å.
2Fo-Fc electron density for the ordered (B) and disordered (C) C-terminal loop is shown
contoured at 1.0 σ.
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Figure 4.
Stereo views of the electron density maps obtained from the 0.93 Å Mn-MnP structure
contoured at 1.6σ (top) and 1.05Å Mn free MnP structure obtained from EDTA-treated crystal
contoured at 1.0σbottom). MnII (cyan) and all its ligands including two water molecules are
labeled for the MnII-bound structure. Glu39 is in a single conformation in this structure where
it is in two conformations in the MnII-free structure, as shown in the figure.
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Figure 5.
2Fo – Fc (green, 1 σ and red, 3 σ and Fo – Fc (blue, 3 σ) electron density maps in the distal
(A) and proximal (B) regions. The peaks (blue) in Fo – Fc map indicate potential hydrogen
peaks near the proximal Asp242 and distal His46.
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Table 1
Data collection statistics
Dataset
MnP (-Mn)
Mn-MnP
Resolution range (Å)
∞ - 1.05
∞ - 0.93
No. of observations
653,391
1,246,031
Unique reflections
150,666
264,958
Completeness (%)
91.2 (79.3)
97.5 (93.0)
Redundancy
4.1 (2.7)
4.9 (2.0)
<I>/<σ(I)>
9.0 (3.4)
26.3 (1.9)
Rsym
0.05 (0.21)
0.066 (0.512)
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Table 2
Refinement statistics
MnP (-Mn)
Mn-MnP
Resolution range (Å)
8.0 – 1.05
8.0 – 0.93
No of atoms
Amino acid residues
2,622
2,622
Sugar residues
39
39
Heme
43
43
CaII
2
2
MnII
-
1
Glycerol
24
12
Water
549
477
Rwork [working set, Fo > 4σ(Fo)]
11.1 (144,730)
10.7 (191,300)
Rwork (working set, all reflections)
11.7 (156,881)
12.4 (243,170)
Rfree [test set, Fo > 4σ(Fo)]
13.6 (3,181)
12.4 (3,967)
Rfree (test set, all reflections)
13. 9 (3,352)
13.4 (4,736)
Rcryst [working + test set, Fo > 4σ(Fo)]
11.1 (147,911)
10.8 (195,267)
Rcryst (working + test set, all reflections)
11.6 (160,232)
12.4 (247,906)
No. of Parameters
30,135
29,226
No. of observations/No. of parameters
5.3
8.5
No. of restraints
36,426
35,548
Mean isotropic B-factor (Å2)
All atoms
12.973
13.670
Main chain atoms
9.279
10.098
Side chain atoms
11.776
13.276
Sugar residues
17.171
16.416
Heme and metal ions
8.394
7.150
Solvent
24.909
25.687
RMS deviation from ideal geometry
Bond length (1-2) (Å)
0.017
0.018
Angle distance (1-3) (Å)
0.038
0.033
Chiral volume (Å3)
0.129
0.096
Non-zero chiral volume (Å3)
0.219
0.105
Deviation from planes (Å)
0.029
0.028
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Table 3
Metal-ligand distances
Metal – ligand
MnP (-Mn) (Å)
Mn-MnP (Å)
H2O*
MnII
Heme propionate
2.25
2.12
Glu35 OE1
2.36
2.14
Glu39 OE1
--
2.27
Asp179 OD1
--
2.28
Water 1 (1040)
2.56
2.25
Water 2 (1108)
2.64
2.26
Proximal CaII
Carbonyl O-Ser174
2.36
2.37
Side chain OG1-Ser174
2.47
2.47
Side chain carboxyl Asp191
2.42
2.44
Carbonyl O-Thr193
2.37
2.38
Side chain OG1-Thr193
2.50
2.49
Carbonyl O-Thr196
2.49
2.51
Side chain carboxyl Asp198
2.46
2.48
Distal CaII
Carbonyl O-Asp47
2.44
2.44
Side chain carboxyl Asp47
2.31
2.31
Carbonyl O-Gly62
2.44
2.46
Side chain carboxyl Asp64
2.39
2.42
Side chain O-Ser66
2.49
2.49
Water 1 (1085)
2.39
2.41
Water 2 (1027)
2.35
2.33
*A water or unidentified monovalent cation occupies the Mn-binding site in the substrate free structure.
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Sundaramoorthy et al.
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Table 4
Heme parameters
Distance (Å)
MnP (-Mn) (Å)
Mn-MnP (Å)
CcP (Å)
Fe—His N
2.10
2.07
2.07
Fe—pyrrole N
2.02
2.04
2.05
Fe—pyrrole N plane
0.28
J Inorg Biochem. Author manuscript; available in PMC 2011 June 1.
|
3M5X
|
Crystal structure of the mutant V182A,I199A of orotidine 5'-monophosphate decarboxylase from Methanobacterium thermoautotrophicum
|
Conformational Changes in Orotidine 5′-Monophosphate
Decarboxylase: “Remote” Residues that Stabilize the Active
Conformation†
B. McKay Wood‡, Tina L. Amyes§, Alexander A. Fedorov∥, Elena V. Fedorov∥, Andrew
Shabila‡, Steven C. Almo∥, John P. Richard§, and John A. Gerlt*,‡
‡Departments of Biochemistry and Chemistry, University of Illinois at Urbana-Champaign
§Department of Chemistry, University at Buffalo, Buffalo, NY 14260
∥Department of Biochemistry, Albert Einstein College of Medicine, Bronx, NY 10461
Abstract
The structural factors responsible for the extraordinary rate enhancement (~1017) of the reaction
catalyzed by orotidine 5′-monophosphate decarboxylase (OMPDC) have not been defined.
Catalysis requires a conformational change that closes an active site loop and “clamps” the orotate
base proximal to hydrogen-bonded networks that destabilize the substrate and stabilize the
intermediate. In the OMPDC from Methanobacter thermoautotrophicus, a “remote” structurally
conserved cluster of hydrophobic residues that includes Val 182 in the active site loop is
assembled in the closed, catalytically active conformation. Substitution of these residues with Ala
decreases kcat/Km with a minimal effect on kcat, providing evidence that the cluster stabilizes the
closed conformation. The intrinsic binding energies of the 5′-phosphate group of OMP for the
mutant enzymes are similar to that for the wild type, supporting this conclusion.
Orotidine 5′-monophosphate decarboxylase (OMPDC1) is one of Nature’s best catalysts: the
reaction occurs with a rate enhancement of ~1017 and a proficiency of ~1023 M (2). The
reaction coordinate includes a vinyl carbanion intermediate (Scheme 1) (3, 4).
Structures of OMPDCs complexed with UMP or 6-hydroxyUMP, a n intermediate analog
(5-9), reveal hydrogen-bonded networks proximal to 1) C6 of the pyrimidine (Asp 70-Lys
72-Asp 75 in the OMPDC from Methanobacter thermoautotrophicus; MtOMPDC); and 2)
O2, N3, and O4 of the base (Ser 127-Gln 185). These destabilize the substrate (9) and
stabilize the intermediate, although the structural strategy for the latter is uncertain.
The 5′-phosphate group binds in a conserved motif at the ends of the seventh and eighth β-
strands of the (β/α)8-barrel structure, with interactions to backbone NH groups as well as the
guanidinium group of a conserved Arg (Arg 203 in MtOMPDC). These interactions are
important for catalysis: 1) kcat/Km for OMP exceeds that for orotidine by a factor of ~1011
for the OMPDC from Saccharomyces cerevisiae (ScOMPDC) (10); 2) kcat/Km for OMP
†This work was supported by NIH Grants GM039754 to J.P.R. and GM065155 to J.A.G.
*To whom correspondence should be addressed: J.A.G.: telephone, (217) 244-7414; fax: (217) 244-6538; j-gerlt@uiuc.edu..
SUPPORTING INFORMATION AVAILABLE
Descriptions of the experimental procedures. This material is available free of charge via the Internet at http://pubs.acs.org.
1Abbreviations: OMP, orotidine 5′-monophosphate; OMPDC, OMP decarboxylase; MtOMPDC, OMPDC from Methanobacter
thermoautotrophicus; ScOMPDC, OMPDC from Saccharomyces cerevisiae; EO, 1-(β-D-erythrofuranosyl)-orotic acid; HPi,
phosphate dianion; IBE, intrinsic binding energy.
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Published in final edited form as:
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exceeds that for 1-(β-D-erythrofuranosyl)orotic acid (EO; 5′-truncated OMP analog) by
factors of 5.2 × 108 and 3.6 × 108 for ScOMPDC and MtOMPDC, respectively (11, 12); and
3 ) phosphite dianion ( H Pi) activates decarboxylation of EO by factors of 5.6 × 105 M−1
and 2.9 × 105 M−1 for ScOMPDC and MtOMPDC, respectively (11, 12). The “intrinsic
binding energy” [IBE; (13)] of the 5′-phosphate/HPi 1) increases the affinity for the
substrate (e.g., OMP vs. orotidine/EO); and 2) enables decarboxylation by juxtaposition of
the substrate with the active site hydrogen-bonded networks (substrate destabilization and
intermediate stabilization). How the IBE promotes catalysis is unknown but required to
understand the structural basis for the rate enhancement.
A loop located at the end of the seventh β-strand closes over the active site when OMP binds
(Figure 1). Although the active site loops differ in both length and sequence in divergent
OMPDCs (12), each includes a spatially conserved Gln (Gln 185 in MtOMPDC) hydrogen-
bonded to both the 5′-phosphate and O2 of the pyrimidine as well as a conserved Ser at the
end of the fifth β-strand that also is hydrogen-bonded to N3 of the pyrimidine (Ser 127). We
characterized the importance of these “clamp” residues in ScOMPDC (Gln 215-Ser 154)
using EO/HPi and confirmed that the 5′-phosphate/HPi “switch” is required to activate the
enzyme (14).
The structures of divergent OMPDCs reveal that OMP binding is always accompanied by a
conformational change (Figure 1). The most obvious component is closure of the active site
loop. However, the (β/α)8-barrel structure can be divided into two domains, one formed
from the second, third, fourth, and fifth β-strands (where the hydrogen-bonded Asp 70-Lys
72-Asp 75 motif and Ser 127 are located) and the second from the sixth, seventh, eighth, and
first β-strands (where the phosphate binding motif and the active site loop, including Gln
185, are located) (15). OMP binding reorients the domains, with the latter domain moving
toward the former, enforcing the orotate carboxylate group to be juxtaposed vis a vis Asp
70- Lys 72-Asp 75 and, also, allowing formation of the Ser 127-Gln 185 “clamp”. Thus, the
transition between the open and closed conformations is more complicated than “simple”
hinge motion of the loop on the rigid framework of the (β/α)8-barrel structure. In this Report
we identify “remote” residues involved in this conformational change and quantitate their
importance in promoting and stabilizing the catalytically competent form of the enzyme.
The active site loop of MtOMPDC, P180-G181-V182-G183-A184-Q185-G186-G187-D188,
is disordered in the absence of substrate but ordered and closed in its presence (Figure 1). In
the liganded structure two residues in the loop make contacts with the (β/α)8-barrel scaffold:
1) Gln 185 is hydrogen-bonded to Ser 127 (vide infra); and 2) Val 182 is embedded in a
hydrophobic cluster also formed by Ile 199, Val 201, and Ile 218. The conservation of this
hydrophobic cluster in all OMPDCs suggests its importance in a common catalytic strategy.
We probed this strategy by mutagenesis of these hydrophobic residues.
Ala substitutions for residues in the hydrophobic cluster cause substantial decreases in kcat/
Km for decarboxylation of OMP but have little effect on kcat (Table 1). We determined high
resolution X-ray structures (≤ 1.4 Å) for each mutant in the presence of 6-hydroxyUMP
(Figure 2). The liganded structures superimpose well with that of wild type, with only small
differences observed at the sites of the substitutions (panel A). The active sites are identical
to that of wild type (panel B), explaining the minimal impact on kcat.
The mutated residues are remote from the active site (Figures 1 and 2). Thus, the effects of
the substitutions on kcat/Km cannot be explained by altered direct interactions with the
substrate. Instead, the effects can be explained by decreased stabilities of the closed
conformation in which the substrate is destabilized (9) and the anionic intermediate is
stabilized. A consistent model (Scheme 2) is that an equilibrium of open (Eo) and closed
Wood et al.
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(Ec) conformations exists in the absence or presence of substrate (KC’ or KC; KC’ << 1). The
substitutions destabilize Ec (decrease KC and KC ’), so kcat/Km is decreased (the energy
difference between Eo + S and [Ec•S]‡ is increased). However, the invariance of kcat
establishes that the substitutions do not alter the reactivitiy of Ec•S (the energy difference
between Ec•S and [Ec•S]‡). The energy to form Ec•S from Eo + S is achieved from 1)
interactions of the 5′-phosphate group with its binding motif (IBE; vide infra), and 2) an
increased concentration of OMP to form Ec•S (KC/KS or, equivalently, KC’/KS’, although
the former is expected to be the relevant pathway).
We also used the two part EO•HPi substrate. The values of kcat/Km for EO are decreased
relative to that for wild type (Table 1); these can be explained by decreased populations of
Ec (Scheme 2), assuming that EO, without a 5′-phosphate group, is unable to promote the
transition from Eo to Ec. The values of kcat/Km for OMP and kcat/Km for EO allow
calculation of the IBE for the 5′-phosphate group of OMP (Table 1).
HPi activates the mutants as judged by the values of the third-order rate constant, (kcat/
Km)EO•HPi/Kd. The equivalent changes (ΔΔG‡) in kcat/Km for both OMP and EO and the
third-order rate constant indicate that all three measure the effects of the substitutions on the
values of KC.
The values of the IBEs for the 5′-phosphate group for the mutants are the same as that for
the wild type, establishing that decarboxylation in the Ec•S complex occurs with equivalent
amounts of ground state destabilization (9) and transition state stabilization (as also reflected
by the invariant values of kcat). The IBEs provide further support for the role of the “remote”
hydrophobic cluster in stabilizing Ec relative to Eo but do not directly participating in
catalysis.
Our experiments implicate a structurally conserved hydrophobic cluster, Val 182, Ile 199,
Val 201, and Ile 218 in MtOMPDC, in stabilizing the closed conformation required for
catalysis. Its identification provides evidence that structural elements distal from the active
site, in addition to the proximal active site loop that closes to “clamp” the substrate, are
required for OMPDC’s extraordinary catalytic efficiency and proficiency.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
REFERENCES
(1). Wood BM, Chan KK, Amyes TL, Richard JP, Gerlt JA. Biochemistry. 2009; 48:5510–5517.
[PubMed: 19435313]
(2). Radzicka A, Wolfenden R. Science. 1995; 267:90–93. [PubMed: 7809611]
(3). Toth K, Amyes TL, Wood BM, Chan K, Gerlt JA, Richard JP. J. Am. Chem. Soc. 2007;
129:12946–12947. [PubMed: 17918849]
(4). Amyes TL, Wood BM, Chan K, Gerlt JA, Richard JP. J. Am. Chem. Soc. 2008; 130:1574–1575.
[PubMed: 18186641]
(5). Miller BG, Hassell AM, Wolfenden R, Milburn MV, Short SA. Proc. Natl. Acad. Sci. U A. 2000;
97:2011–2016.
(6). Appleby TC, Kinsland C, Begley TP, Ealick SE. Proc atl Acad Sci U S A. 2000; 97:2005–2010.
(7). Harris P, Navarro Poulsen JC, Jensen KF, Larsen S. Biochemistry. 2000; 39:4217–4224.
[PubMed: 10757968]
(8). Wu N, Mo Y, Gao J, Pai EF. Proc. Natl. Acad. Sci. U S A. 2000; 97:2017–2022. [PubMed:
10681441]
Wood et al.
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(9). Chan KK, Wood BM, Fedorov AA, Fedorov EV, Imker HJ, Amyes TL, Richard JP, Almo SC,
Gerlt JA. Biochemistry. 2009; 48:5518–5531. [PubMed: 19435314]
(10). Sievers A, Wolfenden R. Bioorg. Chem. 2005; 33:45–52. [PubMed: 15668182]
(11). Amyes TL, Richard JP, Tait JJ. J. Am. Chem. Soc. 2005; 127:15708–15709. [PubMed:
16277505]
(12). Toth K, Amyes TL, Wood BM, Chan KK, Gerlt JA, Richard JP. Biochemistry. 2009; 48:8006–
8013. [PubMed: 19618917]
(13). Jencks WP. Adv. Enzymol. Rel. Areas. Mol. Biol. 1975; 43:219–410.
(14). Barnett SA, Amyes TL, Wood BM, Gerlt JA, Richard JP. Biochemistry. 2008; 47:7785–4487.
[PubMed: 18598058]
(15). Harris P, Poulsen JC, Jensen KF, Larsen S. J. Mol. Biol. 2002; 318:1019–1029. [PubMed:
12054799]
(16). Go MK, Amyes TL, Richard JP. Biochemistry. 2009; 48:5769–5778. [PubMed: 19425580]
Wood et al.
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Figure 1.
Superposition of the active site of wild type MtOMPDC in the absence (cyan; disordered
loop depicted with the dotted line ) and presence ( grey ) of 6-hydroxyUMP. The carbons of
6-hydroxyUMP are highlighted in yellow; the carbons of Val 182 and Arg 203 in the
liganded structure are highlighted in orange.
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Figure 2.
Superpositions of the 6-hydroxyUMP-liganded structures of wild type MtOMPDC and the
single mutants in the hydrophobic cluster. Panel A, hydrophobic cluster; panel B, active
sites.
Wood et al.
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Scheme 1.
Wood et al.
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Scheme 2.
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Table 1
Kinetic Constants for OMP, EO and EO/HPi and Intrinsic Binding Energies of the 5′-Phosphate Group of OMP at pH 7.1 and 25 °C.
MtOMPDC
kcat
OMP
s−1
kcat/Km
OMP
M−1 s−1
ΔΔG‡
kcal/mola
kcat/Km
EO
M−1 s−1
ΔΔG‡
kcal/mola
(kcat/Km)/K
D
EO•HPib
M−2 s−1
ΔΔG‡
kcal/mola
5′-Phosphate
IBEc
kcal/mol
Wild type
4.6
2.9 × 106
8.7 × 10−3
2500
11.6 d
V182A
3.4
1.4 × 105
1.8
1.3 × 10−3
1.1
190
1.5
10.9
I199A
3.9
9.1 × 105
0.7
1.9 × 10−3
0.9
980
0.6
11.8
V201A
4.0
9.5 × 105
0.7
3.1 × 10−3
0.6
690
0.8
11.5
I218A
3.3
2.8 × 105
1.4
2.3 × 10−3
0.8
340
1.2
11.0
V182A/I199A
3.1
4.9 × 104
2.4
3.9 × 10−4
1.8
81
2.0
11.0
V182A/V201A
2.5
4.9 × 104
2.4
5.0 × 10−4
1.7
30
2.6
10.9
aCalculated from the ratio of the second-order or third-order rate constants for the wild type and mutant enzyme.
bThird-order rate constant for reaction of EO/HPi.
cTransition state stabilization by the 5′-phosphate group of OMP, calculated from the ratio of the values of kcat/Km for OMP and EO. IBEs for phosphite dianion can be calculated from the ratio of (kcat/
Km)/KD for EOHPi and kcat/Km for EO.
Biochemistry. Author manuscript; available in PMC 2011 December 26.
|
3M5Y
|
Crystal structure of the mutant V182A,V201A of orotidine 5'-monophosphate decarboxylase from Methanobacterium thermoautotrophicum
|
Conformational Changes in Orotidine 5′-Monophosphate
Decarboxylase: “Remote” Residues that Stabilize the Active
Conformation†
B. McKay Wood‡, Tina L. Amyes§, Alexander A. Fedorov∥, Elena V. Fedorov∥, Andrew
Shabila‡, Steven C. Almo∥, John P. Richard§, and John A. Gerlt*,‡
‡Departments of Biochemistry and Chemistry, University of Illinois at Urbana-Champaign
§Department of Chemistry, University at Buffalo, Buffalo, NY 14260
∥Department of Biochemistry, Albert Einstein College of Medicine, Bronx, NY 10461
Abstract
The structural factors responsible for the extraordinary rate enhancement (~1017) of the reaction
catalyzed by orotidine 5′-monophosphate decarboxylase (OMPDC) have not been defined.
Catalysis requires a conformational change that closes an active site loop and “clamps” the orotate
base proximal to hydrogen-bonded networks that destabilize the substrate and stabilize the
intermediate. In the OMPDC from Methanobacter thermoautotrophicus, a “remote” structurally
conserved cluster of hydrophobic residues that includes Val 182 in the active site loop is
assembled in the closed, catalytically active conformation. Substitution of these residues with Ala
decreases kcat/Km with a minimal effect on kcat, providing evidence that the cluster stabilizes the
closed conformation. The intrinsic binding energies of the 5′-phosphate group of OMP for the
mutant enzymes are similar to that for the wild type, supporting this conclusion.
Orotidine 5′-monophosphate decarboxylase (OMPDC1) is one of Nature’s best catalysts: the
reaction occurs with a rate enhancement of ~1017 and a proficiency of ~1023 M (2). The
reaction coordinate includes a vinyl carbanion intermediate (Scheme 1) (3, 4).
Structures of OMPDCs complexed with UMP or 6-hydroxyUMP, a n intermediate analog
(5-9), reveal hydrogen-bonded networks proximal to 1) C6 of the pyrimidine (Asp 70-Lys
72-Asp 75 in the OMPDC from Methanobacter thermoautotrophicus; MtOMPDC); and 2)
O2, N3, and O4 of the base (Ser 127-Gln 185). These destabilize the substrate (9) and
stabilize the intermediate, although the structural strategy for the latter is uncertain.
The 5′-phosphate group binds in a conserved motif at the ends of the seventh and eighth β-
strands of the (β/α)8-barrel structure, with interactions to backbone NH groups as well as the
guanidinium group of a conserved Arg (Arg 203 in MtOMPDC). These interactions are
important for catalysis: 1) kcat/Km for OMP exceeds that for orotidine by a factor of ~1011
for the OMPDC from Saccharomyces cerevisiae (ScOMPDC) (10); 2) kcat/Km for OMP
†This work was supported by NIH Grants GM039754 to J.P.R. and GM065155 to J.A.G.
*To whom correspondence should be addressed: J.A.G.: telephone, (217) 244-7414; fax: (217) 244-6538; j-gerlt@uiuc.edu..
SUPPORTING INFORMATION AVAILABLE
Descriptions of the experimental procedures. This material is available free of charge via the Internet at http://pubs.acs.org.
1Abbreviations: OMP, orotidine 5′-monophosphate; OMPDC, OMP decarboxylase; MtOMPDC, OMPDC from Methanobacter
thermoautotrophicus; ScOMPDC, OMPDC from Saccharomyces cerevisiae; EO, 1-(β-D-erythrofuranosyl)-orotic acid; HPi,
phosphate dianion; IBE, intrinsic binding energy.
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 December 26.
Published in final edited form as:
Biochemistry. 2010 May 4; 49(17): 3514–3516. doi:10.1021/bi100443a.
NIH-PA Author Manuscript
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NIH-PA Author Manuscript
exceeds that for 1-(β-D-erythrofuranosyl)orotic acid (EO; 5′-truncated OMP analog) by
factors of 5.2 × 108 and 3.6 × 108 for ScOMPDC and MtOMPDC, respectively (11, 12); and
3 ) phosphite dianion ( H Pi) activates decarboxylation of EO by factors of 5.6 × 105 M−1
and 2.9 × 105 M−1 for ScOMPDC and MtOMPDC, respectively (11, 12). The “intrinsic
binding energy” [IBE; (13)] of the 5′-phosphate/HPi 1) increases the affinity for the
substrate (e.g., OMP vs. orotidine/EO); and 2) enables decarboxylation by juxtaposition of
the substrate with the active site hydrogen-bonded networks (substrate destabilization and
intermediate stabilization). How the IBE promotes catalysis is unknown but required to
understand the structural basis for the rate enhancement.
A loop located at the end of the seventh β-strand closes over the active site when OMP binds
(Figure 1). Although the active site loops differ in both length and sequence in divergent
OMPDCs (12), each includes a spatially conserved Gln (Gln 185 in MtOMPDC) hydrogen-
bonded to both the 5′-phosphate and O2 of the pyrimidine as well as a conserved Ser at the
end of the fifth β-strand that also is hydrogen-bonded to N3 of the pyrimidine (Ser 127). We
characterized the importance of these “clamp” residues in ScOMPDC (Gln 215-Ser 154)
using EO/HPi and confirmed that the 5′-phosphate/HPi “switch” is required to activate the
enzyme (14).
The structures of divergent OMPDCs reveal that OMP binding is always accompanied by a
conformational change (Figure 1). The most obvious component is closure of the active site
loop. However, the (β/α)8-barrel structure can be divided into two domains, one formed
from the second, third, fourth, and fifth β-strands (where the hydrogen-bonded Asp 70-Lys
72-Asp 75 motif and Ser 127 are located) and the second from the sixth, seventh, eighth, and
first β-strands (where the phosphate binding motif and the active site loop, including Gln
185, are located) (15). OMP binding reorients the domains, with the latter domain moving
toward the former, enforcing the orotate carboxylate group to be juxtaposed vis a vis Asp
70- Lys 72-Asp 75 and, also, allowing formation of the Ser 127-Gln 185 “clamp”. Thus, the
transition between the open and closed conformations is more complicated than “simple”
hinge motion of the loop on the rigid framework of the (β/α)8-barrel structure. In this Report
we identify “remote” residues involved in this conformational change and quantitate their
importance in promoting and stabilizing the catalytically competent form of the enzyme.
The active site loop of MtOMPDC, P180-G181-V182-G183-A184-Q185-G186-G187-D188,
is disordered in the absence of substrate but ordered and closed in its presence (Figure 1). In
the liganded structure two residues in the loop make contacts with the (β/α)8-barrel scaffold:
1) Gln 185 is hydrogen-bonded to Ser 127 (vide infra); and 2) Val 182 is embedded in a
hydrophobic cluster also formed by Ile 199, Val 201, and Ile 218. The conservation of this
hydrophobic cluster in all OMPDCs suggests its importance in a common catalytic strategy.
We probed this strategy by mutagenesis of these hydrophobic residues.
Ala substitutions for residues in the hydrophobic cluster cause substantial decreases in kcat/
Km for decarboxylation of OMP but have little effect on kcat (Table 1). We determined high
resolution X-ray structures (≤ 1.4 Å) for each mutant in the presence of 6-hydroxyUMP
(Figure 2). The liganded structures superimpose well with that of wild type, with only small
differences observed at the sites of the substitutions (panel A). The active sites are identical
to that of wild type (panel B), explaining the minimal impact on kcat.
The mutated residues are remote from the active site (Figures 1 and 2). Thus, the effects of
the substitutions on kcat/Km cannot be explained by altered direct interactions with the
substrate. Instead, the effects can be explained by decreased stabilities of the closed
conformation in which the substrate is destabilized (9) and the anionic intermediate is
stabilized. A consistent model (Scheme 2) is that an equilibrium of open (Eo) and closed
Wood et al.
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(Ec) conformations exists in the absence or presence of substrate (KC’ or KC; KC’ << 1). The
substitutions destabilize Ec (decrease KC and KC ’), so kcat/Km is decreased (the energy
difference between Eo + S and [Ec•S]‡ is increased). However, the invariance of kcat
establishes that the substitutions do not alter the reactivitiy of Ec•S (the energy difference
between Ec•S and [Ec•S]‡). The energy to form Ec•S from Eo + S is achieved from 1)
interactions of the 5′-phosphate group with its binding motif (IBE; vide infra), and 2) an
increased concentration of OMP to form Ec•S (KC/KS or, equivalently, KC’/KS’, although
the former is expected to be the relevant pathway).
We also used the two part EO•HPi substrate. The values of kcat/Km for EO are decreased
relative to that for wild type (Table 1); these can be explained by decreased populations of
Ec (Scheme 2), assuming that EO, without a 5′-phosphate group, is unable to promote the
transition from Eo to Ec. The values of kcat/Km for OMP and kcat/Km for EO allow
calculation of the IBE for the 5′-phosphate group of OMP (Table 1).
HPi activates the mutants as judged by the values of the third-order rate constant, (kcat/
Km)EO•HPi/Kd. The equivalent changes (ΔΔG‡) in kcat/Km for both OMP and EO and the
third-order rate constant indicate that all three measure the effects of the substitutions on the
values of KC.
The values of the IBEs for the 5′-phosphate group for the mutants are the same as that for
the wild type, establishing that decarboxylation in the Ec•S complex occurs with equivalent
amounts of ground state destabilization (9) and transition state stabilization (as also reflected
by the invariant values of kcat). The IBEs provide further support for the role of the “remote”
hydrophobic cluster in stabilizing Ec relative to Eo but do not directly participating in
catalysis.
Our experiments implicate a structurally conserved hydrophobic cluster, Val 182, Ile 199,
Val 201, and Ile 218 in MtOMPDC, in stabilizing the closed conformation required for
catalysis. Its identification provides evidence that structural elements distal from the active
site, in addition to the proximal active site loop that closes to “clamp” the substrate, are
required for OMPDC’s extraordinary catalytic efficiency and proficiency.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
REFERENCES
(1). Wood BM, Chan KK, Amyes TL, Richard JP, Gerlt JA. Biochemistry. 2009; 48:5510–5517.
[PubMed: 19435313]
(2). Radzicka A, Wolfenden R. Science. 1995; 267:90–93. [PubMed: 7809611]
(3). Toth K, Amyes TL, Wood BM, Chan K, Gerlt JA, Richard JP. J. Am. Chem. Soc. 2007;
129:12946–12947. [PubMed: 17918849]
(4). Amyes TL, Wood BM, Chan K, Gerlt JA, Richard JP. J. Am. Chem. Soc. 2008; 130:1574–1575.
[PubMed: 18186641]
(5). Miller BG, Hassell AM, Wolfenden R, Milburn MV, Short SA. Proc. Natl. Acad. Sci. U A. 2000;
97:2011–2016.
(6). Appleby TC, Kinsland C, Begley TP, Ealick SE. Proc atl Acad Sci U S A. 2000; 97:2005–2010.
(7). Harris P, Navarro Poulsen JC, Jensen KF, Larsen S. Biochemistry. 2000; 39:4217–4224.
[PubMed: 10757968]
(8). Wu N, Mo Y, Gao J, Pai EF. Proc. Natl. Acad. Sci. U S A. 2000; 97:2017–2022. [PubMed:
10681441]
Wood et al.
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(9). Chan KK, Wood BM, Fedorov AA, Fedorov EV, Imker HJ, Amyes TL, Richard JP, Almo SC,
Gerlt JA. Biochemistry. 2009; 48:5518–5531. [PubMed: 19435314]
(10). Sievers A, Wolfenden R. Bioorg. Chem. 2005; 33:45–52. [PubMed: 15668182]
(11). Amyes TL, Richard JP, Tait JJ. J. Am. Chem. Soc. 2005; 127:15708–15709. [PubMed:
16277505]
(12). Toth K, Amyes TL, Wood BM, Chan KK, Gerlt JA, Richard JP. Biochemistry. 2009; 48:8006–
8013. [PubMed: 19618917]
(13). Jencks WP. Adv. Enzymol. Rel. Areas. Mol. Biol. 1975; 43:219–410.
(14). Barnett SA, Amyes TL, Wood BM, Gerlt JA, Richard JP. Biochemistry. 2008; 47:7785–4487.
[PubMed: 18598058]
(15). Harris P, Poulsen JC, Jensen KF, Larsen S. J. Mol. Biol. 2002; 318:1019–1029. [PubMed:
12054799]
(16). Go MK, Amyes TL, Richard JP. Biochemistry. 2009; 48:5769–5778. [PubMed: 19425580]
Wood et al.
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Figure 1.
Superposition of the active site of wild type MtOMPDC in the absence (cyan; disordered
loop depicted with the dotted line ) and presence ( grey ) of 6-hydroxyUMP. The carbons of
6-hydroxyUMP are highlighted in yellow; the carbons of Val 182 and Arg 203 in the
liganded structure are highlighted in orange.
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Figure 2.
Superpositions of the 6-hydroxyUMP-liganded structures of wild type MtOMPDC and the
single mutants in the hydrophobic cluster. Panel A, hydrophobic cluster; panel B, active
sites.
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Scheme 1.
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Scheme 2.
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Table 1
Kinetic Constants for OMP, EO and EO/HPi and Intrinsic Binding Energies of the 5′-Phosphate Group of OMP at pH 7.1 and 25 °C.
MtOMPDC
kcat
OMP
s−1
kcat/Km
OMP
M−1 s−1
ΔΔG‡
kcal/mola
kcat/Km
EO
M−1 s−1
ΔΔG‡
kcal/mola
(kcat/Km)/K
D
EO•HPib
M−2 s−1
ΔΔG‡
kcal/mola
5′-Phosphate
IBEc
kcal/mol
Wild type
4.6
2.9 × 106
8.7 × 10−3
2500
11.6 d
V182A
3.4
1.4 × 105
1.8
1.3 × 10−3
1.1
190
1.5
10.9
I199A
3.9
9.1 × 105
0.7
1.9 × 10−3
0.9
980
0.6
11.8
V201A
4.0
9.5 × 105
0.7
3.1 × 10−3
0.6
690
0.8
11.5
I218A
3.3
2.8 × 105
1.4
2.3 × 10−3
0.8
340
1.2
11.0
V182A/I199A
3.1
4.9 × 104
2.4
3.9 × 10−4
1.8
81
2.0
11.0
V182A/V201A
2.5
4.9 × 104
2.4
5.0 × 10−4
1.7
30
2.6
10.9
aCalculated from the ratio of the second-order or third-order rate constants for the wild type and mutant enzyme.
bThird-order rate constant for reaction of EO/HPi.
cTransition state stabilization by the 5′-phosphate group of OMP, calculated from the ratio of the values of kcat/Km for OMP and EO. IBEs for phosphite dianion can be calculated from the ratio of (kcat/
Km)/KD for EOHPi and kcat/Km for EO.
Biochemistry. Author manuscript; available in PMC 2011 December 26.
|
3M5Z
|
Crystal structure of the mutant V182A,I218A of orotidine 5'-monophosphate decarboxylase from Methanobacterium thermoautotrophicum
|
Conformational Changes in Orotidine 5′-Monophosphate
Decarboxylase: “Remote” Residues that Stabilize the Active
Conformation†
B. McKay Wood‡, Tina L. Amyes§, Alexander A. Fedorov∥, Elena V. Fedorov∥, Andrew
Shabila‡, Steven C. Almo∥, John P. Richard§, and John A. Gerlt*,‡
‡Departments of Biochemistry and Chemistry, University of Illinois at Urbana-Champaign
§Department of Chemistry, University at Buffalo, Buffalo, NY 14260
∥Department of Biochemistry, Albert Einstein College of Medicine, Bronx, NY 10461
Abstract
The structural factors responsible for the extraordinary rate enhancement (~1017) of the reaction
catalyzed by orotidine 5′-monophosphate decarboxylase (OMPDC) have not been defined.
Catalysis requires a conformational change that closes an active site loop and “clamps” the orotate
base proximal to hydrogen-bonded networks that destabilize the substrate and stabilize the
intermediate. In the OMPDC from Methanobacter thermoautotrophicus, a “remote” structurally
conserved cluster of hydrophobic residues that includes Val 182 in the active site loop is
assembled in the closed, catalytically active conformation. Substitution of these residues with Ala
decreases kcat/Km with a minimal effect on kcat, providing evidence that the cluster stabilizes the
closed conformation. The intrinsic binding energies of the 5′-phosphate group of OMP for the
mutant enzymes are similar to that for the wild type, supporting this conclusion.
Orotidine 5′-monophosphate decarboxylase (OMPDC1) is one of Nature’s best catalysts: the
reaction occurs with a rate enhancement of ~1017 and a proficiency of ~1023 M (2). The
reaction coordinate includes a vinyl carbanion intermediate (Scheme 1) (3, 4).
Structures of OMPDCs complexed with UMP or 6-hydroxyUMP, a n intermediate analog
(5-9), reveal hydrogen-bonded networks proximal to 1) C6 of the pyrimidine (Asp 70-Lys
72-Asp 75 in the OMPDC from Methanobacter thermoautotrophicus; MtOMPDC); and 2)
O2, N3, and O4 of the base (Ser 127-Gln 185). These destabilize the substrate (9) and
stabilize the intermediate, although the structural strategy for the latter is uncertain.
The 5′-phosphate group binds in a conserved motif at the ends of the seventh and eighth β-
strands of the (β/α)8-barrel structure, with interactions to backbone NH groups as well as the
guanidinium group of a conserved Arg (Arg 203 in MtOMPDC). These interactions are
important for catalysis: 1) kcat/Km for OMP exceeds that for orotidine by a factor of ~1011
for the OMPDC from Saccharomyces cerevisiae (ScOMPDC) (10); 2) kcat/Km for OMP
†This work was supported by NIH Grants GM039754 to J.P.R. and GM065155 to J.A.G.
*To whom correspondence should be addressed: J.A.G.: telephone, (217) 244-7414; fax: (217) 244-6538; j-gerlt@uiuc.edu..
SUPPORTING INFORMATION AVAILABLE
Descriptions of the experimental procedures. This material is available free of charge via the Internet at http://pubs.acs.org.
1Abbreviations: OMP, orotidine 5′-monophosphate; OMPDC, OMP decarboxylase; MtOMPDC, OMPDC from Methanobacter
thermoautotrophicus; ScOMPDC, OMPDC from Saccharomyces cerevisiae; EO, 1-(β-D-erythrofuranosyl)-orotic acid; HPi,
phosphate dianion; IBE, intrinsic binding energy.
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exceeds that for 1-(β-D-erythrofuranosyl)orotic acid (EO; 5′-truncated OMP analog) by
factors of 5.2 × 108 and 3.6 × 108 for ScOMPDC and MtOMPDC, respectively (11, 12); and
3 ) phosphite dianion ( H Pi) activates decarboxylation of EO by factors of 5.6 × 105 M−1
and 2.9 × 105 M−1 for ScOMPDC and MtOMPDC, respectively (11, 12). The “intrinsic
binding energy” [IBE; (13)] of the 5′-phosphate/HPi 1) increases the affinity for the
substrate (e.g., OMP vs. orotidine/EO); and 2) enables decarboxylation by juxtaposition of
the substrate with the active site hydrogen-bonded networks (substrate destabilization and
intermediate stabilization). How the IBE promotes catalysis is unknown but required to
understand the structural basis for the rate enhancement.
A loop located at the end of the seventh β-strand closes over the active site when OMP binds
(Figure 1). Although the active site loops differ in both length and sequence in divergent
OMPDCs (12), each includes a spatially conserved Gln (Gln 185 in MtOMPDC) hydrogen-
bonded to both the 5′-phosphate and O2 of the pyrimidine as well as a conserved Ser at the
end of the fifth β-strand that also is hydrogen-bonded to N3 of the pyrimidine (Ser 127). We
characterized the importance of these “clamp” residues in ScOMPDC (Gln 215-Ser 154)
using EO/HPi and confirmed that the 5′-phosphate/HPi “switch” is required to activate the
enzyme (14).
The structures of divergent OMPDCs reveal that OMP binding is always accompanied by a
conformational change (Figure 1). The most obvious component is closure of the active site
loop. However, the (β/α)8-barrel structure can be divided into two domains, one formed
from the second, third, fourth, and fifth β-strands (where the hydrogen-bonded Asp 70-Lys
72-Asp 75 motif and Ser 127 are located) and the second from the sixth, seventh, eighth, and
first β-strands (where the phosphate binding motif and the active site loop, including Gln
185, are located) (15). OMP binding reorients the domains, with the latter domain moving
toward the former, enforcing the orotate carboxylate group to be juxtaposed vis a vis Asp
70- Lys 72-Asp 75 and, also, allowing formation of the Ser 127-Gln 185 “clamp”. Thus, the
transition between the open and closed conformations is more complicated than “simple”
hinge motion of the loop on the rigid framework of the (β/α)8-barrel structure. In this Report
we identify “remote” residues involved in this conformational change and quantitate their
importance in promoting and stabilizing the catalytically competent form of the enzyme.
The active site loop of MtOMPDC, P180-G181-V182-G183-A184-Q185-G186-G187-D188,
is disordered in the absence of substrate but ordered and closed in its presence (Figure 1). In
the liganded structure two residues in the loop make contacts with the (β/α)8-barrel scaffold:
1) Gln 185 is hydrogen-bonded to Ser 127 (vide infra); and 2) Val 182 is embedded in a
hydrophobic cluster also formed by Ile 199, Val 201, and Ile 218. The conservation of this
hydrophobic cluster in all OMPDCs suggests its importance in a common catalytic strategy.
We probed this strategy by mutagenesis of these hydrophobic residues.
Ala substitutions for residues in the hydrophobic cluster cause substantial decreases in kcat/
Km for decarboxylation of OMP but have little effect on kcat (Table 1). We determined high
resolution X-ray structures (≤ 1.4 Å) for each mutant in the presence of 6-hydroxyUMP
(Figure 2). The liganded structures superimpose well with that of wild type, with only small
differences observed at the sites of the substitutions (panel A). The active sites are identical
to that of wild type (panel B), explaining the minimal impact on kcat.
The mutated residues are remote from the active site (Figures 1 and 2). Thus, the effects of
the substitutions on kcat/Km cannot be explained by altered direct interactions with the
substrate. Instead, the effects can be explained by decreased stabilities of the closed
conformation in which the substrate is destabilized (9) and the anionic intermediate is
stabilized. A consistent model (Scheme 2) is that an equilibrium of open (Eo) and closed
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(Ec) conformations exists in the absence or presence of substrate (KC’ or KC; KC’ << 1). The
substitutions destabilize Ec (decrease KC and KC ’), so kcat/Km is decreased (the energy
difference between Eo + S and [Ec•S]‡ is increased). However, the invariance of kcat
establishes that the substitutions do not alter the reactivitiy of Ec•S (the energy difference
between Ec•S and [Ec•S]‡). The energy to form Ec•S from Eo + S is achieved from 1)
interactions of the 5′-phosphate group with its binding motif (IBE; vide infra), and 2) an
increased concentration of OMP to form Ec•S (KC/KS or, equivalently, KC’/KS’, although
the former is expected to be the relevant pathway).
We also used the two part EO•HPi substrate. The values of kcat/Km for EO are decreased
relative to that for wild type (Table 1); these can be explained by decreased populations of
Ec (Scheme 2), assuming that EO, without a 5′-phosphate group, is unable to promote the
transition from Eo to Ec. The values of kcat/Km for OMP and kcat/Km for EO allow
calculation of the IBE for the 5′-phosphate group of OMP (Table 1).
HPi activates the mutants as judged by the values of the third-order rate constant, (kcat/
Km)EO•HPi/Kd. The equivalent changes (ΔΔG‡) in kcat/Km for both OMP and EO and the
third-order rate constant indicate that all three measure the effects of the substitutions on the
values of KC.
The values of the IBEs for the 5′-phosphate group for the mutants are the same as that for
the wild type, establishing that decarboxylation in the Ec•S complex occurs with equivalent
amounts of ground state destabilization (9) and transition state stabilization (as also reflected
by the invariant values of kcat). The IBEs provide further support for the role of the “remote”
hydrophobic cluster in stabilizing Ec relative to Eo but do not directly participating in
catalysis.
Our experiments implicate a structurally conserved hydrophobic cluster, Val 182, Ile 199,
Val 201, and Ile 218 in MtOMPDC, in stabilizing the closed conformation required for
catalysis. Its identification provides evidence that structural elements distal from the active
site, in addition to the proximal active site loop that closes to “clamp” the substrate, are
required for OMPDC’s extraordinary catalytic efficiency and proficiency.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
REFERENCES
(1). Wood BM, Chan KK, Amyes TL, Richard JP, Gerlt JA. Biochemistry. 2009; 48:5510–5517.
[PubMed: 19435313]
(2). Radzicka A, Wolfenden R. Science. 1995; 267:90–93. [PubMed: 7809611]
(3). Toth K, Amyes TL, Wood BM, Chan K, Gerlt JA, Richard JP. J. Am. Chem. Soc. 2007;
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(4). Amyes TL, Wood BM, Chan K, Gerlt JA, Richard JP. J. Am. Chem. Soc. 2008; 130:1574–1575.
[PubMed: 18186641]
(5). Miller BG, Hassell AM, Wolfenden R, Milburn MV, Short SA. Proc. Natl. Acad. Sci. U A. 2000;
97:2011–2016.
(6). Appleby TC, Kinsland C, Begley TP, Ealick SE. Proc atl Acad Sci U S A. 2000; 97:2005–2010.
(7). Harris P, Navarro Poulsen JC, Jensen KF, Larsen S. Biochemistry. 2000; 39:4217–4224.
[PubMed: 10757968]
(8). Wu N, Mo Y, Gao J, Pai EF. Proc. Natl. Acad. Sci. U S A. 2000; 97:2017–2022. [PubMed:
10681441]
Wood et al.
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(9). Chan KK, Wood BM, Fedorov AA, Fedorov EV, Imker HJ, Amyes TL, Richard JP, Almo SC,
Gerlt JA. Biochemistry. 2009; 48:5518–5531. [PubMed: 19435314]
(10). Sievers A, Wolfenden R. Bioorg. Chem. 2005; 33:45–52. [PubMed: 15668182]
(11). Amyes TL, Richard JP, Tait JJ. J. Am. Chem. Soc. 2005; 127:15708–15709. [PubMed:
16277505]
(12). Toth K, Amyes TL, Wood BM, Chan KK, Gerlt JA, Richard JP. Biochemistry. 2009; 48:8006–
8013. [PubMed: 19618917]
(13). Jencks WP. Adv. Enzymol. Rel. Areas. Mol. Biol. 1975; 43:219–410.
(14). Barnett SA, Amyes TL, Wood BM, Gerlt JA, Richard JP. Biochemistry. 2008; 47:7785–4487.
[PubMed: 18598058]
(15). Harris P, Poulsen JC, Jensen KF, Larsen S. J. Mol. Biol. 2002; 318:1019–1029. [PubMed:
12054799]
(16). Go MK, Amyes TL, Richard JP. Biochemistry. 2009; 48:5769–5778. [PubMed: 19425580]
Wood et al.
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Figure 1.
Superposition of the active site of wild type MtOMPDC in the absence (cyan; disordered
loop depicted with the dotted line ) and presence ( grey ) of 6-hydroxyUMP. The carbons of
6-hydroxyUMP are highlighted in yellow; the carbons of Val 182 and Arg 203 in the
liganded structure are highlighted in orange.
Wood et al.
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Figure 2.
Superpositions of the 6-hydroxyUMP-liganded structures of wild type MtOMPDC and the
single mutants in the hydrophobic cluster. Panel A, hydrophobic cluster; panel B, active
sites.
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Scheme 1.
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Scheme 2.
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Table 1
Kinetic Constants for OMP, EO and EO/HPi and Intrinsic Binding Energies of the 5′-Phosphate Group of OMP at pH 7.1 and 25 °C.
MtOMPDC
kcat
OMP
s−1
kcat/Km
OMP
M−1 s−1
ΔΔG‡
kcal/mola
kcat/Km
EO
M−1 s−1
ΔΔG‡
kcal/mola
(kcat/Km)/K
D
EO•HPib
M−2 s−1
ΔΔG‡
kcal/mola
5′-Phosphate
IBEc
kcal/mol
Wild type
4.6
2.9 × 106
8.7 × 10−3
2500
11.6 d
V182A
3.4
1.4 × 105
1.8
1.3 × 10−3
1.1
190
1.5
10.9
I199A
3.9
9.1 × 105
0.7
1.9 × 10−3
0.9
980
0.6
11.8
V201A
4.0
9.5 × 105
0.7
3.1 × 10−3
0.6
690
0.8
11.5
I218A
3.3
2.8 × 105
1.4
2.3 × 10−3
0.8
340
1.2
11.0
V182A/I199A
3.1
4.9 × 104
2.4
3.9 × 10−4
1.8
81
2.0
11.0
V182A/V201A
2.5
4.9 × 104
2.4
5.0 × 10−4
1.7
30
2.6
10.9
aCalculated from the ratio of the second-order or third-order rate constants for the wild type and mutant enzyme.
bThird-order rate constant for reaction of EO/HPi.
cTransition state stabilization by the 5′-phosphate group of OMP, calculated from the ratio of the values of kcat/Km for OMP and EO. IBEs for phosphite dianion can be calculated from the ratio of (kcat/
Km)/KD for EOHPi and kcat/Km for EO.
Biochemistry. Author manuscript; available in PMC 2011 December 26.
|
3M62
|
Crystal structure of Ufd2 in complex with the ubiquitin-like (UBL) domain of Rad23
|
The Yeast E4 Ubiquitin Ligase Ufd2 Interacts with the
Ubiquitin-like Domains of Rad23 and Dsk2 via a Novel and
Distinct Ubiquitin-like Binding Domain*□
S
Received for publication,February 12, 2010, and in revised form, March 22, 2010 Published, JBC Papers in Press,April 28, 2010, DOI 10.1074/jbc.M110.112532
Petra Ha¨nzelmann‡1, Julian Stingele§1, Kay Hofmann¶, Hermann Schindelin‡2, and Shahri Raasi§3
From the ‡Rudolf Virchow Center for Experimental Biomedicine, University of Wu¨rzburg, Josef-Schneider-Strasse 2,
97080 Wu¨rzburg, the §Laboratory of Cellular Biochemistry, Department of Biology, University of Konstanz, 78457 Konstanz,
and the ¶Bioinformatics Group, Miltenyi Biotec GmbH, Friedrich-Ebert-Strasse 68, 51429 Bergisch-Gladbach, Germany
Proteins containing ubiquitin-like (UBL) and ubiquitin-asso-
ciated (UBA) domains interact with various binding partners
and function as hubs during ubiquitin-mediated protein degra-
dation. A common interaction of the budding yeast UBL-UBA
proteins Rad23 and Dsk2 with the E4 ubiquitin ligase Ufd2 has
been described in endoplasmic reticulum-associated degrada-
tion among other pathways. The UBL domains of Rad23 and
Dsk2 play a prominent role in this process by interacting with
Ufd2 and different subunits of the 26 S proteasome. Here, we
report crystal structures of Ufd2 in complex with the UBL
domains of Rad23 and Dsk2. The N-terminal UBL-interacting
region of Ufd2 exhibits a unique sequence pattern, which is dis-
tinct from any known ubiquitin- or UBL-binding domain iden-
tified so far. Residue-specific differences exist in the interac-
tions of these UBL domains with Ufd2, which are coupled to
subtle differences in their binding affinities. The molecular
details of their differential interactions point to a role for adap-
tive evolution in shaping these interfaces.
The ubiquitin proteasome system regulates diverse cellular
processes including cell cycle progression, immune response,
neurodegenerative diseases, and protein quality control (1–4).
Ubiquitin-like (UBL)4 domains and ubiquitin- or UBL-binding
domains (UBD) (5) are small and highly diversified domains
that occur as integral parts of larger proteins (6–9). Integral
UBLs display a similar fold as ubiquitin (Ub) and like Ub are
described as protein-protein interaction modules without the
modifier function of Ub (5, 10). So far more than 20 different
classes of UBDs have been reported with a wide range of Ub
binding specificities (11, 12). The ubiquitin-associated (UBA)
domain was the first identified UBD, which exhibits the highest
representation of all UBDs in the eukaryotic genome (13) with
diverse Ub and Ub chain binding properties (14, 15). Although
the source of this binding diversity in vivo remained elusive so
far, remarkable structural studies have recently unraveled the
unique poly-Ub binding mode of a few other UBDs (16–20) and
contributed further to the understanding of how UBDs might
have acquired their respective ligand specificity.
UBL-UBA proteins contain both a UBL domain and at least
one UBA domain. Via these domains they interact simulta-
neously with ubiquitylated substrates and 26 S proteasome,
thereby delivering substrates to the proteasome for degradation
(21). Interestingly, UBL-UBA proteins are also binding partners
of other proteins (22–25). For instance, the budding yeast UBL-
UBA proteins Rad23 and Dsk2 can interact with the E4 ligase
Ufd2 via their UBL domains (22, 26, 27). A common involve-
ment of Ufd2, Rad23, and Dsk2 has been described in the endo-
plasmic reticulum-associated degradation, ubiquitin fusion
degradation, and OLE-1 gene induction pathway (22, 28–30),
where the UBL-Ufd2 interaction is indispensable. The associa-
tion of UBL-UBA proteins with Ub ligases, their reported sub-
strate specificity (31, 32), and the inhibitory effect of UBL-UBA
proteins on Ub chain disassembly (33, 34) support the idea that
UBL-UBA proteins might function as important regulatory and
specificity factors in Ub-mediated cellular proteolysis (21).
Therefore, understanding the binding behavior of the UBL
domains of UBL-UBA proteins with their various interacting
proteins will shed light on the regulatory role of these proteins.
Despite the identification of a large number of UBDs, structural
details of integral UBL-binding domains are limited. In some
cases, the intra- and intermolecular interactions between these
UBLs with known UBDs such as UBA or the ubiquitin-interact-
ing motif (UIM) have been demonstrated by solution NMR
(35–38).
Here, we are reporting crystal structures of budding yeast
Ufd2 in complex with the UBL domains of Rad23 and Dsk2 and
the molecular details of their interaction interfaces. We identify
a novel sequence pattern in the N-terminal UBL-binding region
of budding yeast Ufd2, which is conserved in lower eukaryotes
and is distinct from any known UBD identified so far. More-
over, despite engaging the same binding region, residue-spe-
* This work was supported by Deutsche Forschungsgemeinschaft Grant
RA1643/2-1 (to S. R.) and Rudolf Virchow Center for Experimental Biomed-
icine Grant FZ 82 (to H. S.).
The atomic coordinates and structure factors (codes 3M62 and 3M63) have been
deposited in the Protein Data Bank, Research Collaboratory for Structural
Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).
□
S The on-line version of this article (available at http://www.jbc.org) contains
supplemental Experimental Procedures, Tables S1 and S2, and Figs. S1–S5.
1 Both authors contributed equally to this work.
2 To whom correspondence may be addressed. E-mail: hermann.schindelin@
virchow.uni-wuerzburg.de.
3 To whom correspondence may be addressed. E-mail: shahri.raasi@
uni-konstanz.de.
4 The abbreviations used are: UBL, ubiquitin-like; UBA, ubiquitin-associated;
UIM, ubiquitin-interacting motif; UBD, ubiquitin-binding or ubiquitin-like
binding domain; UFD, ubiquitin fusion degradation; Ub, ubiquitin; GST,
glutathione S-transferase; ITC, isothermal titration calorimetry; SPR, sur-
face plasmon resonance; r.m.s., root mean square; WT, wild type; PDB, Pro-
tein Data Bank; h, human; Sc, S. cerevisiae; Sp, S. pombe.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 26, pp. 20390–20398, June 25, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
20390
JOURNAL OF BIOLOGICAL CHEMISTRY
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cific differences exist in the interactions of the UBL domains of
Rad23 and Dsk2 with Ufd2, which are coupled to subtle differ-
ences in their overall binding affinities. Mutational analyses of
the binding surface of the UBL domains and a closer inspection
of the thermodynamic contributions of those residues point to
adaptive evolution as a factor shaping these interfaces.
EXPERIMENTAL PROCEDURES
Cloning, site-directed mutagenesis, protein expression, and
purification
are
described
in
the
supplemental
Experi-
mental Procedures.
Crystallization of Ufd2Rad23-UBL and Ufd2Dsk2-UBL—
For crystallization of the Ufd2Rad23-UBL and Ufd2Dsk2-UBL
complexes, Ufd2 was incubated with Rad23-UBL or Dsk2-UBL
at a molar ratio of 1:1.5 (77 M Ufd2 and 115.5 M UBL) for 1 h
at 4 °C in the presence of 2 mM dithiothreitol. Crystals were
grown by vapor diffusion in hanging drops containing equal
volumes of protein in 50 mM HEPES, pH 7.4, 150 mM NaCl, and
2 mM dithiothreitol and a reservoir solution consisting of
16–18% (w/v) polyethylene glycol 3500 and 200 mM K3-citrate,
pH 8.3, equilibrated against the reservoir solution. Crystals
were cryo-protected by soaking in mother liquor containing
15–20% (v/v) glycerol. They belong to space group P212121 with
approximate cell dimensions of a 65 Å, b 126 Å, and c
181 Å with one complex per asymmetric unit.
Data Collection and Structure Determination—Crystals were
flash-cooled in liquid nitrogen, and data collection was per-
formed at 100 K. Data were collected at beamlines ID14–4
(European Synchrotron Radiation Facility (ESRF), Grenoble,
France) and BL 14.1 (Berliner Elektronenspeicherring-Gesell-
schaft fu¨r Synchrotronstrahlung (BESSY), Berlin, Germany)
and processed using Mosflm and Scala (39, 40). Data collection
statistics are summarized in supplemental Table S1. For subse-
quent calculations, the CCP4 suite was utilized (41) with excep-
tions as indicated. The Ufd2 structure was solved by molecular
replacement using Phaser (42) with Protein Data Bank (PDB)
entry 2QIZ as search model. Because Phaser could not find a
solution for the UBL domain with different search models, this
domain was fitted manually into the electron density using
human ubiquilin 3 (PDB entry 1YQB) for the Ufd2Rad23-UBL
complex and the Dsk2-UBL domain (PDB entry 2BWF) for the
Ufd2Dsk2-UBL complex as a model. The structures were
refined with Phenix (43) and REFMAC5 incorporating transla-
tion, libration, screw-rotation (TLS) refinement in all cycles
(44, 45). Solvent molecules were automatically added with Coot
(46). The figures were produced with PyMOL (65).
In Vitro Binding Assays—For pulldown assays, GST-tagged
Ufd2 and variants were immobilized on glutathione (GSH)
beads. In all experiments, 20 l of GSH beads were incubated
with 0.95 M purified Ufd2 in 400 l of phosphate-buffered
saline buffer with 1 mM dithiothreitol and 0.1% (v/v) Triton
X-100 at 4 °C for 1 h. WT-Ufd2 and GST alone were included as
controls. After centrifugation (1250 g, 30 s), beads were
washed five times with 400 l of binding buffer. Purified UBL
proteins (0.95 M) in a total volume of 400 l of binding buffer
were added to immobilized Ufd2 and treated in the same way as
in the first step. Immobilized proteins were analyzed by 17%
(v/v) SDS-PAGE or by immunoblotting with an anti-His
antibody.
Isothermal Titration Calorimetry (ITC)—Proteins were
extensively dialyzed against phosphate-buffered saline buffer
(pH 7.4, 1 mM -mercaptoethanol) followed by degassing. In all
experiments, 75–150 M Rad23- and Dsk2-UBL proteins were
titrated as the ligand into the sample cell containing 5–10 M
Ufd2. A volume of 10 l of ligand was added at a time with a
total number of 30 injections, resulting in a final molar ratio of
ligand-to-protein varying between 3:1 and 4:1. All experiments
were performed using a VP-ITC instrument (MicroCal, GE
Healthcare) at 25 °C. Buffer-to-buffer, buffer-to-Ufd2, as well
as Rad23-UBL/Dsk2-UBL-to-buffer titrations were performed
as described above. Corrected data were analyzed with a single-
site binding model using software supplied by the ITC manu-
facturer and non-linear least squares fitting to calculate the
dissociation constant (Kd).
Surface Plasmon Resonance (SPR) Measurements—SPR
binding assays were performed alternatively on BIAcore X or
BIAcore T100 instruments (GE Healthcare) at 25 °C in 10 mM
HEPES, pH 7.4, 150 mM NaCl, 50 M EDTA, 1 mM -mercap-
toethanol, and 0.005% (v/v) Surfactant P20. 100 response units
of His-tagged Rad23- or Dsk2-UBL were captured on a nickel-
nitrilotriacetic acid (Ni-NTA) sensor chip. GST-tagged Ufd2
for comparative binding assays and untagged Ufd2 for affin-
ity analysis were applied to the UBL surfaces in random
duplicates at a flow rate of 50 l/min. After each cycle, the
surface was regenerated using 350 mM EDTA in running
buffer to remove bound Ni2 and captured proteins. The
BIAcore T100 evaluation software was used to calculate the
steady state affinity constants. Data were plotted using
GraphPad Prism. For comparative assays, the relative bind-
ing responses of the mutants to WT proteins were deter-
mined by obtaining the maximum response for each interac-
tion at the end of each injection.
RESULTS
Ufd2 Binds the UBL Domains of Rad23 and Dsk2 with High
Affinity—Although Rad23 and Dsk2 interact with Ufd2 via
their UBL domains (22, 26), yeast two hybrid assays could only
identify the isolated N-terminal fragment (residues 1–380) of
Ufd2 as its UBL-interacting region (26). Additional details
regarding the Ufd2-UBL interactions have not been unraveled
so far. To further characterize the interactions of Ufd2 with the
UBLs of Rad23 and Dsk2, we performed GST pulldown assays
with GST-tagged full-length Ufd2 and C-terminally His-tagged
UBLs (Fig. 1A). Both UBLs were readily captured using immo-
bilized GST-Ufd2. In contrast, the UBL domain of Ddi1, a third
UBL-UBA protein, does not interact with Ufd2 (Fig. 1A) (22).
The differential binding of the Rad23- and Dsk2-UBLs to the
proteasomal subunits Rpn1 and Rpn10 has been described (25,
47, 48). Hence, we used SPR interaction analysis to search for
quantitative differences in their interactions. Steady state affin-
ity analysis of Ufd2 on both Rad23-UBL (Fig. 1B, left panel, and
1C) and Dsk2-UBL surfaces (Fig. 1B, right panel, and 1C) pro-
vided a Kd of 55 3 nM for the interaction of Rad23-UBL and a
lower affinity for Dsk2-UBL with a Kd of 418 56 nM.
UBL-binding Domain of Ufd2
JUNE 25, 2010•VOLUME 285•NUMBER 26
JOURNAL OF BIOLOGICAL CHEMISTRY 20391
The binding of the UBLs of Rad23 and Dsk2 to Ufd2 was also
analyzed by ITC to allow for a complete thermodynamic char-
acterization (Fig. 1D). These studies resulted in a Kd of 70 6
nM for the interaction of Rad23-UBL with Ufd2 and for the
binding of Dsk2-UBL to Ufd2 in a 2–3-fold higher Kd of 175
19 nM. Although there is an excellent agreement between SPR
and ITC for the Rad23-Ufd2 interaction, the two methods show
an 2-fold difference for the Dsk2-Ufd2 interaction. More
importantly, the enthalpic and entropic components to the free
energy are highly different between the two UBLs. The interac-
tion of Rad23-UBL and Ufd2 is more exothermic (H 17.3
kcal/mol) when compared with Dsk2-UBL (H 10.1 kcal/
mol). However, this is offset by a substantial decrease in entropy
for Rad23-UBL (TS 7.4 kcal/mol), whereas the entropic
contribution
is
minimal
for
the
Dsk2-UBL
interaction
(TS 0.8 kcal/mol).
Crystal Structures of Ufd2 in Complex with Rad23- and
Dsk2-UBL—We solved the structures of Ufd2 in complex with
Rad23-UBL carrying either an N-terminal or a C-terminal His
tag, which showed no significant structural differences. Due to
better data quality, the structure of Ufd2 with a C-terminal
His-tagged UBL is presented here. The Ufd2Rad23-UBL com-
plex was refined at 2.4 Å resolution to a crystallographic R-fac-
tor of 20.3% and a free R-factor of 25.7% (Table 1). As described
previously (49), Ufd2 is composed of an N-terminal variable
domain, a core domain, and a C-terminal U-box domain with a
fold similar to that of RING (really interesting new gene)
domains, which are present in certain Ub ligases (Fig. 2A).
Despite some conformational variability of the U-box domain,
our Ufd2 structure in the complex is quite similar (1.5 Å root
mean square (r.m.s.) deviation for 954 C atoms) to the pub-
lished Ufd2 structure (49).
The N-terminal variable region of Ufd2 that binds to the UBL
domain consists of eight -helices. Helices 1 to 4 are
arranged in a four-helix bundle, whereas helices 5 and 6
interact with 3 and 4 through hydrophobic contacts that are
partly mediated by their connecting loops (Fig. 2B). The struc-
ture of Rad23-UBL is comprised of a five-stranded -sheet, one
-helix, and one 310-helix (Fig. 2B). It displays a high degree of
similarity with Ub (PDB entry 1UBQ, 1.1 Å r.m.s. deviation for
72 C atoms, z-score 14, 25% sequence identity) and the UBL
domain of hHR23A (PDB entry 1P98, 1.6 Å r.m.s. deviation for
FIGURE 1. Interactions of Ufd2 with the UBL domains of Rad23 and Dsk2. A, GST-Ufd2 immobilized on GSH-beads was tested for binding to C-terminally
His-tagged UBLs of Rad23, Dsk2, and Ddi1. Captured UBLs were visualized by immunoblotting (WB) with an anti-His antibody. 2% of the input and GST beads
incubatedwithUBLswereloadedascontrols.B,aseriesof2-foldUfd2dilutions(233–3.6nM)wasappliedonaRad23-orDsk2-UBLsurfacefor120s(leftandright
panel,respectively).RU,responseunits.C,SPRbindingisothermsofWT-Rad23-andWT-Dsk2-UBLandthequintupleandseptupleDsk2-UBLvariantswithUfd2.
conc., concentration. D, ITC analysis of Ufd2Rad23-UBL (closed circles) and Ufd2Dsk2-UBL (open circles) complexes.
TABLE 1
Refinement statistics
Ufd2Rad23-UBL
Ufd2Dsk2-UBL
Resolution limit (Å)
45.2-2.4
73.5-2.4
No. of reflections
56,268
55,087
No. of protein/ligand/solvent atoms
8303/17/298
8288/17/182
Rcryst (Rfree)a,b
0.203 (0.257)
0.210 (0.270)
r.m.s. deviations in:
Bond lengths (Å)
0.016
0.015
Bond angles (°)
1.711
1.610
Estimated coordinate error (Å)
0.25
0.26
Overall average B-factor (Å2)
25.7
42.9
Ramachandran statistics (%)c
93.1/97.9/2.1
93.8/98.4/1.6
aRcryst hklFo Fc/hklFo where Fo and Fc are the observed and calculated
structure factor amplitudes.
bRfree, same as Rcryst for 5% of the data randomly omitted from the refinement. The
estimated coordinate error is based on Rfree.
c Ramachandran statistics indicate the fraction of residues in the favored (98%),
allowed ( 99.8%), and disallowed regions of the Ramachandran diagram, as
defined by MolProbity (64).
UBL-binding Domain of Ufd2
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72 C atoms, z-score 11.3, 26% sequence identity), one of the
two human homologs of budding yeast Rad23.
Subsequently, we solved the structure of Ufd2 with Dsk2-
UBL by molecular replacement. The UBL domain in the
Ufd2Dsk2-UBL complex exhibits increased flexibility; in par-
ticular, with a C-terminally tagged UBL domain, the first 30
amino acids of this domain were largely disordered (data not
shown). With an N-terminally tagged protein, the Ufd2Dsk2-
UBL structure was refined at 2.4 Å resolution to a crystallo-
graphic R-factor of 21.0% and a free R-factor of 27.0% (Table 1).
Both Rad23-UBL and Dsk2-UBL structures can be superim-
posed with an r.m.s. deviation of 1.1 Å for 71 aligned residues
(z-score 13.6, 30% sequence identity).
Analysis of the Ufd2Rad23-UBL Interface—The Ufd2UBL
interface in the structure of the complex buries a total molecu-
lar surface of about 1260 Å2, which is comprised to 590 Å2 of
the molecular surface of Ufd2 (1.3% of the total surface area)
and 670 Å2 from UBL (14.6% of the total surface area). This
interface is composed of almost equal parts of non-polar resi-
dues (38%), polar residues (33%), and charged residues (29%);
however, there are only one salt bridge (UBL-Lys-10 N–Ufd2-
Glu-49 O1 with a distance of 2.6 Å) and two direct hydrogen
bonds (UBL-Ser-47 O–Ufd2-Arg-92 N2, UBL-Gln-52 N2–
Ufd2-Glu-141 O at distances of 2.3 and 3.2 Å, respectively)
present (Fig. 3A).
Three UBL segments are contacting Ufd2 (Fig. 3A). Segment
I is located in the loop connecting -strands one and two, seg-
ment II involves -strands three and four, and segment III is
located in -strand five. Ufd2 residues from helix 2 and 4 as
well as the loop connecting 4 with 5 contribute to the
Ufd2UBL interface. These residues contact the hydrophobic
surface of the UBL -sheet in the region of -strands 3, 4, and 5.
Participating residues from Ufd2 include Leu-44, Tyr-97, Val-
100, and Trp-107, which are located in the hydrophobic UBL
pocket formed by residues Phe-9, Ile-45, Val-50, Val-69, and
Met-71 of Rad23 (Fig. 3, A and B).
For comparison, the principal recognition determinants in
Ub are: 1) the hydrophobic pocket formed by the side chains of
Leu-8 (Phe-9 in Rad23), Ile-44 (Ile-45 in Rad23), His-68 (Val-69
in Rad23), and Val-70 (Met-71 in Rad23) and 2) the main chain
amide group of Gly-47 (Gly-48 in Rad23), which is involved in
hydrogen bonding (50). Although the hydrophobic patch of
Rad23-UBL is also crucial for its interaction with Ufd2, the
main chain of Gly-48 does not form a hydrogen bond. Instead,
the -turn (Ser47–Gly48) connecting -strands 3 and 4 is stabi-
lized by the aforementioned strong hydrogen bond between
Ufd2-Arg-92 and UBL-Ser-47, whereas Ufd2-Gly-96 and Ufd2-
Tyr-97 contact UBL-Gly-48 (Fig. 3A). The aromatic ring of
Ufd2-Tyr-97 is involved in a stacking interaction with the pep-
tide bond between UBL residues 47 and 48 in this -turn.
Probing the Ufd2Rad23-UBL Interface—The importance of
interface residues was analyzed by mutagenesis experiments.
Eleven residues from Ufd2 and nine from Rad23-UBL were
each replaced with Ala. With the exception of the Rad23-UBL-
G48A variant that showed a reduced expression, all Ufd2 and
Rad23-UBL variants behaved like the WT protein during and
after purification, indicating that they were correctly folded
(data not shown). Initially, the contribution of these residues
was studied by GST pulldown and comparative SPR binding
assays (Table 2, supplemental Figs. S1 and S2A). In SPR studies,
the relative binding responses of mutants to WT proteins were
determined and compared. The majority of Rad23-UBL single
mutants revealed reduced binding to Ufd2 with Rad23-UBL-
I45A displaying the most prominent binding defect. The con-
tribution of the remaining residues to the interaction is aug-
mented in double mutants (supplemental Fig. S1C). Analysis of
the Ufd2 variants by SPR showed a largely reduced binding of
the residues located in the hydrophobic region of the UBL-
binding pocket (Leu-44, Tyr-97, Val-100, and Phe-107) and
Asp-40 (Table 2 and supplemental Fig. S2A).
ITC studies confirmed these results and allowed for a
quantitative analysis (Table 2, supplemental Fig. S3 and
supplemental Table S2). The most significant effect for Ufd2
was observed for all residues located in the hydrophobic UBL
pocket. Mutation of Val-100 and Phe-107 to Ala completely
abolished binding, the Y97A variant strongly reduced binding
(1900-fold), and the I104A and L44A variants showed signifi-
cantly decreased affinities (20- and 120-fold, respectively).
Although not directly involved in complex formation (Fig. 3A),
the Ufd2-D40A variant showed a 110-fold reduced affinity
(Table 2), which probably is the result of the missing intramo-
lecular hydrogen bond between Ufd2-Asp-40 and Ufd2-Tyr-97
(O2–OH 2.5 Å). This hydrogen bond seems to be crucial for
proper positioning of the aromatic side chain of Tyr-97 in the
interface region and might be important to align helices 2 and
4 for interaction with the Rad23-UBL.
FIGURE 2. Structure of Ufd2 in complex with the UBL domain of Rad23.
A, ribbon representation of the overall structure of the Ufd2Rad23-UBL com-
plex. The Rad23-UBL domain is shown in green, the N-terminal Ufd2 region is
in orange, the Ufd2 core domain is in gray, and the Ufd2 U-box domain is in
red. B, close-up view of the N-terminal Ufd2 domain in complex with Rad23-
UBL with secondary structural elements labeled and color-coded as in A.
UBL-binding Domain of Ufd2
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In Rad23-UBL, Ile-45 was shown to be integral for binding to
Ufd2 by the detrimental effect (130-fold decrease) after
exchange to Ala (Table 2). Mutation of Phe-9, Val-50, and Val-
69, residues adjacent to Ile-45 in the hydrophobic patch, also
decreased the affinity of Rad23-UBL 5–7-fold. Ser-47, which is
in hydrogen-bonding distance to Ufd2-Arg-92 and next to
UBL-Gly-48, showed a 9-fold reduced affinity. In Ub and in the
human Rad23 homolog hHR23A, Ser-47 is replaced by Ala.
Charged residues found in the interface (Ufd2, Glu-26, Glu-49,
and Arg-92; UBL, Lys-10) do not contribute significantly to the
interaction. In summary, our data indicate that the most prom-
inent contact between Ufd2 and Rad23-UBL is the strong
hydrophobic interaction between UBL-Ile-45 and Ufd2-Val-
100 as well as Ufd2-Phe-107, which defines the core of the UBL-
interacting region of Ufd2.
Molecular Discrimination between Rad23 and Dsk2—De-
spite a similar fold, the UBL domains of Rad23 and Dsk2 display
only 30% sequence identity, which could give rise to differences
in their interactions. A superposition of the bound Rad23-UBL
and Dsk2-UBL in the two complex structures showed signifi-
cant changes (Fig. 3C). Of the three UBL segments involved in
the Ufd2 interaction (Fig. 3A), segment II including Ile-45
(Ile-44 in Ub) is highly conserved, and there are no conforma-
tional changes in both UBL structures, whereas segments I and
III are not conserved and display structural changes (Fig. 3C).
The loop, connecting -strands one and two, adopts different
conformations, and -strand five shows a displacement that
might affect binding (Fig. 3C).
Segment I includes Phe-9 in Rad23-UBL, corresponding to
Leu-8 in Ub, where this residue is also involved in Ub recogni-
tion by UBDs (50, 51). Phe-9 is replaced by Gly-10 in Dsk2-UBL,
and there is no corresponding hydrophobic interacting residue
(supplemental Fig. S4A). Dsk2 residues Gly-10 and Gln-11
adopt different conformations when compared with Leu-8/
Thr-9 of Ub and Phe-9/Lys-10 of Rad23-UBL. In the
Ufd2Dsk2-UBL structure, the Ufd2Rad23-UBL salt bridge
(Lys-10/Glu-49) is missing due to the Lys-10 to Gln-11
exchange, with the latter side chain no longer being located in
the protein interface (supplemental Fig. S4A). The missing
interaction from segment I in Dsk2 might be compensated by
the displacement of -strand five toward Ufd2 and a replace-
ment of Val-69 to His-69 found in segment III resulting in a
more pronounced interaction in this region when compared
with Rad23-UBL (supplemental Fig. S4A). The presence of the
salt bridge seems to be the reason for the more exothermic
character of the Ufd2Rad23-UBL interaction, a view that is also
supported by the corresponding Ufd2-E49A and Rad23-K10A
variants, which both display binding enthalpies similar to the
FIGURE 3. The Ufd2Rad23-UBL interface. A, residues involved in binding are shown in stick representation. Carbon atoms of Ufd2 residues are colored in
orange and in green for Rad23-UBL. Dashed lines indicate H-bonds. B, structure-based sequence alignment of Rad23-UBL, Dsk2-UBL, hHR23A-UBL, and Ub.
Secondary structure elements of Rad23-UBL were assigned using DSSP (61) and are labeled above the sequences. The alignment was performed using DaliLite
(62), and the figure was prepared with ESPript (63). Strictly conserved amino acids are highlighted with a red background, and similar amino acids are shown as
redletters.ThethreeUfd2-bindingsegmentsareindicated.ResiduesinvolvedinUfd2Rad23-UBLinteractionarelabeledwithgreenstars.C,superpositionofthe
Ufd2Rad23-UBL/Dsk2-UBL complex structures with the N-terminal binding domain of Ufd2 in orange (Rad23 complex) and gray (Dsk2 complex), with Rad23-
UBL in green and Dsk2-UbL in yellow.
UBL-binding Domain of Ufd2
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Ufd2Dsk2-UBL complex (supplemental Table S2). At the same
time, the absence of the salt bridge in both mutants is accom-
panied by a more favorable entropic contribution, which is on a
level similar to the Ufd2Dsk2-UBL complex.
To identify residues important for the subtle molecular dis-
crimination between the UBL domains of Rad23 and Dsk2, the
interaction of Dsk2-UBL with Ufd2 mutants was analyzed by
GST pulldown assays (data not shown), SPR, and ITC (Table 2,
supplemental Figs. S2B and S3C). Quantitative ITC analysis
showed reduced binding of Dsk2 to Ufd2 mutants Y97A (470-
fold), V100A (22-fold), I104A (6-fold), and F107A (20-fold)
(Table 2). However, binding of the V100A and F107A variants is
not completely abolished, and when compared with Rad23-
UBL, the binding affinities are less affected by a factor of about
3–7 in most of the mutants analyzed. In addition, the L44A
mutant, which has a 120-fold reduced affinity with Rad23-UBL,
is only three times reduced in the case of Dsk2-UBL.
In general agreement with the ITC affinity data, the compar-
ative SPR binding assay revealed significant differences in the
association of Ufd2 variants Y97A, V100A, I104A, and F107A
with Rad23- and Dsk2-UBL surfaces (supplemental Fig. S4B).
The observed SPR decrease for the binding of the T48A and
E49A variants of Ufd2 to Dsk2-UBL seems to be compensated
by slower dissociations, thus explaining why these mutants
show no significant defect in the ITC analysis.
To further analyze the contribution of segments I and III to
complex formation, a G10F/Q11K/S67Q/H69V/V71M quintu-
ple Dsk2-UBL mutant was generated, where key residues in
binding segments I and III were replaced with the correspond-
ing residues from Rad23-UBL. Comparative binding as well as
steady state affinity analysis by SPR revealed only a small
increase (Kd 348 nM) in binding affinity for Ufd2 when com-
pared with WT-Dsk2-UBL (Kd 418 nM) (data not shown and
Fig. 1C). In addition, neither a crystal structure of the quintuple
Ufd2Dsk2-UBL complex (data not shown) nor the KD of 240
nM deduced by ITC revealed significant differences from
WT-Dsk2-UBL (Kd 175 nM). The ITC analysis did, however,
reveal that the binding is now driven by an increase in entropy
(TS 6.5 kcal/mol versus 0.8 and 7.4 kcal/mol for
WT-Dsk2-UBL and -Rad23-UBL, respectively), whereas the
binding enthalpy is reduced to only 2.5 kcal/mol when com-
pared with 10.1 and 17.3 kcal/mol (supplemental Table S2).
Interestingly, SPR and ITC analysis of a G10F/Q11K/I50V/
K52Q/S67Q/H69V/V71M septuple Dsk2-UBL mutant, which
has the additional I50V and K52Q substitutions in segment II,
showed an even lower affinity (SPR, Kd 648 nM; ITC, Kd 875
nM) to Ufd2 when compared with WT-Dsk2-UBL (Fig. 1C).
The N Terminus of Ufd2 Represents a Unique and Conserved
UBL-binding Domain—A multiple sequence alignment of Ufd2
from different yeast species displays a distinct pattern of con-
served residues involved in UBL binding (Fig. 4A). Among the
available yeast genomes, the Schizosaccharomyces pombe
sequence is most similar to those from higher eukaryotes; thus
we isolated cDNA fragments for the coding region of the UBL
domains of Rad23 and Dsk2 and full-length Ufd2 from this
organism and examined their interactions by GST pulldown
assays (Fig. 4B) as well as SPR (data not shown). We could show
that SpUfd2 interacts strongly with the UBL domains of
SpRad23 and SpDsk2 as well as with the UBL domains of
ScRad23 and ScDsk2 and vice versa. This cross species interac-
tion, despite the diversified UBL and Ufd2 amino acid
sequences, indicates that the identified sequence pattern repre-
sents a real UBL-interacting domain. A surface representation
of this motif is shown in Fig. 4C.
The N terminus of budding yeast Ufd2 displays only limited
sequence homology with the human Ufd2s, E4A and E4B
(supplemental Fig. S5) and other Ufd2s from higher eukaryotes.
In agreement with this finding, there are no reports that
hHR23A/B interacts with either of the human homologs of
Ufd2. Interestingly, our SPR studies showed that the UBL
domain of hHR23A interacts with ScUfd2, albeit with lower
affinity (data not shown). Apparently, the high affinity interac-
tion of the UBL domains of Rad23 and Dsk2 has been lost dur-
ing the evolution of this domain. The absence of conservation
of the Ufd2-UBL interface could potentially be used for thera-
peutic interventions against pathogenic yeasts such as Candida
albicans by designing low molecular weight compounds that
disrupt this interface. However, further functional studies in
pathogenic yeasts are required to examine the suitability of this
surface as a drug target.
TABLE 2
ITC and SPR parameters of Ufd2, Rad23-UBL, Dsk2-UBL, and variants
indicates no change; ND indicates not detected (corresponding to at least a
104-fold decrease in binding affinity).
Ufd2
WT-UBL
ITC
SPRa (% of
relative
response)
Kd
Fold decrease
nM
WT
Rad23
70
100
Dsk2
175
100
E26A
Rad23
284
4
91
Dsk2
521
3
83
D40A
Rad23
7900
110
20
Dsk2
7600
40
0
L44A
Rad23
8300
120
31
Dsk2
463
3
52
T48A
Rad23
72
70
Dsk2
296
2
29
E49A
Rad23
413
6
69
Dsk2
314
2
44
R92A
Rad23
265
4
76
Dsk2
128
59
G96A
Rad23
592
8
51
Dsk2
216
60
Y97A
Rad23
134,000
1900
3
Dsk2
83,000
470
0
V100A
Rad23
ND
10,000
9
Dsk2
3900
22
1
I104A
Rad23
1600
20
43
Dsk2
1100
6
12
F107A
Rad23
ND
10,000
11
Dsk2
3600
20
0
Ufd2
Rad23-UBL
ITC
SPRa (% of
relative
response)
Kd
Fold decrease
nM
WT
F9A
376
5
80
WT
K10A
162
2
96
WT
I45A
9100
130
17
WT
S47A
606
9
62
WT
V50A
441
6
88
WT
Q52A
415
6
79
WT
Q67A
113
2
92
WT
V69A
478
7
70
WT
M71A
221
3
88
a For comparative SPR assays, the relative binding responses of the mutants to wt
proteins were determined by obtaining the maximum response for each interac-
tion at the end of injection.
UBL-binding Domain of Ufd2
JUNE 25, 2010•VOLUME 285•NUMBER 26
JOURNAL OF BIOLOGICAL CHEMISTRY 20395
DISCUSSION
Families and superfamilies of protein domains or folds have
evolved through a process of homologous recombination and
gene duplication (52) followed by sequence divergence. Mem-
bers of different classes of UBDs such as UBA or UBL domains
result from these processes. For instance, the UBL domains of
Rad23 and Dsk2 display only 30% sequence identity but adopt
the same fold and utilize the same binding surface to recognize
a common UBL-binding domain of Ufd2 to form complexes
that display similarly high affinity. Nevertheless, not all inter-
acting residues are conserved; in particular, there is sequence
diversity in binding segments I and III of UBLs. Our attempts to
interconvert the UBL domains by altering non-conserved inter-
facial residues were not successful, thus suggesting that addi-
tional elements exist and play a role in the respective Ufd2-UBL
interaction. Interestingly, these results resemble earlier studies
on WW domains (53, 54), where a statistical analysis of multiple
sequence alignments was utilized to identify co-evolving resi-
dues. The authors demonstrated that not only interfacial resi-
dues but also buried residues distal to the interface co-evolved
with interfacial residues and contribute significantly to the
interactions. They concluded that certain sequence patterns in
interacting domains are due to adaptive evolution. In agree-
ment with these findings, our data prove that substitution of
key interfacial residues of Dsk2-UBL has no significant effect on
its overall binding affinity to Ufd2. In case of the septuple
mutant, we even observed a decrease in binding affinity, which
could be due to the imposed disorder into the evolutionary
inter-residue relations within the UBL fold. This is supported
by the fact that when compared with Dsk2-UBL and in partic-
ular Rad23-UBL, the binding of the quintuple Dsk2-UBL
mutant is driven strongly by entropy. These findings indicate
that binding interfaces can be modulated by changes in residues
that affect either the binding enthalpy or the entropy, thus pro-
viding additional freedom to maintain an interaction during the
course of evolution, an effect that has been described previously
as entropy/enthalpy compensation (55, 56).
Our studies suggest that UBL domains have co-evolved with
Ufd2 to reach optimal binding affinities by altering specific res-
idue-to-residue interactions (co-evolution at the residue level)
(57), while at the same time, all functional aspects of Rad23 or
Dsk2 are preserved. Therefore, the primary sequence degener-
FIGURE 4. The N terminus of Ufd2 represents a conserved UBL-interacting domain in lower eukaryotes. A, alignment of the N-terminal sequences of
fungal Ufd2s. Invariant or conserved residues with surface access are colored in dark blue, buried ones are in light blue. Residues labeled with red stars represent
the core region of the binding domain, which is essential for UBL interaction, whereas residues labeled with yellow stars contribute moderately to the
interaction. K. lactis, Kluyveromyces lactis; C. glabrata, Candida glabrata; Z. rouxii, Zygosaccharomyces rouxii; L. thermotolerans, Lachancea thermotolerans; C.
tropicalis, Candida tropicalis; C. dubliniensis, Candida dubliniensis; P. guilliermondii, Pichia guilliermondii; D. hansenii, Debaryomyces hansenii. B, GST pulldown
assay demonstrates the cross interactions of S. pombe and S. cerevisiae proteins. 5% of inputs and GST beads incubated with UBLs were loaded as controls. WB,
Western blot. C, surface representation of the N-terminal UBL-binding domain of Ufd2, color-coded as in A.
UBL-binding Domain of Ufd2
20396
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 26•JUNE 25, 2010
acy of protein domains such as UBAs or UBLs has been toler-
ated and established in favor of the cooperative nature of the
interactions and their functionality within their respective pro-
tein complexes. This further suggests that differential binding
properties observed for the interactions of Ufd2 with UBLs (this
study) or for the interactions of UBAs with Ub and Ub chains
(14) can arise not necessarily due to their interaction with dif-
ferent ligands but can also result from the adaptive co-evolution
of these domains with the same interacting partners. Seem-
ingly, these interfacial domains have evolved to hold protein-
protein interactions in a suitable form within multicomponent
complexes until they are challenged by downstream events.
Numerous structures of Ub receptors in complex with their
respective Ub/UBL-binding domains have been reported. The
so far characterized Ub receptors of the 26 S proteasome in
budding yeast encompass the two proteasomal subunits Rpn10
(S5a in humans) and Rpn13 and the three UBL-UBA proteins
Rad23, Dsk2, and Ddi1, which associate with the proteasome
and function as shuttle factors (21). Experimental evidence for
the existence of additional candidates exist (21, 58). Rad23 and
Dsk2 interact with the proteasomal subunit Rpn1 via their UBL
domains (21, 47). Aside from their known interactions with Ub,
Rpn13 and Rpn10/S5a alternatively interact with UBL-UBA
proteins (21, 35, 37, 38, 48, 51, 59). For instance, the preferential
association of Rpn1 with Rad23 and Rpn10 with Dsk2 has been
reported (25, 38, 47, 48). Based on the binding of hpLIC2 (Dsk2
homolog) with Rpn13, an interac-
tion of Dsk2 with Rpn13 has been
proposed (51, 59).
Although
the
aforementioned
examples engage essentially the
same
surface
of
Ub/UBL,
they
diverge in both structure and pat-
terns of Ub/UBL recognition (Fig.
5). For instance, hRpn10/S5a recog-
nizes the UBL domain of hHR23A,
one of the two human homologs of
Rad23, via a Ub-interacting motif,
which consists of a single -helix
(35, 37). Rpn13 binds Ub via a pleck-
strin homology domain, which is a
seven-stranded -sandwich capped
by an -helix (51). The Ub-binding
surface of Rpn13 is formed by
three loops that bridge -strands.
Another Ub-binding element is the
UBA domain found for example in
Dsk2 (60). The UBA domain is com-
posed of a three-helix bundle. With
the exception of Rpn13, which
exclusively binds via loops, it seems
that the majority of Ub/UBL-bind-
ing domains fold into -helical
structures
including
the
known
UBDs, UIM, and UBA, and the
UBL-binding domain of Ufd2 iden-
tified in this study. Despite the pre-
dominant
interaction
involving
-helices as Ub/UBL-binding elements, the three-dimensional
structure of the UBL-binding domain of Ufd2 differs from other
known examples, hence providing the first structural descrip-
tion for how Ufd2 acts as a UBL receptor while at the same time
further enhancing the diversity of UBDs in general.
Acknowledgments—We thank Martin Scheffner and Keith Wilkinson
for critical reading of the manuscript. We thank Stefan Jentsch for
providing the original plasmids for the expression of Rad23, Dsk2, and
Ufd2 and for Ufd2-specific antibodies used in the initial phase of this
study. We also thank David Fischer and Rodrigo Villasen˜or for the
contribution to this study and Sven Eiselein for providing us with
C-terminal GST-tagging plasmid.
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UBL-binding Domain of Ufd2
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VOLUME 285•NUMBER 26•JUNE 25, 2010
|
3M63
|
Crystal structure of Ufd2 in complex with the ubiquitin-like (UBL) domain of Dsk2
|
The Yeast E4 Ubiquitin Ligase Ufd2 Interacts with the
Ubiquitin-like Domains of Rad23 and Dsk2 via a Novel and
Distinct Ubiquitin-like Binding Domain*□
S
Received for publication,February 12, 2010, and in revised form, March 22, 2010 Published, JBC Papers in Press,April 28, 2010, DOI 10.1074/jbc.M110.112532
Petra Ha¨nzelmann‡1, Julian Stingele§1, Kay Hofmann¶, Hermann Schindelin‡2, and Shahri Raasi§3
From the ‡Rudolf Virchow Center for Experimental Biomedicine, University of Wu¨rzburg, Josef-Schneider-Strasse 2,
97080 Wu¨rzburg, the §Laboratory of Cellular Biochemistry, Department of Biology, University of Konstanz, 78457 Konstanz,
and the ¶Bioinformatics Group, Miltenyi Biotec GmbH, Friedrich-Ebert-Strasse 68, 51429 Bergisch-Gladbach, Germany
Proteins containing ubiquitin-like (UBL) and ubiquitin-asso-
ciated (UBA) domains interact with various binding partners
and function as hubs during ubiquitin-mediated protein degra-
dation. A common interaction of the budding yeast UBL-UBA
proteins Rad23 and Dsk2 with the E4 ubiquitin ligase Ufd2 has
been described in endoplasmic reticulum-associated degrada-
tion among other pathways. The UBL domains of Rad23 and
Dsk2 play a prominent role in this process by interacting with
Ufd2 and different subunits of the 26 S proteasome. Here, we
report crystal structures of Ufd2 in complex with the UBL
domains of Rad23 and Dsk2. The N-terminal UBL-interacting
region of Ufd2 exhibits a unique sequence pattern, which is dis-
tinct from any known ubiquitin- or UBL-binding domain iden-
tified so far. Residue-specific differences exist in the interac-
tions of these UBL domains with Ufd2, which are coupled to
subtle differences in their binding affinities. The molecular
details of their differential interactions point to a role for adap-
tive evolution in shaping these interfaces.
The ubiquitin proteasome system regulates diverse cellular
processes including cell cycle progression, immune response,
neurodegenerative diseases, and protein quality control (1–4).
Ubiquitin-like (UBL)4 domains and ubiquitin- or UBL-binding
domains (UBD) (5) are small and highly diversified domains
that occur as integral parts of larger proteins (6–9). Integral
UBLs display a similar fold as ubiquitin (Ub) and like Ub are
described as protein-protein interaction modules without the
modifier function of Ub (5, 10). So far more than 20 different
classes of UBDs have been reported with a wide range of Ub
binding specificities (11, 12). The ubiquitin-associated (UBA)
domain was the first identified UBD, which exhibits the highest
representation of all UBDs in the eukaryotic genome (13) with
diverse Ub and Ub chain binding properties (14, 15). Although
the source of this binding diversity in vivo remained elusive so
far, remarkable structural studies have recently unraveled the
unique poly-Ub binding mode of a few other UBDs (16–20) and
contributed further to the understanding of how UBDs might
have acquired their respective ligand specificity.
UBL-UBA proteins contain both a UBL domain and at least
one UBA domain. Via these domains they interact simulta-
neously with ubiquitylated substrates and 26 S proteasome,
thereby delivering substrates to the proteasome for degradation
(21). Interestingly, UBL-UBA proteins are also binding partners
of other proteins (22–25). For instance, the budding yeast UBL-
UBA proteins Rad23 and Dsk2 can interact with the E4 ligase
Ufd2 via their UBL domains (22, 26, 27). A common involve-
ment of Ufd2, Rad23, and Dsk2 has been described in the endo-
plasmic reticulum-associated degradation, ubiquitin fusion
degradation, and OLE-1 gene induction pathway (22, 28–30),
where the UBL-Ufd2 interaction is indispensable. The associa-
tion of UBL-UBA proteins with Ub ligases, their reported sub-
strate specificity (31, 32), and the inhibitory effect of UBL-UBA
proteins on Ub chain disassembly (33, 34) support the idea that
UBL-UBA proteins might function as important regulatory and
specificity factors in Ub-mediated cellular proteolysis (21).
Therefore, understanding the binding behavior of the UBL
domains of UBL-UBA proteins with their various interacting
proteins will shed light on the regulatory role of these proteins.
Despite the identification of a large number of UBDs, structural
details of integral UBL-binding domains are limited. In some
cases, the intra- and intermolecular interactions between these
UBLs with known UBDs such as UBA or the ubiquitin-interact-
ing motif (UIM) have been demonstrated by solution NMR
(35–38).
Here, we are reporting crystal structures of budding yeast
Ufd2 in complex with the UBL domains of Rad23 and Dsk2 and
the molecular details of their interaction interfaces. We identify
a novel sequence pattern in the N-terminal UBL-binding region
of budding yeast Ufd2, which is conserved in lower eukaryotes
and is distinct from any known UBD identified so far. More-
over, despite engaging the same binding region, residue-spe-
* This work was supported by Deutsche Forschungsgemeinschaft Grant
RA1643/2-1 (to S. R.) and Rudolf Virchow Center for Experimental Biomed-
icine Grant FZ 82 (to H. S.).
The atomic coordinates and structure factors (codes 3M62 and 3M63) have been
deposited in the Protein Data Bank, Research Collaboratory for Structural
Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).
□
S The on-line version of this article (available at http://www.jbc.org) contains
supplemental Experimental Procedures, Tables S1 and S2, and Figs. S1–S5.
1 Both authors contributed equally to this work.
2 To whom correspondence may be addressed. E-mail: hermann.schindelin@
virchow.uni-wuerzburg.de.
3 To whom correspondence may be addressed. E-mail: shahri.raasi@
uni-konstanz.de.
4 The abbreviations used are: UBL, ubiquitin-like; UBA, ubiquitin-associated;
UIM, ubiquitin-interacting motif; UBD, ubiquitin-binding or ubiquitin-like
binding domain; UFD, ubiquitin fusion degradation; Ub, ubiquitin; GST,
glutathione S-transferase; ITC, isothermal titration calorimetry; SPR, sur-
face plasmon resonance; r.m.s., root mean square; WT, wild type; PDB, Pro-
tein Data Bank; h, human; Sc, S. cerevisiae; Sp, S. pombe.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 26, pp. 20390–20398, June 25, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
20390
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 26•JUNE 25, 2010
cific differences exist in the interactions of the UBL domains of
Rad23 and Dsk2 with Ufd2, which are coupled to subtle differ-
ences in their overall binding affinities. Mutational analyses of
the binding surface of the UBL domains and a closer inspection
of the thermodynamic contributions of those residues point to
adaptive evolution as a factor shaping these interfaces.
EXPERIMENTAL PROCEDURES
Cloning, site-directed mutagenesis, protein expression, and
purification
are
described
in
the
supplemental
Experi-
mental Procedures.
Crystallization of Ufd2Rad23-UBL and Ufd2Dsk2-UBL—
For crystallization of the Ufd2Rad23-UBL and Ufd2Dsk2-UBL
complexes, Ufd2 was incubated with Rad23-UBL or Dsk2-UBL
at a molar ratio of 1:1.5 (77 M Ufd2 and 115.5 M UBL) for 1 h
at 4 °C in the presence of 2 mM dithiothreitol. Crystals were
grown by vapor diffusion in hanging drops containing equal
volumes of protein in 50 mM HEPES, pH 7.4, 150 mM NaCl, and
2 mM dithiothreitol and a reservoir solution consisting of
16–18% (w/v) polyethylene glycol 3500 and 200 mM K3-citrate,
pH 8.3, equilibrated against the reservoir solution. Crystals
were cryo-protected by soaking in mother liquor containing
15–20% (v/v) glycerol. They belong to space group P212121 with
approximate cell dimensions of a 65 Å, b 126 Å, and c
181 Å with one complex per asymmetric unit.
Data Collection and Structure Determination—Crystals were
flash-cooled in liquid nitrogen, and data collection was per-
formed at 100 K. Data were collected at beamlines ID14–4
(European Synchrotron Radiation Facility (ESRF), Grenoble,
France) and BL 14.1 (Berliner Elektronenspeicherring-Gesell-
schaft fu¨r Synchrotronstrahlung (BESSY), Berlin, Germany)
and processed using Mosflm and Scala (39, 40). Data collection
statistics are summarized in supplemental Table S1. For subse-
quent calculations, the CCP4 suite was utilized (41) with excep-
tions as indicated. The Ufd2 structure was solved by molecular
replacement using Phaser (42) with Protein Data Bank (PDB)
entry 2QIZ as search model. Because Phaser could not find a
solution for the UBL domain with different search models, this
domain was fitted manually into the electron density using
human ubiquilin 3 (PDB entry 1YQB) for the Ufd2Rad23-UBL
complex and the Dsk2-UBL domain (PDB entry 2BWF) for the
Ufd2Dsk2-UBL complex as a model. The structures were
refined with Phenix (43) and REFMAC5 incorporating transla-
tion, libration, screw-rotation (TLS) refinement in all cycles
(44, 45). Solvent molecules were automatically added with Coot
(46). The figures were produced with PyMOL (65).
In Vitro Binding Assays—For pulldown assays, GST-tagged
Ufd2 and variants were immobilized on glutathione (GSH)
beads. In all experiments, 20 l of GSH beads were incubated
with 0.95 M purified Ufd2 in 400 l of phosphate-buffered
saline buffer with 1 mM dithiothreitol and 0.1% (v/v) Triton
X-100 at 4 °C for 1 h. WT-Ufd2 and GST alone were included as
controls. After centrifugation (1250 g, 30 s), beads were
washed five times with 400 l of binding buffer. Purified UBL
proteins (0.95 M) in a total volume of 400 l of binding buffer
were added to immobilized Ufd2 and treated in the same way as
in the first step. Immobilized proteins were analyzed by 17%
(v/v) SDS-PAGE or by immunoblotting with an anti-His
antibody.
Isothermal Titration Calorimetry (ITC)—Proteins were
extensively dialyzed against phosphate-buffered saline buffer
(pH 7.4, 1 mM -mercaptoethanol) followed by degassing. In all
experiments, 75–150 M Rad23- and Dsk2-UBL proteins were
titrated as the ligand into the sample cell containing 5–10 M
Ufd2. A volume of 10 l of ligand was added at a time with a
total number of 30 injections, resulting in a final molar ratio of
ligand-to-protein varying between 3:1 and 4:1. All experiments
were performed using a VP-ITC instrument (MicroCal, GE
Healthcare) at 25 °C. Buffer-to-buffer, buffer-to-Ufd2, as well
as Rad23-UBL/Dsk2-UBL-to-buffer titrations were performed
as described above. Corrected data were analyzed with a single-
site binding model using software supplied by the ITC manu-
facturer and non-linear least squares fitting to calculate the
dissociation constant (Kd).
Surface Plasmon Resonance (SPR) Measurements—SPR
binding assays were performed alternatively on BIAcore X or
BIAcore T100 instruments (GE Healthcare) at 25 °C in 10 mM
HEPES, pH 7.4, 150 mM NaCl, 50 M EDTA, 1 mM -mercap-
toethanol, and 0.005% (v/v) Surfactant P20. 100 response units
of His-tagged Rad23- or Dsk2-UBL were captured on a nickel-
nitrilotriacetic acid (Ni-NTA) sensor chip. GST-tagged Ufd2
for comparative binding assays and untagged Ufd2 for affin-
ity analysis were applied to the UBL surfaces in random
duplicates at a flow rate of 50 l/min. After each cycle, the
surface was regenerated using 350 mM EDTA in running
buffer to remove bound Ni2 and captured proteins. The
BIAcore T100 evaluation software was used to calculate the
steady state affinity constants. Data were plotted using
GraphPad Prism. For comparative assays, the relative bind-
ing responses of the mutants to WT proteins were deter-
mined by obtaining the maximum response for each interac-
tion at the end of each injection.
RESULTS
Ufd2 Binds the UBL Domains of Rad23 and Dsk2 with High
Affinity—Although Rad23 and Dsk2 interact with Ufd2 via
their UBL domains (22, 26), yeast two hybrid assays could only
identify the isolated N-terminal fragment (residues 1–380) of
Ufd2 as its UBL-interacting region (26). Additional details
regarding the Ufd2-UBL interactions have not been unraveled
so far. To further characterize the interactions of Ufd2 with the
UBLs of Rad23 and Dsk2, we performed GST pulldown assays
with GST-tagged full-length Ufd2 and C-terminally His-tagged
UBLs (Fig. 1A). Both UBLs were readily captured using immo-
bilized GST-Ufd2. In contrast, the UBL domain of Ddi1, a third
UBL-UBA protein, does not interact with Ufd2 (Fig. 1A) (22).
The differential binding of the Rad23- and Dsk2-UBLs to the
proteasomal subunits Rpn1 and Rpn10 has been described (25,
47, 48). Hence, we used SPR interaction analysis to search for
quantitative differences in their interactions. Steady state affin-
ity analysis of Ufd2 on both Rad23-UBL (Fig. 1B, left panel, and
1C) and Dsk2-UBL surfaces (Fig. 1B, right panel, and 1C) pro-
vided a Kd of 55 3 nM for the interaction of Rad23-UBL and a
lower affinity for Dsk2-UBL with a Kd of 418 56 nM.
UBL-binding Domain of Ufd2
JUNE 25, 2010•VOLUME 285•NUMBER 26
JOURNAL OF BIOLOGICAL CHEMISTRY 20391
The binding of the UBLs of Rad23 and Dsk2 to Ufd2 was also
analyzed by ITC to allow for a complete thermodynamic char-
acterization (Fig. 1D). These studies resulted in a Kd of 70 6
nM for the interaction of Rad23-UBL with Ufd2 and for the
binding of Dsk2-UBL to Ufd2 in a 2–3-fold higher Kd of 175
19 nM. Although there is an excellent agreement between SPR
and ITC for the Rad23-Ufd2 interaction, the two methods show
an 2-fold difference for the Dsk2-Ufd2 interaction. More
importantly, the enthalpic and entropic components to the free
energy are highly different between the two UBLs. The interac-
tion of Rad23-UBL and Ufd2 is more exothermic (H 17.3
kcal/mol) when compared with Dsk2-UBL (H 10.1 kcal/
mol). However, this is offset by a substantial decrease in entropy
for Rad23-UBL (TS 7.4 kcal/mol), whereas the entropic
contribution
is
minimal
for
the
Dsk2-UBL
interaction
(TS 0.8 kcal/mol).
Crystal Structures of Ufd2 in Complex with Rad23- and
Dsk2-UBL—We solved the structures of Ufd2 in complex with
Rad23-UBL carrying either an N-terminal or a C-terminal His
tag, which showed no significant structural differences. Due to
better data quality, the structure of Ufd2 with a C-terminal
His-tagged UBL is presented here. The Ufd2Rad23-UBL com-
plex was refined at 2.4 Å resolution to a crystallographic R-fac-
tor of 20.3% and a free R-factor of 25.7% (Table 1). As described
previously (49), Ufd2 is composed of an N-terminal variable
domain, a core domain, and a C-terminal U-box domain with a
fold similar to that of RING (really interesting new gene)
domains, which are present in certain Ub ligases (Fig. 2A).
Despite some conformational variability of the U-box domain,
our Ufd2 structure in the complex is quite similar (1.5 Å root
mean square (r.m.s.) deviation for 954 C atoms) to the pub-
lished Ufd2 structure (49).
The N-terminal variable region of Ufd2 that binds to the UBL
domain consists of eight -helices. Helices 1 to 4 are
arranged in a four-helix bundle, whereas helices 5 and 6
interact with 3 and 4 through hydrophobic contacts that are
partly mediated by their connecting loops (Fig. 2B). The struc-
ture of Rad23-UBL is comprised of a five-stranded -sheet, one
-helix, and one 310-helix (Fig. 2B). It displays a high degree of
similarity with Ub (PDB entry 1UBQ, 1.1 Å r.m.s. deviation for
72 C atoms, z-score 14, 25% sequence identity) and the UBL
domain of hHR23A (PDB entry 1P98, 1.6 Å r.m.s. deviation for
FIGURE 1. Interactions of Ufd2 with the UBL domains of Rad23 and Dsk2. A, GST-Ufd2 immobilized on GSH-beads was tested for binding to C-terminally
His-tagged UBLs of Rad23, Dsk2, and Ddi1. Captured UBLs were visualized by immunoblotting (WB) with an anti-His antibody. 2% of the input and GST beads
incubatedwithUBLswereloadedascontrols.B,aseriesof2-foldUfd2dilutions(233–3.6nM)wasappliedonaRad23-orDsk2-UBLsurfacefor120s(leftandright
panel,respectively).RU,responseunits.C,SPRbindingisothermsofWT-Rad23-andWT-Dsk2-UBLandthequintupleandseptupleDsk2-UBLvariantswithUfd2.
conc., concentration. D, ITC analysis of Ufd2Rad23-UBL (closed circles) and Ufd2Dsk2-UBL (open circles) complexes.
TABLE 1
Refinement statistics
Ufd2Rad23-UBL
Ufd2Dsk2-UBL
Resolution limit (Å)
45.2-2.4
73.5-2.4
No. of reflections
56,268
55,087
No. of protein/ligand/solvent atoms
8303/17/298
8288/17/182
Rcryst (Rfree)a,b
0.203 (0.257)
0.210 (0.270)
r.m.s. deviations in:
Bond lengths (Å)
0.016
0.015
Bond angles (°)
1.711
1.610
Estimated coordinate error (Å)
0.25
0.26
Overall average B-factor (Å2)
25.7
42.9
Ramachandran statistics (%)c
93.1/97.9/2.1
93.8/98.4/1.6
aRcryst hklFo Fc/hklFo where Fo and Fc are the observed and calculated
structure factor amplitudes.
bRfree, same as Rcryst for 5% of the data randomly omitted from the refinement. The
estimated coordinate error is based on Rfree.
c Ramachandran statistics indicate the fraction of residues in the favored (98%),
allowed ( 99.8%), and disallowed regions of the Ramachandran diagram, as
defined by MolProbity (64).
UBL-binding Domain of Ufd2
20392
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VOLUME 285•NUMBER 26•JUNE 25, 2010
72 C atoms, z-score 11.3, 26% sequence identity), one of the
two human homologs of budding yeast Rad23.
Subsequently, we solved the structure of Ufd2 with Dsk2-
UBL by molecular replacement. The UBL domain in the
Ufd2Dsk2-UBL complex exhibits increased flexibility; in par-
ticular, with a C-terminally tagged UBL domain, the first 30
amino acids of this domain were largely disordered (data not
shown). With an N-terminally tagged protein, the Ufd2Dsk2-
UBL structure was refined at 2.4 Å resolution to a crystallo-
graphic R-factor of 21.0% and a free R-factor of 27.0% (Table 1).
Both Rad23-UBL and Dsk2-UBL structures can be superim-
posed with an r.m.s. deviation of 1.1 Å for 71 aligned residues
(z-score 13.6, 30% sequence identity).
Analysis of the Ufd2Rad23-UBL Interface—The Ufd2UBL
interface in the structure of the complex buries a total molecu-
lar surface of about 1260 Å2, which is comprised to 590 Å2 of
the molecular surface of Ufd2 (1.3% of the total surface area)
and 670 Å2 from UBL (14.6% of the total surface area). This
interface is composed of almost equal parts of non-polar resi-
dues (38%), polar residues (33%), and charged residues (29%);
however, there are only one salt bridge (UBL-Lys-10 N–Ufd2-
Glu-49 O1 with a distance of 2.6 Å) and two direct hydrogen
bonds (UBL-Ser-47 O–Ufd2-Arg-92 N2, UBL-Gln-52 N2–
Ufd2-Glu-141 O at distances of 2.3 and 3.2 Å, respectively)
present (Fig. 3A).
Three UBL segments are contacting Ufd2 (Fig. 3A). Segment
I is located in the loop connecting -strands one and two, seg-
ment II involves -strands three and four, and segment III is
located in -strand five. Ufd2 residues from helix 2 and 4 as
well as the loop connecting 4 with 5 contribute to the
Ufd2UBL interface. These residues contact the hydrophobic
surface of the UBL -sheet in the region of -strands 3, 4, and 5.
Participating residues from Ufd2 include Leu-44, Tyr-97, Val-
100, and Trp-107, which are located in the hydrophobic UBL
pocket formed by residues Phe-9, Ile-45, Val-50, Val-69, and
Met-71 of Rad23 (Fig. 3, A and B).
For comparison, the principal recognition determinants in
Ub are: 1) the hydrophobic pocket formed by the side chains of
Leu-8 (Phe-9 in Rad23), Ile-44 (Ile-45 in Rad23), His-68 (Val-69
in Rad23), and Val-70 (Met-71 in Rad23) and 2) the main chain
amide group of Gly-47 (Gly-48 in Rad23), which is involved in
hydrogen bonding (50). Although the hydrophobic patch of
Rad23-UBL is also crucial for its interaction with Ufd2, the
main chain of Gly-48 does not form a hydrogen bond. Instead,
the -turn (Ser47–Gly48) connecting -strands 3 and 4 is stabi-
lized by the aforementioned strong hydrogen bond between
Ufd2-Arg-92 and UBL-Ser-47, whereas Ufd2-Gly-96 and Ufd2-
Tyr-97 contact UBL-Gly-48 (Fig. 3A). The aromatic ring of
Ufd2-Tyr-97 is involved in a stacking interaction with the pep-
tide bond between UBL residues 47 and 48 in this -turn.
Probing the Ufd2Rad23-UBL Interface—The importance of
interface residues was analyzed by mutagenesis experiments.
Eleven residues from Ufd2 and nine from Rad23-UBL were
each replaced with Ala. With the exception of the Rad23-UBL-
G48A variant that showed a reduced expression, all Ufd2 and
Rad23-UBL variants behaved like the WT protein during and
after purification, indicating that they were correctly folded
(data not shown). Initially, the contribution of these residues
was studied by GST pulldown and comparative SPR binding
assays (Table 2, supplemental Figs. S1 and S2A). In SPR studies,
the relative binding responses of mutants to WT proteins were
determined and compared. The majority of Rad23-UBL single
mutants revealed reduced binding to Ufd2 with Rad23-UBL-
I45A displaying the most prominent binding defect. The con-
tribution of the remaining residues to the interaction is aug-
mented in double mutants (supplemental Fig. S1C). Analysis of
the Ufd2 variants by SPR showed a largely reduced binding of
the residues located in the hydrophobic region of the UBL-
binding pocket (Leu-44, Tyr-97, Val-100, and Phe-107) and
Asp-40 (Table 2 and supplemental Fig. S2A).
ITC studies confirmed these results and allowed for a
quantitative analysis (Table 2, supplemental Fig. S3 and
supplemental Table S2). The most significant effect for Ufd2
was observed for all residues located in the hydrophobic UBL
pocket. Mutation of Val-100 and Phe-107 to Ala completely
abolished binding, the Y97A variant strongly reduced binding
(1900-fold), and the I104A and L44A variants showed signifi-
cantly decreased affinities (20- and 120-fold, respectively).
Although not directly involved in complex formation (Fig. 3A),
the Ufd2-D40A variant showed a 110-fold reduced affinity
(Table 2), which probably is the result of the missing intramo-
lecular hydrogen bond between Ufd2-Asp-40 and Ufd2-Tyr-97
(O2–OH 2.5 Å). This hydrogen bond seems to be crucial for
proper positioning of the aromatic side chain of Tyr-97 in the
interface region and might be important to align helices 2 and
4 for interaction with the Rad23-UBL.
FIGURE 2. Structure of Ufd2 in complex with the UBL domain of Rad23.
A, ribbon representation of the overall structure of the Ufd2Rad23-UBL com-
plex. The Rad23-UBL domain is shown in green, the N-terminal Ufd2 region is
in orange, the Ufd2 core domain is in gray, and the Ufd2 U-box domain is in
red. B, close-up view of the N-terminal Ufd2 domain in complex with Rad23-
UBL with secondary structural elements labeled and color-coded as in A.
UBL-binding Domain of Ufd2
JUNE 25, 2010•VOLUME 285•NUMBER 26
JOURNAL OF BIOLOGICAL CHEMISTRY 20393
In Rad23-UBL, Ile-45 was shown to be integral for binding to
Ufd2 by the detrimental effect (130-fold decrease) after
exchange to Ala (Table 2). Mutation of Phe-9, Val-50, and Val-
69, residues adjacent to Ile-45 in the hydrophobic patch, also
decreased the affinity of Rad23-UBL 5–7-fold. Ser-47, which is
in hydrogen-bonding distance to Ufd2-Arg-92 and next to
UBL-Gly-48, showed a 9-fold reduced affinity. In Ub and in the
human Rad23 homolog hHR23A, Ser-47 is replaced by Ala.
Charged residues found in the interface (Ufd2, Glu-26, Glu-49,
and Arg-92; UBL, Lys-10) do not contribute significantly to the
interaction. In summary, our data indicate that the most prom-
inent contact between Ufd2 and Rad23-UBL is the strong
hydrophobic interaction between UBL-Ile-45 and Ufd2-Val-
100 as well as Ufd2-Phe-107, which defines the core of the UBL-
interacting region of Ufd2.
Molecular Discrimination between Rad23 and Dsk2—De-
spite a similar fold, the UBL domains of Rad23 and Dsk2 display
only 30% sequence identity, which could give rise to differences
in their interactions. A superposition of the bound Rad23-UBL
and Dsk2-UBL in the two complex structures showed signifi-
cant changes (Fig. 3C). Of the three UBL segments involved in
the Ufd2 interaction (Fig. 3A), segment II including Ile-45
(Ile-44 in Ub) is highly conserved, and there are no conforma-
tional changes in both UBL structures, whereas segments I and
III are not conserved and display structural changes (Fig. 3C).
The loop, connecting -strands one and two, adopts different
conformations, and -strand five shows a displacement that
might affect binding (Fig. 3C).
Segment I includes Phe-9 in Rad23-UBL, corresponding to
Leu-8 in Ub, where this residue is also involved in Ub recogni-
tion by UBDs (50, 51). Phe-9 is replaced by Gly-10 in Dsk2-UBL,
and there is no corresponding hydrophobic interacting residue
(supplemental Fig. S4A). Dsk2 residues Gly-10 and Gln-11
adopt different conformations when compared with Leu-8/
Thr-9 of Ub and Phe-9/Lys-10 of Rad23-UBL. In the
Ufd2Dsk2-UBL structure, the Ufd2Rad23-UBL salt bridge
(Lys-10/Glu-49) is missing due to the Lys-10 to Gln-11
exchange, with the latter side chain no longer being located in
the protein interface (supplemental Fig. S4A). The missing
interaction from segment I in Dsk2 might be compensated by
the displacement of -strand five toward Ufd2 and a replace-
ment of Val-69 to His-69 found in segment III resulting in a
more pronounced interaction in this region when compared
with Rad23-UBL (supplemental Fig. S4A). The presence of the
salt bridge seems to be the reason for the more exothermic
character of the Ufd2Rad23-UBL interaction, a view that is also
supported by the corresponding Ufd2-E49A and Rad23-K10A
variants, which both display binding enthalpies similar to the
FIGURE 3. The Ufd2Rad23-UBL interface. A, residues involved in binding are shown in stick representation. Carbon atoms of Ufd2 residues are colored in
orange and in green for Rad23-UBL. Dashed lines indicate H-bonds. B, structure-based sequence alignment of Rad23-UBL, Dsk2-UBL, hHR23A-UBL, and Ub.
Secondary structure elements of Rad23-UBL were assigned using DSSP (61) and are labeled above the sequences. The alignment was performed using DaliLite
(62), and the figure was prepared with ESPript (63). Strictly conserved amino acids are highlighted with a red background, and similar amino acids are shown as
redletters.ThethreeUfd2-bindingsegmentsareindicated.ResiduesinvolvedinUfd2Rad23-UBLinteractionarelabeledwithgreenstars.C,superpositionofthe
Ufd2Rad23-UBL/Dsk2-UBL complex structures with the N-terminal binding domain of Ufd2 in orange (Rad23 complex) and gray (Dsk2 complex), with Rad23-
UBL in green and Dsk2-UbL in yellow.
UBL-binding Domain of Ufd2
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Ufd2Dsk2-UBL complex (supplemental Table S2). At the same
time, the absence of the salt bridge in both mutants is accom-
panied by a more favorable entropic contribution, which is on a
level similar to the Ufd2Dsk2-UBL complex.
To identify residues important for the subtle molecular dis-
crimination between the UBL domains of Rad23 and Dsk2, the
interaction of Dsk2-UBL with Ufd2 mutants was analyzed by
GST pulldown assays (data not shown), SPR, and ITC (Table 2,
supplemental Figs. S2B and S3C). Quantitative ITC analysis
showed reduced binding of Dsk2 to Ufd2 mutants Y97A (470-
fold), V100A (22-fold), I104A (6-fold), and F107A (20-fold)
(Table 2). However, binding of the V100A and F107A variants is
not completely abolished, and when compared with Rad23-
UBL, the binding affinities are less affected by a factor of about
3–7 in most of the mutants analyzed. In addition, the L44A
mutant, which has a 120-fold reduced affinity with Rad23-UBL,
is only three times reduced in the case of Dsk2-UBL.
In general agreement with the ITC affinity data, the compar-
ative SPR binding assay revealed significant differences in the
association of Ufd2 variants Y97A, V100A, I104A, and F107A
with Rad23- and Dsk2-UBL surfaces (supplemental Fig. S4B).
The observed SPR decrease for the binding of the T48A and
E49A variants of Ufd2 to Dsk2-UBL seems to be compensated
by slower dissociations, thus explaining why these mutants
show no significant defect in the ITC analysis.
To further analyze the contribution of segments I and III to
complex formation, a G10F/Q11K/S67Q/H69V/V71M quintu-
ple Dsk2-UBL mutant was generated, where key residues in
binding segments I and III were replaced with the correspond-
ing residues from Rad23-UBL. Comparative binding as well as
steady state affinity analysis by SPR revealed only a small
increase (Kd 348 nM) in binding affinity for Ufd2 when com-
pared with WT-Dsk2-UBL (Kd 418 nM) (data not shown and
Fig. 1C). In addition, neither a crystal structure of the quintuple
Ufd2Dsk2-UBL complex (data not shown) nor the KD of 240
nM deduced by ITC revealed significant differences from
WT-Dsk2-UBL (Kd 175 nM). The ITC analysis did, however,
reveal that the binding is now driven by an increase in entropy
(TS 6.5 kcal/mol versus 0.8 and 7.4 kcal/mol for
WT-Dsk2-UBL and -Rad23-UBL, respectively), whereas the
binding enthalpy is reduced to only 2.5 kcal/mol when com-
pared with 10.1 and 17.3 kcal/mol (supplemental Table S2).
Interestingly, SPR and ITC analysis of a G10F/Q11K/I50V/
K52Q/S67Q/H69V/V71M septuple Dsk2-UBL mutant, which
has the additional I50V and K52Q substitutions in segment II,
showed an even lower affinity (SPR, Kd 648 nM; ITC, Kd 875
nM) to Ufd2 when compared with WT-Dsk2-UBL (Fig. 1C).
The N Terminus of Ufd2 Represents a Unique and Conserved
UBL-binding Domain—A multiple sequence alignment of Ufd2
from different yeast species displays a distinct pattern of con-
served residues involved in UBL binding (Fig. 4A). Among the
available yeast genomes, the Schizosaccharomyces pombe
sequence is most similar to those from higher eukaryotes; thus
we isolated cDNA fragments for the coding region of the UBL
domains of Rad23 and Dsk2 and full-length Ufd2 from this
organism and examined their interactions by GST pulldown
assays (Fig. 4B) as well as SPR (data not shown). We could show
that SpUfd2 interacts strongly with the UBL domains of
SpRad23 and SpDsk2 as well as with the UBL domains of
ScRad23 and ScDsk2 and vice versa. This cross species interac-
tion, despite the diversified UBL and Ufd2 amino acid
sequences, indicates that the identified sequence pattern repre-
sents a real UBL-interacting domain. A surface representation
of this motif is shown in Fig. 4C.
The N terminus of budding yeast Ufd2 displays only limited
sequence homology with the human Ufd2s, E4A and E4B
(supplemental Fig. S5) and other Ufd2s from higher eukaryotes.
In agreement with this finding, there are no reports that
hHR23A/B interacts with either of the human homologs of
Ufd2. Interestingly, our SPR studies showed that the UBL
domain of hHR23A interacts with ScUfd2, albeit with lower
affinity (data not shown). Apparently, the high affinity interac-
tion of the UBL domains of Rad23 and Dsk2 has been lost dur-
ing the evolution of this domain. The absence of conservation
of the Ufd2-UBL interface could potentially be used for thera-
peutic interventions against pathogenic yeasts such as Candida
albicans by designing low molecular weight compounds that
disrupt this interface. However, further functional studies in
pathogenic yeasts are required to examine the suitability of this
surface as a drug target.
TABLE 2
ITC and SPR parameters of Ufd2, Rad23-UBL, Dsk2-UBL, and variants
indicates no change; ND indicates not detected (corresponding to at least a
104-fold decrease in binding affinity).
Ufd2
WT-UBL
ITC
SPRa (% of
relative
response)
Kd
Fold decrease
nM
WT
Rad23
70
100
Dsk2
175
100
E26A
Rad23
284
4
91
Dsk2
521
3
83
D40A
Rad23
7900
110
20
Dsk2
7600
40
0
L44A
Rad23
8300
120
31
Dsk2
463
3
52
T48A
Rad23
72
70
Dsk2
296
2
29
E49A
Rad23
413
6
69
Dsk2
314
2
44
R92A
Rad23
265
4
76
Dsk2
128
59
G96A
Rad23
592
8
51
Dsk2
216
60
Y97A
Rad23
134,000
1900
3
Dsk2
83,000
470
0
V100A
Rad23
ND
10,000
9
Dsk2
3900
22
1
I104A
Rad23
1600
20
43
Dsk2
1100
6
12
F107A
Rad23
ND
10,000
11
Dsk2
3600
20
0
Ufd2
Rad23-UBL
ITC
SPRa (% of
relative
response)
Kd
Fold decrease
nM
WT
F9A
376
5
80
WT
K10A
162
2
96
WT
I45A
9100
130
17
WT
S47A
606
9
62
WT
V50A
441
6
88
WT
Q52A
415
6
79
WT
Q67A
113
2
92
WT
V69A
478
7
70
WT
M71A
221
3
88
a For comparative SPR assays, the relative binding responses of the mutants to wt
proteins were determined by obtaining the maximum response for each interac-
tion at the end of injection.
UBL-binding Domain of Ufd2
JUNE 25, 2010•VOLUME 285•NUMBER 26
JOURNAL OF BIOLOGICAL CHEMISTRY 20395
DISCUSSION
Families and superfamilies of protein domains or folds have
evolved through a process of homologous recombination and
gene duplication (52) followed by sequence divergence. Mem-
bers of different classes of UBDs such as UBA or UBL domains
result from these processes. For instance, the UBL domains of
Rad23 and Dsk2 display only 30% sequence identity but adopt
the same fold and utilize the same binding surface to recognize
a common UBL-binding domain of Ufd2 to form complexes
that display similarly high affinity. Nevertheless, not all inter-
acting residues are conserved; in particular, there is sequence
diversity in binding segments I and III of UBLs. Our attempts to
interconvert the UBL domains by altering non-conserved inter-
facial residues were not successful, thus suggesting that addi-
tional elements exist and play a role in the respective Ufd2-UBL
interaction. Interestingly, these results resemble earlier studies
on WW domains (53, 54), where a statistical analysis of multiple
sequence alignments was utilized to identify co-evolving resi-
dues. The authors demonstrated that not only interfacial resi-
dues but also buried residues distal to the interface co-evolved
with interfacial residues and contribute significantly to the
interactions. They concluded that certain sequence patterns in
interacting domains are due to adaptive evolution. In agree-
ment with these findings, our data prove that substitution of
key interfacial residues of Dsk2-UBL has no significant effect on
its overall binding affinity to Ufd2. In case of the septuple
mutant, we even observed a decrease in binding affinity, which
could be due to the imposed disorder into the evolutionary
inter-residue relations within the UBL fold. This is supported
by the fact that when compared with Dsk2-UBL and in partic-
ular Rad23-UBL, the binding of the quintuple Dsk2-UBL
mutant is driven strongly by entropy. These findings indicate
that binding interfaces can be modulated by changes in residues
that affect either the binding enthalpy or the entropy, thus pro-
viding additional freedom to maintain an interaction during the
course of evolution, an effect that has been described previously
as entropy/enthalpy compensation (55, 56).
Our studies suggest that UBL domains have co-evolved with
Ufd2 to reach optimal binding affinities by altering specific res-
idue-to-residue interactions (co-evolution at the residue level)
(57), while at the same time, all functional aspects of Rad23 or
Dsk2 are preserved. Therefore, the primary sequence degener-
FIGURE 4. The N terminus of Ufd2 represents a conserved UBL-interacting domain in lower eukaryotes. A, alignment of the N-terminal sequences of
fungal Ufd2s. Invariant or conserved residues with surface access are colored in dark blue, buried ones are in light blue. Residues labeled with red stars represent
the core region of the binding domain, which is essential for UBL interaction, whereas residues labeled with yellow stars contribute moderately to the
interaction. K. lactis, Kluyveromyces lactis; C. glabrata, Candida glabrata; Z. rouxii, Zygosaccharomyces rouxii; L. thermotolerans, Lachancea thermotolerans; C.
tropicalis, Candida tropicalis; C. dubliniensis, Candida dubliniensis; P. guilliermondii, Pichia guilliermondii; D. hansenii, Debaryomyces hansenii. B, GST pulldown
assay demonstrates the cross interactions of S. pombe and S. cerevisiae proteins. 5% of inputs and GST beads incubated with UBLs were loaded as controls. WB,
Western blot. C, surface representation of the N-terminal UBL-binding domain of Ufd2, color-coded as in A.
UBL-binding Domain of Ufd2
20396
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 26•JUNE 25, 2010
acy of protein domains such as UBAs or UBLs has been toler-
ated and established in favor of the cooperative nature of the
interactions and their functionality within their respective pro-
tein complexes. This further suggests that differential binding
properties observed for the interactions of Ufd2 with UBLs (this
study) or for the interactions of UBAs with Ub and Ub chains
(14) can arise not necessarily due to their interaction with dif-
ferent ligands but can also result from the adaptive co-evolution
of these domains with the same interacting partners. Seem-
ingly, these interfacial domains have evolved to hold protein-
protein interactions in a suitable form within multicomponent
complexes until they are challenged by downstream events.
Numerous structures of Ub receptors in complex with their
respective Ub/UBL-binding domains have been reported. The
so far characterized Ub receptors of the 26 S proteasome in
budding yeast encompass the two proteasomal subunits Rpn10
(S5a in humans) and Rpn13 and the three UBL-UBA proteins
Rad23, Dsk2, and Ddi1, which associate with the proteasome
and function as shuttle factors (21). Experimental evidence for
the existence of additional candidates exist (21, 58). Rad23 and
Dsk2 interact with the proteasomal subunit Rpn1 via their UBL
domains (21, 47). Aside from their known interactions with Ub,
Rpn13 and Rpn10/S5a alternatively interact with UBL-UBA
proteins (21, 35, 37, 38, 48, 51, 59). For instance, the preferential
association of Rpn1 with Rad23 and Rpn10 with Dsk2 has been
reported (25, 38, 47, 48). Based on the binding of hpLIC2 (Dsk2
homolog) with Rpn13, an interac-
tion of Dsk2 with Rpn13 has been
proposed (51, 59).
Although
the
aforementioned
examples engage essentially the
same
surface
of
Ub/UBL,
they
diverge in both structure and pat-
terns of Ub/UBL recognition (Fig.
5). For instance, hRpn10/S5a recog-
nizes the UBL domain of hHR23A,
one of the two human homologs of
Rad23, via a Ub-interacting motif,
which consists of a single -helix
(35, 37). Rpn13 binds Ub via a pleck-
strin homology domain, which is a
seven-stranded -sandwich capped
by an -helix (51). The Ub-binding
surface of Rpn13 is formed by
three loops that bridge -strands.
Another Ub-binding element is the
UBA domain found for example in
Dsk2 (60). The UBA domain is com-
posed of a three-helix bundle. With
the exception of Rpn13, which
exclusively binds via loops, it seems
that the majority of Ub/UBL-bind-
ing domains fold into -helical
structures
including
the
known
UBDs, UIM, and UBA, and the
UBL-binding domain of Ufd2 iden-
tified in this study. Despite the pre-
dominant
interaction
involving
-helices as Ub/UBL-binding elements, the three-dimensional
structure of the UBL-binding domain of Ufd2 differs from other
known examples, hence providing the first structural descrip-
tion for how Ufd2 acts as a UBL receptor while at the same time
further enhancing the diversity of UBDs in general.
Acknowledgments—We thank Martin Scheffner and Keith Wilkinson
for critical reading of the manuscript. We thank Stefan Jentsch for
providing the original plasmids for the expression of Rad23, Dsk2, and
Ufd2 and for Ufd2-specific antibodies used in the initial phase of this
study. We also thank David Fischer and Rodrigo Villasen˜or for the
contribution to this study and Sven Eiselein for providing us with
C-terminal GST-tagging plasmid.
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UBL-binding Domain of Ufd2
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|
3M6B
|
Crystal Structure of the Ertapenem Pre-isomerized Covalent Adduct with TB B-lactamase
|
Biochemical and Structural Characterization of Mycobacterium
tuberculosis β-Lactamase (BlaC) with the Carbapenems
Ertapenem and Doripenem
Lee W. Tremblay#, Fan Fan#, and John S Blanchard‡,*
Department of Biochemistry, Albert Einstein College of Medicine, 1300 Morris Park Avenue,
Bronx, New York 10461
Abstract
Despite the enormous success of β-lactams as broad-spectrum antibacterials, they have never been
widely used for the treatment of TB due to intrinsic resistance that is caused by the presence of a
chromosomally-encoded gene (blaC) in Mycobacterium tuberculosis. Our previous studies of TB
BlaC revealed that this enzyme is an extremely broad-spectrum β-lactamase hydrolyzing all β-
lactam classes. Carbapenems are slow substrates that acylate the enzyme but are only slowly
deacylated and can therefore act also as potent inhibitors of BlaC. We carried out the in vitro
characterization of doripenem and ertapenem with BlaC. A steady-state kinetic burst was observed
with both compounds with magnitudes proportional to the concentration of BlaC used. The results
show apparent Km and kcat values of 0.18 µM and 0.016 min−1 for doripenem and 0.18 µM and
0.017 min−1 for ertapenem. FTICR mass spectrometry demonstrated that the doripenem and
ertapenem acyl-enzyme complexes remain stable over a time period of 90 min. The BlaC-
doripenem covalent complex obtained after 90 minutes of soaking was solved to 2.2 Å, while the
BlaC-ertapenem complex obtained after a 90 minute soak was solved to 2.0 Å. The 1.3 Å
diffraction data from a 10 minute ertapenem-soaked crystal revealed an isomerization occurring in
the BlaC-ertapenem adduct in which the original Δ2 pyrroline ring was tautomerized to generate
the Δ1 pyrroline ring. The isomerization leads to the flipping of the carbapenem-hydroxyethyl
group to hydrogen bond to the carboxyl O2 of Glu166. The hydroxyethyl flip results in both
decreased basicity of Glu166 and in a significant increase in the distance between the carboxyl O2
of Glu166 and the catalytic water molecule, slowing hydrolysis.
Tuberculosis (TB), caused by Mycobacterium tuberculosis, continues to be a worldwide
health concern (1). There were an estimated 9.3 million new cases of TB in 2007 and
approximately 1.3 million HIV-negative patient fatalities as well as nearly half a million
deaths amongst HIV-positive populations (2). Even fifty years after the introduction of
powerful antibiotics to treat TB, it has been estimated that one person is infected in the
world every few seconds (3). The failure to control TB is due to the emergence of M.
tuberculosis strains that are multiply drug resistant towards the front line antimycobacterial
drugs such as isoniazid and rifampicin.
Phone: (718) 430-3096; Fax: (718) 430-8565.
*AUTHOR EMAIL ADDRESS: blanchar@aecom.yu.edu
#These authors contributed equally to this work.
Supporting Information Available
One figure showing the dependence of kburst on the [ertapenem] and [doripenem]. This material is available free of charge via the
Internet at http://pubs.acs.org
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 May 4.
Published in final edited form as:
Biochemistry. 2010 May 4; 49(17): 3766–3773. doi:10.1021/bi100232q.
NIH-PA Author Manuscript
NIH-PA Author Manuscript
NIH-PA Author Manuscript
As one of the most important antibiotic families, β-lactams include a broad range of
molecules including penicillin derivatives, cephalosporins, monobactams, carbapenems, and
β-lactamase inhibitors. The carbapenems exhibit the broadest spectrum of activity among
the β-lactam antimicrobials, providing safe and efficacious therapies in the treatment of
serious infections caused by Gram-positive, Gram-negative, and anaerobic bacterial
pathogens (4,5). Carbapenem antibiotics were originally developed from thienamycin, a
natural product identified in culture filtrates of Streptomyces cattleya (6). There are four
carbapenems approved thus far for human use: imipenem, meropenem, ertapenem, and
doripenem (5). Imipenem was the first carbapenem approved by the US Food and Drug
Administration (FDA) in 1985, and is by far the most widely used carbapenem. The use of
meropenem was approved in 1995, followed by ertapenem and doripenem in 2001 and 2007,
respectively. Except for imipenem, all carbapenems are stable against the mammalian
kidney dehydropeptidase (7). In clinical usage, imipenem and meropenem have to be given
frequently to maintain high circulating levels. Also, weight-dosage adjustment of imipenem
is required to minimize the chance of seizures (8). Ertapenem and doripenem can be given
once per day due to their high target affinity and circulating stability (5,9). The lower
effective doses of these latter drugs reduces potential side effects, as well as the
development of resistance (10). Currently, ertapenem and doripenem are used for
complicated intra-abdominal, and urinary tract infections (11,12).
Despite the general success of β-lactam antibiotics, they have not been widely used for the
treatment of TB due to intrinsic resistance that is caused by the presence of a
chromosomally-encoded gene (blaC) in M. tuberculosis for a Class A Ambler β-lactamase
(BlaC). Like other Class A β-lactamases, BlaC catalyzes the opening of the β- lactam ring
via nucleophilic attack by an active site serine residue to generate the acylenzyme, followed
by the hydrolysis of the ester bond to generate the ring-opened, inactive product. Our
previous studies of TB BlaC revealed that this enzyme is an extremely broad-spectrum β-
lactamase hydrolyzing all β-lactam classes, including the carbapenems meropenem and
imipenem (13). Being slow substrates that exhibit rapid acylation followed by a slow
deacylation step, meropenem and imipenem also act as potent inhibitors of BlaC (14).
FTICR mass spectrometry demonstrated that the acylated intermediate remains stable for
many minutes (14). Such slow turnover rates allowed the determination of three-
dimensional structure of BlaC in complex with meropenem at a resolution of 1.8 Å. In vivo
studies showed that meropenem in combination with the β-lactamase inhibitor, clavulante, is
bactericidal against clinical TB strains that are phenotypically exensively drug resistant
(XDR-TB) (14). As an extension of our prior work, we carried out an in vitro
characterization of doripenem and ertapenem with BlaC.
Materials and Methods
All chromatographic materials were purchased from Pharmacia. Meropenem and faropenem
were from IKT Laboratories. Doripenem (as Doribax) was from Ortho-McNeil
Pharmaceutical Inc (Raritan, NJ). Ertapenem (as Invanz) was from Merck & Co. Inc. The
potassium salt of clavulanic acid was from Sigma Aldrich. All other chemicals were
purchased from Sigma or Aldrich. Nitrocefin was purchased from Beckton Dickinson.
Purification of BlaC
Recombinant and truncated BlaC from M. tuberculosis expressed from plasmid pET28a(+)
and purified to homogeneity as described by Hugonnet and Blanchard (13).
Tremblay et al.
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NIH-PA Author Manuscript
NIH-PA Author Manuscript
Kinetics
The steady state rate of hydrolysis of β-lactam ring was monitored as a decrease in the
absorbance in the UV region, as described previously (13). Assays using doripenem,
ertapenem, faropenem and meropenem were performed at 296 nm (ε = 7,540 M−1 cm−1),
295 nm (ε = 9,970 M−1 cm−1), 306 nm (ε = 3,445 M−1 cm−1), and 297 nm (ε = 6,152 M−1
cm−1), respectively. Assays using the chromogenic substrate nitrocefin were performed at
486 nm (ε = 20,500 M−1 cm−1). Assays were performed in 100 mM MES (pH 6.5).
Reactions were initiated by the addition of enzyme at concentrations between 0.1–25 µM
using 100 µM of the carbapenem substrate.
Inhibition Studies
Carbapenems at concentrations ranging from 0.1–10 µM were tested as inhibitors of 1.5 nM
BlaC using 60 µM nitrocefin as substrate. Time courses were followed for 15 min. For slow
onset inhibition, reaction velocities as a function of time were fitted to eq 1:
(1)
where [P] is the concentration of the product, vi and vs are the initial and final reaction
velocities respectively for the reaction in the presence of inhibitor and kiso is the apparent
first order rate constant for the inter-conversion between vi and vs, and t is time.
The general mechanism can be modeled as:
(2)
where k1 and k−1 represent the reversible binding to and dissociation from the carbapenem
to BlaC, k2 represents the irreversible cleavage of the carbapenem β-lactam ring and k3
represents the hydrolysis of the BlaC-carbapenem adduct.
For this model, the rate constant that describes kiso is given by eq 3, where Kd equals k−1/k1.
(3)
In eq 4, the Km value can be expressed as:
(4)
In addition, from the determined k2 and k3 values, kcat is calculated from eq 7, assuming
k2,k3≪k1,k−1.
(5)
Tremblay et al.
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Mass Spectrometry
All mass spectra were acquired on a 9.6 T Fourier Transform Ion Cyclotron Resonance
(FTICR) mass spectrometer (Ionspec, Lake Forest, CA). To avoid salt interference, BlaC
was dialyzed against 20 mM ammonium bicarbonate, pH 6.5. The molecular mass of each
protein sample was determined for the 25+ charge state using the equation m = (m/z x
25)−25 on the isotopic centroid. To monitor the intermediate of steady state turnover or
small molecular mass spectrometry, 51 µM of enzyme was incubated with 25 µM
carbapenem in a total volume of 20 µL. An aliquot of 1 µL was withdrawn at desired time
(0, 30, 60, and 90 min) and mixed with 9 µL of mixing solution (containing 50% acetonitrile
and 0.1% formic acid). The resulting mixture was injected into the FTICR mass
spectrometer.
Crystallization
BlaC was crystallized in the hanging drop vapor diffusion configuration over well
conditions of 0.1 M HEPES, pH 7.5 and 2 M NH4H2PO4. The final pH of the well solution
was 4.1. Protein at a concentration of 10 mg/ml was mixed 1:1 with the well solution and
incubated at 18 °C. Initial crystals grew within a week but were small, sparse and
amorphous. New wells were sealed and allowed to equilibrate overnight. Equilibrated drops
were micro-seeded, which resulted in efficient crystal growth as well as improved
morphology. Iterative seeding resulted in diffraction quality crystals of active enzyme.
Data collection and refinement
Crystals were soaked with either ~ 50 mM ertapenem or doripenem in mother liquor plus
20% glycerol as a cryo-protectant. Data were collected after 10 and 90 minute soaks with
ertapenem and a 90 minute soak with doripenem at Brookhaven National Laboratory on
beamlines X12C and X29, in which various resolutions of diffraction were obtained
dependent on the soaking times and beamline. The data were processed using either
HKL2000 (15) or Mosflm (16). Our previous structure of clavulanate bound M. tuberculosis
β-lactamase (17) (PDB entry 3CG5) was used to phase all the data, using the CCP4 software
suite (18). Iterative rounds of structural refinement and model building were performed in
Refmac5 (19,20) and Coot (21). Table 1 lists the data collection statistics for the structures
as well as the final refinement statistics.
RESULTS and DISCUSSION
Kinetics
The accurate determination of the kinetic parameters for doripenem and ertapenem was
severely hampered by apparent very low Km values, very low kcat values and the modest
extinction coefficients accompanying hydrolysis. At the [BlaC] required to see any
significant rate of reaction (~2 µM), variation of the [doripenem] or [ertapenem] at
concentrations from 2–20 µM showed almost no difference in rate, suggesting their Km
values were less than 2 µM. The steady-state kinetic parameters determined for faropenem, a
structurally distinct penem, were Km = 55 ± 11 µM, and kcat = 0.65 ± 0.04 min−1 (data not
shown). This Km value is ~17 times larger and the kcat value 8 times faster than those of
meropenem (14).
Detailed investigations of the kinetics of carbapenem hydrolysis under near stoichiometric
enzyme concentrations were carried out over 30 minute time periods. As shown in Figure 1,
a steady-state kinetic burst was observed with both compounds where the magnitudes of the
burst are proportional to the concentration of BlaC used. Extrapolation of the rates of
hydrolysis to the y-axis demonstrates that the acylation is stoichiometric with the
concentration of enzyme.
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Due to the extremely feeble turnover rate, we further tested these carbapenems as inhibitors
of the reaction of nitrocefin with BlaC. Nitrocefin is an extremely good substrate for BlaC
and its β-lactam ring-opened form is extremely chromogenic. As shown in Figure 2,
doripenem and ertapenem act as slow-onset, tight binding inhibitors of BlaC when the
hydrolysis of nitrocefin was monitored. In contrast, faropenem exhibited standard,
competitive inhibition with no time dependent component (data not shown). This type of
time-dependent inhibition for a dead-end inhibitor is modeled as being due to the reversible
formation of a non-covalent complex (E–I), followed by the reversible conversion to an
isomerized complex (E–I*). However, in the case of a slow substrate for BlaC, the initially
formed Michaelis complex reacts with the enzyme in an irreversible step to generate the
BlaC-carbapenem covalent intermediate. This is then hydrolyzed slowly to regenerate the
free enzyme that can react with nitrocefin. While the same equation is used to fit the two
models, the kinetic constants that contribute to kiso (Figure S1) and Ki (or Kd) are different.
Using the fits of the slow-onset data and eq 3 to calculate Kd, k2 and k3, we can then used
eqs 4 and 5 to calculate the apparent Km and kcat values for doripenem (0.18 µM and 0.016
min−1, respectively), and for ertapenem (0.18 µM and 0.017 min−1, respectively). We have
not corrected for the concentration of nitrocefin used in these experiments because of the
large standard errors (>40%) associated with these kinetic parameters (the reported Km
values are apparent values). However, the extremely tight binding and extremely low
turnover of these carbapenems is evident from these rather imprecise kinetic data.
Mass Spectrometry
The rapid acylation and slow deacylation of BlaC by the carbapenems allows the
observation of the covalently bound, acyl-enzyme intermediate by Fourier transform ion
cyclotron resonance. A freshly prepared solution containing excess BlaC and doripenem
displayed three peaks: the first peak corresponds to free BlaC with mass/charge ratio (m/z) =
28,785.0, a second peak corresponding to the covalently acylated BlaC-doripenem complex
with mass/charge ratio (m/z) = 29, 204.1 and a third peak whose mass corresponds to the
mass of the covalently acylated BlaC-doripenem complex minus 44 mass unit (m/z = 29,
161.0), as shown in Figure 3. With ertapenem, the two covalent acylated BlaC complex
peaks observed had molecular masses of 29,260.0 and 29,217.1, corresponding to acylated
BlaC-ertapenem complex and acylated BlaC-doripenem complete minus 44 mass units,
respectively. This data demonstrates that both doripenem and ertapenem undergo the same
chemical breakdown in the active site as meropenem (14). Once the acyl-enzyme forms, the
carbapenems partition between hydrolysis and enzyme-catalyzed decomposition of the C6
hydroxyethyl substituent, via a retro-Aldol decomposition, which yields acetaldehyde (14).
Intriguingly, the intensities of the acyl-enzyme complexes remain stable over the time period
of 90 min for doripenem and ertapenem. This is in contrast with previous observations with
meropenem, where the acylated forms of the enzyme started to diminish after several
minutes. These data suggest that doripenem and ertapenem form more stable complexes
with BlaC than meropenem, reinforcing the kinetic data.
X-ray Crystallography
The 2.2 Å data from a 90 min doripenem-soaked crystal were refined to an Rwork of 0.161
and an Rfree of 0.205. The 1.3 Å diffraction data from a 10 minute ertapenem-soaked crystal
refined to an Rwork of 0.147 and an Rfree of 0.176. The 2.0 Å diffraction data from a 90 min
ertapenem-soaked crystal were refined to an Rwork of 0.175 and an Rfree of 0.222. In these
three structures, the active site Ambler residue Ser70 has been covalently linked with the
ring open form of these β-lactams in accordance with the acylation chemistry of the first half
of the enzymatic reaction (Scheme 1). The quality of the electron density is displayed in
Figure 4 and Figure 5 under a Fo-Fc omit calculated map contoured at 2.0 σ.
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The C3 atom of the pyrroline rings of doripenem and ertapenem covalent adducts are sp3
hybridized in the 90 min soaks. These results require an isomerization event occurring in the
BlaC-carbapenem adducts in which the original Δ2 pyrroline ring was tautomerized to
generate the Δ1 pyrroline ring, evidenced by the collinear positioning of the C5, N1, C2, and
C3 atoms. In addition, the BlaC-doripenem and ertapenem covalent adduct densities allow
for the positioning of the thioether sulfur atom in the unambiguous assignment of the S
configuration at C3, requiring protonation at the re face of the C2–C3 double bond. This is
similar to our earlier findings with BlaC crystals soaked with meropenem on similar time
scales (14), and represents the thermodynamically preferred product with the trans
orientation of the C4 methyl and thioether substituents. Interestingly, in the recently reported
structure of the Class D β-lactamase, OXA-1 covalent adduct with doripenem, revealed that
while an identical isomerization had taken place, that the final Δ1 pyrroline ring product was
of the opposite, R, stereochemistry (22).
In the structure determined after the shorter 10-minute soak, the BlaC-ertapenem adduct was
covalently bound in the active site, but in a different geometry. On this shorter time scale,
the BlaC-ertapenem adduct C3 atom is found in its original sp2 hybridization with the
definitive collinear positioning of the thioether sulfer atom in line with the N1, C2, C3, and
C4 bonds indicating the presence of the Δ2 pyrroline ring. This result requires that β-lactam
ring cleavage and isomerization of the methyl pyrroline ring not be concerted.
The active site interactions vary in some subtle ways between the pre and post-isomerization
ertapenem complexes, yet a number of common interactions are observed in all complexes.
Both the BlaC-ertapenem and -doripenem adducts bind as covalent adducts with the active
site Ser70 and position their lactam ring-opened ester carbonyl oxygen atom within the
oxyanion hole formed from hydrogen bonding interactions with the Ser70 and Thr253 amide
nitrogen atoms. All structures contain a hydrophobic interaction between the methyl group
of the pyrroline ring and the sidechain of Ile117 and different forms of hydrogen bonding
interactions between the C6 hydroxyethyl substituent of the carbapenem and Glu166. All
three structures also show a conserved interaction between the sidechain hydroxyl of
Ser130, which consistently hydrogen bonds the pyrroline ring nitrogen atom at a distance of
2.7–2.8 Å. The pyrroline C2 carboxylate group forms hydrogen bonds with the Thr251
hydroxyethyl side chain and an active site water molecule. In structures of other β-
lactamase-carbapenem adducts, this carboxylate electrostatically interacts with a conserved
arginine residue (R244 in TEM-1) (23), but this is not the case for BlaC. In the pre-
isomerized ertapenem structure, there is an additional hydrogen bond between the C2
carboxylate of the pyrroline ring and Thr253, which is broken upon the repositioning of the
meta-amino-benzoate ‘arm’ observed in the post-isomerized ertapenem structure. The
isomerization and stereospecific protonation leads to a reorientation of the terminal portion
of the molecule within the active site, allowing for the formation of the hydrogen bond
between the terminal carboxylate group of the meta-amino-benzoic acid moiety and Ser118.
A final difference between the initially formed Δ2-pyrroline isomer and the final Δ1-
pyrroline form is the orientation of the C6 hydroxyethyl substituent. In the pre-isomerized
complex it is oriented away from Lys73 and hydrogen bonds to the carboxyl O1 of Glu166
as well as the Asn186 nitrogen, but rotates upon isomerization to hydrogen bond with the
carboxyl O2 of Glu166 and the ε-amino group of Lys73 in a manner similar to that observed
for the doripenem complex.
The structures of the pre and post-isomerization states reveal the mechanistic basis for the
relative stability of the carbapenems within the active site of BlaC and their ability to resist
hydrolysis by the enzyme. As seen in Scheme1, deacylation-hydrolysis from the enzyme
requires the activation of the conserved active site water by the side chain carboxyl O2 of
Glu166. The probability of water activation by Glu166 decreases with the increased distance
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between the two. The distance between the carboxyl O2 of Glu166 and the active site water
is significantly increased after isomerization from 2.4 Å to 2.7 Å, making water activation
less probable. The isomerization event does not directly cause these changes but rather alters
the positioning of the adduct such that the adduct-hydroxyethyl flips in the active site,
breaking the hydrogen bond formed between the hydroxyl group with the side chain amide
nitrogen of Asn186 and the adjacent carboxyl O1 of Glu166 oxygen. The hydroxyethyl
substituent then rotates to generate a hydrogen bond network with Lys73 and the carboxyl
O2 of Glu166, effectively ‘pulling’ this essential base away from the conserved active site
water. These residues (Lys 73 and Glu166) are involved as general bases in the acylation
and deacylation reactions, respectively. In addition, by hydrogen bonding carboxyl O2 of
Glu166, the reoriented hydroxyethyl substituent reduced the basicity of Glu166. These two
factors, introduced in the reorientation of the hydroxyethyl substituent in the post-
isomerization complex, reduce water activation and thereby stabilize the acyl-intermediate
in the active site. The studies reported here allow us to directly visualize the changes that
occur between the pre- and post-isomerization adduct structures and are atomic level
observations relevant to the biphasic kinetics previously reported for the reactions between
carbapenems and the RTEM β-lactamase (24).
The Δ2 to Δ1-pyrroline isomerized forms of carbapenems have been known to form within
the active sites of various β-lactamase enzymes (25). In confirmation of this, crystal
structures of carbapenems bound within the active sites of the Class A β-lactamases TEM-1
(PDB entry 1BT5) (26) and SHV-1 (PDB entry 2ZD8) (27) as well as AmpC (PDB entry
1LL5) (28) a Class C β-lactamase all revealed the Δ2 form of the carbapenem bound in the
active sites, while the Class D OXA-1 (PDB entry 3ISG) (22) and the Class A BlaC (PDB
entry 3DWZ) (14) were both bound with carbapenems in the Δ1 isomerized forms with
respective R and S-stereochemistries. Our findings are the first to show the structures of
both the Δ2 and Δ1 forms of a carbapenem bound to a single β-lactamase. Interestingly,
several of the structures of carbapenems bound as the Δ2 isomers show evidence for
alternate conformations for the carbapenem-carbonyl oxygen position. This oxygen is found
buried within the oxyanion-hole as well as bound in a position rotated by 180 degrees,
usually facing an opposing serine residue (Ser130). In these instances it has been proposed
that the flipping of the carbonyl oxygen from the oxyanion-hole blocks formation of the
deacylation tetrahedral intermediate to inhibit the enzyme. In the cases of OXA-1 and BlaC,
the carbapenem-carbonyl oxygen is only found bound tightly within the oxyanion-hole and
no evidence of alternate conformers has been observed. In these cases inhibition by the
carbapenem is likely due to disruption of water activation.
A second possible reason for the observed alternate conformers at the carbapenem-carbonyl
is likely due to the position of the carbapenem-carboxylate moiety within those active sites.
To date, those β-lactamases with alternate conformations for the carbapenem-carbonyl, have
a highly conserved Arg244 reside which electrostatically interacts with the carbapenem-
carboxylate moiety. The OXA-1 and BlaC active sites lack this arginine interaction and
instead use a combination of threonine and/or serine residues coordinated with waters to
bind the carboxylate moiety. These residues are located closer to the oxyanion hole and act
to ‘clamp’ the carboxylate into a proximal position, as opposed to the Arg244 mechanism of
carboxylate binding, where distance introduces flexibility, allowing for the alternate
positioning of the pyrroline ring. This bonding pattern to the carbapenem allows for
alternate “in/out” conformations of the carbapenem carbonyl in the oxyanion hole. In
contrast, the carbapenem carbonyl is tightly bound in the oxyanion hole in BlaC in both the
Δ2 and Δ1 forms reported here.
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Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
This work was supported in part by a grant from the National Institute of Health (AI33696 to J. S. B.) and in part by
the Charles Revson Foundation (to L.W.T.)
ABBREVIATIONS
BlaC
Mycobacterium tuberculosis beta-lactamase
TB
Tuberculosis
XDR-TB
extensively drug resistant
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Figure 1.
Time courses of doripenem (A) and ertapenem (B) hydrolysis with various concentrations of
BlaC.
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Figure 2.
Time courses of nitrocefin hydrolysis by BlaC in the presence of doripenem (upper) and
ertapenem (lower).
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Figure 3.
Mass spectra of enzyme-carbapenem species. The 25+ charge state ions are shown.
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Figure 4.
(A) Overall structure of BlaC displayed in rainbow from N term (blue) to the C term (red),
with the doripenem adduct displayed in red surface mesh. (B) Fo-Fc omit density (green)
contoured at 2.0 σ surrounds the covalent doripenem adduct formed at the Ambler active-
site residue serine 70. All structure figures were produced using Pymol.
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Figure 5.
(A) Fo-Fc omit density (green) contoured at 2.0 σ surrounds the covalent ertapenem adduct
formed at the Ambler active-site residue serine 70 in the pre-isomerization state. (B) Fo-Fc
omit density (green) contoured at 2.0 σ surrounds the covalent ertapenem adduct formed at
the Ambler active-site residue serine 70 in the post-isomerization state. The resolution of the
densities unambiguously demonstrates the shift in stereochemistry with the change from sp2
to sp3 hybridization of the C3 carbapenem carbon atom with the change in the position of
the density associated with the meta-amino-benzoate and the hydoxyethyl ertapenem
moieties.
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Scheme 1.
(A) the structures of doripenem and ertapenem. (B) The chemical mechanism of hydrolysis
of ertapenem by the Mycobacterium tuberculosis BlaC.
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Table 1
Data Collection and Refinement Statistics
Data Collection
Doripenem
Δ1-isomerζ
Ertapenem
Δ2-isomer
Ertapenem
Δ1-isomer
Resolution (Å)
50.0-2.2
(2.32-2.20)
50.0-1.30
(1.33-1.30)
50.0-2.0
(2.07-2.00)
Completeness
100% (100%)
100.0% (100%)
99.5 (99.9)
Redundancy
7.6 (7.4)
7.5 (5.7)
4.4 (4.4)
I/sigma(I)
3.8 (1.6)
21.4 (1.8)
9.8 (4.0)
Rmerge
0.077 (0.47)
0.057 (0.757)
0.158 (0.373)
Space Group
P212121
P212121
P212121
Unit cell (Å)
a =49.989
b =68.068
c =75.792
α = β = γ = 90.0°
a = 49.66
b = 67.92
c = 75.55
α = β = γ = 90.0°
a =49.934
b =67.830
c =75.201
α = β = γ = 90.0°
Reflections
13,695 (1,943)
60,263 (4,388)
17,920 (1,762)
Refinement Statistics
Rwork
0.161 (0.176)
0.147 (0.265)
0.175 (0.191)
Rfree
0.205 (0.237)
0.176 (0.278)
0.222 (0.281)
Average B-factors (Å2)
Protein
6.97
10.49
6.09
Adduct
27.32
18.64
15.50
Solvent
17.51
32.64
14.36
PO4
12.89
14.40
10.53
RMS deviations
bonds (Å)
0.010
0.010
0.012
angles (°)
1.204
1.428
1.386
Ramachandra
Favored= 97.7%
outliers= 0.0%
Favored= 97.7%
outliers= 0.0%
Favored= 98.1%
outliers= 0.0%
PDB accession code
3IQA
3M6B
3M6H
Values in parentheses are for the highest resolution bin.
ζThis data processed using Mosflm
Biochemistry. Author manuscript; available in PMC 2011 May 4.
|
3M6C
|
Crystal structure of Mycobacterium tuberculosis GroEL1 apical domain
|
Structural and Functional Conservation of Mycobacterium
tuberculosis GroEL Paralogs Suggests that GroEL1 is a
Chaperonin
Bernhard Sielaff*, Ki Seog Lee*,2, and Francis T.F. Tsai1
Verna and Marrs McLean Department of Biochemistry and Molecular Biology, and Department of
Molecular and Cellular Biology, Baylor College of Medicine, One Baylor Plaza, Houston, Texas
77030, USA
Abstract
GroEL is a group I chaperonin that facilitates protein folding and prevents protein aggregation in
the bacterial cytosol. Mycobacteria are unusual in encoding two or more copies of GroEL in their
genome. While GroEL2 is essential for viability and likely functions as the general housekeeping
chaperonin, GroEL1 is dispensable but its structure and function remain unclear.
Here we present the 2.2 Å resolution crystal structure of a 23 kDa fragment of Mycobacterium
tuberculosis GroEL1 consisting of an extended apical domain. Our X-ray structure of the GroEL1
apical domain closely resembles those of Escherichia coli GroEL and M. tuberculosis GroEL2;
thus, highlighting the remarkable structural conservation of bacterial chaperonins. Notably, in our
structure, the proposed substrate-binding site of GroEL1 interacts with the N-terminal region of a
symmetry related, neighboring GroEL1 molecule. The latter is consistent with the known GroEL
apical domain function in substrate binding, and is supported by results obtained from using
peptide array technology. Taken together, we show that the apical domains of M. tuberculosis
GroEL paralogs are conserved in three-dimensional structure, suggesting that GroEL1, like
GroEL2, is a chaperonin.
Keywords
Molecular chaperones; Hsp60; apical domain; protein folding; KasA
Introduction
Molecular chaperones assist protein folding by facilitating the productive folding of newly
synthesized polypeptides and by preventing protein aggregation in the crowded environment
© 2010 Elsevier Ltd. All rights reserved.
1Corresponding author: Department of Biochemistry, Baylor College of Medicine, One Baylor Plaza, MS: BCM125, Houston,
Texas 77030. ftsai@bcm.edu; Phone: +1-713-798-8668; Fax: +1-713-796-9436.
*These authors contributed equally to this study.
2Present address: Department of Clinical Laboratory Science, College of Health Science, Catholic University of Pusan, Pusan
609-757, Republic of Korea.
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Author contributions
B.S. designed and performed research, analyzed data, and wrote the paper. K.S.L. performed research, analyzed data, and wrote the
paper. F.T.FT. designed research, analyzed data, and wrote the paper.
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Published in final edited form as:
J Mol Biol. 2011 January 21; 405(3): 831–839. doi:10.1016/j.jmb.2010.11.021.
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of the cell.1 Escherichia coli GroEL is a group I chaperonin that assembles into an 800 kDa
homo-tetradecamer composed of two heptameric rings that are stacked back-to-back.2; 3
Each GroEL subunit has a molecular weight of 57 kDa and consists of an equatorial, an
intermediate, and an apical domain.3 The equatorial domain contains the ATP-binding site
and mediates contacts between subunits in the cis and trans rings. The intermediate domain
functions as a hinge that connects the equatorial domain to the apical domain. The latter
forms the entrance to the GroEL cavity and is involved in GroES binding4 as well as
polypeptide recognition.5; 6 It has been suggested from X-ray crystallographic studies that
helices H and I of the E. coli GroEL apical domain form the substrate binding site.7; 8; 9
Interestingly, many mycobacteria contain genes encoding two or more GroEL paralogs.10
GroEL1 and GroEL2 from the human pathogen Mycobacterium tuberculosis11; 12 are of
particular interest as both proteins are involved in the host immune response to M.
tuberculosis infection.13; 14 While GroEL2 is essential and likely functions as the principal
housekeeping chaperonin,10; 11 GroEL1 is non-essential and is dispensable for viability. It
has been proposed that M. tuberculosis GroEL1 is a nucleoid-associated protein,15 and that
the closely related GroEL1 ortholog from M. smegmatis plays a role in biofilm formation by
modulating mycolic acid biosynthesis through direct interaction with the β-ketoacyl ACP
synthase KasA.10
Like other bacterial chaperonins, M. tuberculosis GroEL1 and GroEL2 are up-regulated
upon heat shock16 as well as in response to oxidative stress,17 indicating that both copies
may have chaperone activity inside cells. In contrast, recombinant GroEL1 and GroEL2
overexpressed in E. coli exist as dimers, and exhibit low ATPase and no folding activities.18
Since native GroEL1 forms higher-order oligomers in M. tuberculosis cells,19 lack of
chaperone activity might be attributed to the inability of the recombinant proteins to self-
assemble.
Consistent with its essential cellular role, the X-ray structure of a M. tuberculosis GroEL2
dimer20 showed that the GroEL2 monomer has the same fold as E. coli GroEL,20
supporting the notion that GroEL2 is a chaperonin. However, at present, no high-resolution
structural information is available for M. tuberculosis GroEL1, and its structure-function
relationship remains unclear.
Here we present the 2.2 Å resolution crystal structure of a 23 kDa M. tuberculosis GroEL1
fragment consisting of the GroEL1 apical domain flanked by flexible segments that are part
of the intermediate domain. This structure is hereafter referred to as the GroEL1 apical
domain. We found that the atomic structure of the GroEL1 apical domain is very similar to
those of M. tuberculosis GroEL220 and E. coli GroEL.7; 8 Fortuitously, in our crystal
structure, the N-terminus of one molecule interacts with the putative GroEL substrate-
binding site of a symmetry related molecule. This interaction is reminiscent of the X-ray
structures of E. coli chaperonin-substrate peptide complexes.7; 8; 9 Moreover, we found
using peptide array technology that both full-length M. tuberculosis GroEL1 and the isolated
GroEL1 apical domain recognize the same peptide motifs present in the M. tuberculosis
KasA sequence, which resemble binding motifs reported for E. coli GroEL.21 Thus, our
combined structural and functional data suggest that M. tuberculosis GroEL1, like GroEL2,
is a chaperonin and support the notion that the apical domain is sufficient for substrate
interaction.
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Results and Discussion
Crystal Structure of the M. tuberculosis GroEL1 Apical Domain
Crystals of the GroEL1 apical domain (residues 184–377) diffracted to 2.2 Å resolution on a
home X-ray source, and belonged to the orthorhombic space group P21212 with one
molecule in the asymmetric unit. After rigid body refinement of the initial model in CNS22
the resulting electron density map was readily interpretable. Several rounds of model
building, simulated annealing, least-squares positional, and individual B-factor refinement
were performed resulting in a final model with a Rfactor and Rfree 23 of 21.0% and 23.2%,
respectively. The final model consists of 194 residues and 65 water molecules. The
Ramachandran plot showed that 91.1% of all residues were in the most favored regions and
8.9% in additional allowed regions. No residue was in a generously allowed or disallowed
region. The refinement statistics are summarized in Table 1.
The structure of the M. tuberculosis GroEL1 apical domain consists of a β-sandwich
scaffold flanked by several α-helices and loops (Fig. 1a and b). Structural comparison of the
M. tuberculosis GroEL1 apical domain with those of M. tuberculosis GroEL2 (PDB ID:
1SJP-A)20 and E. coli GroEL (PDB ID: 1KID and 1DKD-A)7; 8 showed that they are very
similar (Fig. 1c). The Cα atoms of the refined M. tuberculosis GroEL1 apical domain
structure (residues 191–372) superimposed pairwise with those of M. tuberculosis GroEL2
residues 190–371 (PDB ID: 1SJP-A) 20 with a root mean square deviation (RMSD) of 0.82
Å, and with those of E. coli GroEL residues 193–375 (PDB ID: 1KID)7 and residues 193–
336 (PDB ID: 1DKD-A)8 with a RMSD of 0.63 Å and 0.75 Å, respectively.
Protein-protein interactions between symmetry related GroEL1 molecules
Our crystal structure showed two types of protein interfaces with crystallographic symmetry
related GroEL1 apical domain molecules in the crystal lattice. While the observed contacts
may hint at the existence of higher oligomers, none of the observed interactions are
consistent with the known interfaces in the E. coli GroEL tetradecamer structure.2; 3 It has
been proposed that both recombinant, full-length M. tuberculosis GroEL1 and GroEL2 form
dimers18 through inter-subunit contacts between apical domains.20 While our biochemical
results support the notion that recombinantly expressed, full-length GroEL1 is a dimer, we
found that the isolated GroEL1 apical domain is a monomer in solution (Fig. 1d).
In our structure, the N-terminal residues 184 to 187 with sequence Glu-Leu-Glu-Phe interact
with helices H and I of a symmetry-related neighboring molecule (Fig. 2a and b). Notably,
the Leu185 side-chain is in van der Waals contact with the side-chains of Leu232, Leu235,
Ala239, and Leu269, which form a hydrophobic pocket (Fig. 2b and 3a). In addition, there is
a network of hydrogen bond interactions between the polar side chains of Asn263 and
Arg266 and the main chain carbonyl oxygens of Glu184, Leu185, and Glu186 (Fig. 2b and
3a). These interactions are reminiscent of the previously observed E. coli GroEL apical
domain contacts with different peptides (Fig. 3b–c).7; 8; 9
GroEL1 substrate binding site
It has been proposed that the apical domain of E. coli GroEL contains the substrate binding
site.5; 6 This is supported by crystal structures of the E. coli GroEL apical domain bound to
a strong binding peptide selected by phage-display,8 and to an extended N-terminal segment
of a symmetry related molecule,7 respectively. In the latter two structures, protein-protein
interactions were mediated by both non-polar and hydrogen bond interactions with helices H
and I of the GroEL apical domain.7; 8 Although the sequence of the bound peptides differed
in each case, the two previously reported X-ray structures of the E. coli GroEL apical
domain7; 8 and our atomic structure of the M. tuberculosis GroEL1 apical domain display
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common binding features. First, the hydrophobic pocket is lined by the conserved residues
Leu232, Leu235, and Ala239 from helix H and the less conserved Leu269, which
correspond to Leu234, Leu237, Ala241, and Val271 in E. coli GroEL (Fig. 3a–c). Second,
hydrogen bond interactions mainly involve residues from helix I (Fig. 3a–c). Third,
hydrogen bonds are exclusively formed between the side chains of GroEL and the main
chain of the bound peptide (Fig. 3a–c). Notably, in our structure, only four residues (Glu-
Leu-Glu-Phe) contact the substrate binding site of the apical domain, which may represent
the minimal binding region.
GroEL1-KasA peptide interaction
To gain further insight into substrate recognition by M. tuberculosis GroEL1, we
synthesized a miniaturized peptide array of overlapping 12-mer peptides derived from the
M. tuberculosis KasA amino acid sequence. This work followed from an earlier report
demonstrating a direct interaction between KasA and GroEL1 in M. smegmatis cells,10
which serves as a model organism for different mycobacterial species, including M.
tuberculosis. Probing our KasA-derived peptide array with full-length GroEL1 showed
binding to 67 out of 192 peptides (Fig. 4a). Eleven of those peptides (C01, C14, C16 – C18,
E21 – E24, G13, and G15) also bound to a control protein and, therefore, were excluded
from further analysis (data not shown). Remarkably, we found that the isolated GroEL1
apical domain bound to the same peptides as full-length GroEL1, suggesting that the apical
domain is sufficient for substrate binding, although, some spots differed in intensity
indicating different binding affinities (Fig. 4a and b). Moreover, the same KasA peptides, in
addition to other KasA motifs, were also recognized by full-length GroEL2 in a control
experiment (Fig. S1).
Consecutive binding peptides with at least three members were grouped together to identify
consensus motifs within each group (Table 2). A consensus motif was defined as an amino
acid sequence that was present in most members of a group, and was at least four amino acid
residues long. Only one acidic residue (Asp) was found in all consensus motifs (Table 2;
Group B), which is very low compared to the abundance of negatively charged residues (14
%) in the KasA sequence. On the other hand, positively charged (Arg), and hydrophobic
residues (Val and Met) were enriched 2 to 2.6-fold in the consensus motifs relative to the
full-length sequence. The latter is in good agreement with previous studies of peptide
sequences bound by E. coli GroEL, which also revealed a strong preference for hydrophobic
and positively charged residues.21; 24; 25; 26
To provide a spatial explanation for the location of our identified peptides, we mapped our
consensus motifs onto the available X-ray structure of M. tuberculosis KasA27 (Fig. 4c).
We found that six of our consensus motifs (Groups A, D, G, H, I, J) are mostly buried in the
native protein structure, as it might be expected for a chaperonin substrate.28 However,
other consensus motifs (Groups B, C, E, and F) are solvent-exposed (Fig. 4c), suggesting
that GroEL1 may also interact with native, folded KasA. Taken together, our findings
suggest that M. tuberculosis GroEL1 and GroEL2 are chaperonins that recognize both
distinct and overlapping KasA peptide motifs. How GroEL1 modulates fatty-acid synthesis
through interaction with KasA remains unclear and is subject to further investigation.
Materials and Methods
Protein preparation and analysis
Cloning, expression, and purification of full-length GroEL1 from M. tuberculosis H37Rv, as
well as the crystallization and preliminary crystallographic analysis of the GroEL1 fragment
have been described.29 For peptide array analysis, the apical domain of GroEL1 (Glu188 to
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Val373) was cloned by PCR into pProEx Htb generating a plasmid that harbors the GroEL1
apical domain with a Tobacco Etch Virus protease-cleavable N-terminal His6-tag.
Expression and purification of the GroEL1 apical domain was performed as previously
described for full-length GroEL1.29 GroEL2 was cloned, overexpressed, and purified in a
similar manner, except for addition of a complete EDTA-free protease inhibitor cocktail
(Roche Diagnostics) in the lysis buffer.
To determine the oligomeric state, purified full-length GroEL1 and GroEL1 apical domain
were analyzed at 4 °C by size-exclusion chromatography on a Superdex 200 HR 10/30
column (GE Healthcare) in 50 mM Tris pH 7.5 and 150 mM NaCl. Molecular weight
protein standards (BioRad) were run in the same buffer in order to correlate elution volume
with protein size.
X-ray crystallographic analysis and structure refinement
The structure of the GroEL1 apical domain was determined by the molecular replacement
technique using the apical domain (residues 188 to 372) of a M. tuberculosis GroEL2
monomer (PDB ID: 1SJP-A)20 as search model. This model shares 64% sequence identity
with the M. tuberculosis GroEL1 apical domain, distributed evenly over the amino acid
sequence (Fig. 1a). 5% of the observed data were randomly chosen and excluded from
refinement for cross-validation purposes. The model was refined in CNS 1.2.22 Model
refinement was interspersed by manual rebuilding of the atomic structure using COOT.30
Water molecules were selected automatically in CNS 1.2,22 and confirmed manually
according to the peak height and distance criteria in the calculated Fo-Fc and 2Fo-Fc maps.
The stereochemistry of the final model was analyzed using PROCHECK.31
Superpositioning of molecules was done in PyMol32, the SSAP Server33 was used for
RMSD calculations, and ESPript34 was used for preparing the secondary structure
alignment.
Peptide array synthesis
A peptide array of 12-mer overlapping peptides derived from the amino acid sequence of M.
tuberculosis KasA was prepared by the SPOT synthesis technique using a semi-automated
ASP 222 peptide synthesis robot (Intavis) essentially as described.35 The sequence was
walked through by advancing two to three amino acids at each position in order to fit two
complete sets of peptides onto one membrane. The membrane was cut into halves, blocked
for 2 h in 1× Pierce Superblock in TBS-1 (20 mM Tris-HCl pH 7.5, 137 mM NaCl, and 0.1
% Tween-20), and washed for 10 min in blocking buffer consisting of 10% Superblock and
5% sucrose in TBS-2 (20 mM Tris-HCl pH 7.5, 137 mM NaCl, and 0.05 % Tween-20).
Next, each membrane half was incubated in blocking buffer for 1 h in the presence of either
500 nM His6-tagged GroEL1 or 500 nM His6-tagged GroEL1 apical domain. After washing
the membranes three times in TBS-1, bound protein was electro-transferred to a Hybond-
ECL nitrocellulose membrane (GE Healthcare), and probed and detected as previously
described.35 In addition, membranes were also probed directly with 50 ng/ml of a
commercially available, anti-His6 monoclonal antibody-horse radish peroxidase conjugate
(BD Biosciences) for 1 h in blocking buffer without sucrose. Despite slight differences in
spot intensities, both methods generated identical results. After detection, bound protein and
antibody were stripped off the membrane by washing the membrane three times in 8 M urea,
1% SDS, 0.5% 2-mercaptoethanol for 30 min each, followed by three times in 20% acetic
acid and 50% ethanol for 15 min each, and three times in 100% ethanol for 10 min each. A
negative control was performed with His6-tagged GrpE which was expressed and purified as
described.36 After stripping the membrane, another control was performed with His6-tagged
GroEL2. All incubations were carried out under gentle rocking at room temperature.
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PDB accession code
The atomic coordinates and corresponding structure factors have been deposited in the
Protein Data Bank under PDB ID: 3M6C.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We thank Dr. T. Palzkill for advice and suggestions, and Dr. A. Reger for help with X-ray data collection. Work in
F.T.F.T.'s laboratory is supported by grants from the National Institutes of Health (R01-AI076239), the Welch
Foundation (Q-1530), the Department of Defense, and the American Cancer Society. B.S. was a Welch
postdoctoral fellow and a recipient of a training fellowship from the Pharmacoinformatics Training Program of the
Keck Center of the Gulf Coast Consortia (NIH Grant No. R90-DK071505).
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Fig. 1.
Bacterial chaperonins are structurally conserved. (a) Sequence alignment of the apical
domains of M. tuberculosis (Mt) GroEL1, Mt GroEL2, and E. coli (Ec) GroEL. Secondary
structure elements of M. tuberculosis GroEL1 are represented by helices (α- and η [310]-
helices) and arrows (β-strands). Conserved residues are boxed. Nomenclature of α-helices
(H–L) is the same as previously described for the E. coli GroEL apical domain.8 (b) Stereo
view of the M. tuberculosis GroEL1 apical domain. The crystal structure is shown as ribbon
diagram with α-helices colored blue, β-strands red, and the N- and C-terminal extensions
and loops green. η denotes a 310 helix. (c) Superposition of the apical domain structures
from M. tuberculosis GroEL1 (orange), M. tuberculosis GroEL2 (magenta),20 and E. coli
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GroEL residues 191–376 (blue)7 and residues 191–336 (cyan).8 (d) Analysis of full-length
GroEL1 (blue) and GroEL1 apical domain (magenta) by size-exclusion chromatography.
The chromatogram shows that the full-length protein elutes as a dimer, whereas the apical
domain is a monomer.
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Fig. 2.
Structure of the GroEL1 substrate-binding site. (a) Ribbon diagram of the GroEL1 apical
domain (orange) and that of a crystallographic symmetry related molecule (green). The
putative substrate binding site is indicated by a purple box. (b) Close-up stereo view of the
substrate binding site in GroEL1. Interacting residues from helices H and I (orange) and
from the N-terminus of a crystallographic symmetry related molecule (green) are depicted as
ball-and-stick models. Hydrogen bonds are indicated by dashed lines and are shown together
with the corresponding bond lengths.
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Fig. 3.
Schematic representation of molecular interactions at the putative chaperonin-substrate
peptide interface in M. tuberculosis GroEL1 and E. coli GroEL. (a) M. tuberculosis GroEL1
apical domain bound to an N-terminal segment of GroEL1, (b) E. coli GroEL apical domain
bound to a non-native peptide,7 and (c) E. coli GroEL apical domain bound to a strong
binding peptide selected by phage display.8 Gray lines indicate van der Waals contacts, blue
arrows represent hydrogen bonds. Residues surrounded by a dotted box are located on a loop
immediately downstream of helix I.
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Fig. 4.
GroEL1 apical domain is sufficient for substrate recognition. Binding of (a) full-length M.
tuberculosis GroEL1 or (b) the isolated GroEL1 apical domain to a miniaturized peptide
array derived from the M. tuberculosis KasA sequence. Peptides belonging to the same
group are boxed together. Groups are differentiated by the color of boxes: Group A (red),
Group B (blue), Group C (yellow), Group D (green), Group E (orange), Group F (purple),
Group G (white), Group H (pink), Group I (brown), and Group J (teal). (c) Surface
representation of the KasA dimer structure (PDB ID: 2WGF)37 with the KasA monomers
displayed in different hues. Consensus motifs are mapped onto the KasA structure using the
same color scheme as in Fig. 4a and b.
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Table 1
Summary of Data Collection and Refinement Statistics
M. tuberculosis GroEL1 apical domain
Diffraction data statistics
Resolution (Å)
40.0 – 2.2 (2.28 – 2.2)
Space group
P 21212
Unit cell parameters
a = 75.47 Å, b = 78.65 Å, c = 34.89 Å
α = β = γ = 90°
No. of unique reflections
10,994
Completeness (%)
98.7 (91.9)
Redundancy
4.3 (2.6)
I/sigma (I)
12.3 (2.4)
Rsym (%)b
10.1 (36.7)
Refinement statistics
Resolution (Å)
20.0 – 2.2
Rfactor / Rfree (%)c
21.0 / 23.2
Average B-factor (Å2)
30.3
Number of atoms
Protein
1461
Water
65
RMSD of ideal bond length/angle
Bond length (Å)
0.006
Bond angle (°)
1.2
aValues in parentheses are for the highest resolution shell.
bRsym = Σ |I − (I)| / Σ (I), where I is the observed intensity and (I) is the average intensity.
cRfactor Σ |Fobs − Fcalc| / Σ |Fobs|, where Fobs are the observed structure factors and Fcalc are the calculated structure factors. The
crystallographic Rfactor is based on 95% of the data used in refinement, and the Rfree is based on 5% of the data withheld for cross-validation test.
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Table 2
KasA peptides bound by GroEL1
Group
Spots
Sequencea
A
A05-A08
9GGFPSVVVTAVTATTSIS26
B
A23-B01
49EFVTKWDLAVKIGGHL64
C
B09-B17
68DSHMDGRLDMRRMSYVQRMGKLLGGQLWE86
D
C02-C04
103VDPDRFAVVVGTGLGG118
E
C23-D07
145IMPNGAAAVIGLQLGARAGVMTPVSACS172
F
D23-E01
205LPIAAFSMMRAMSTRN220
G
F01-F04
257RGAKPLARLLGAGITSDA274
H
F12-F17
279APAADGVRAGRAMTRSLELAGL300
I
G20-G24
349AVGALESVLTVLTLRDGVIP368
J
H17-H19
391YGDYRYAVNNSFGFGG406
aConsensus motifs are depicted in bold. Lower case numbers indicate the position in the M. tuberculosis KasA sequence.
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|
3M6H
|
Crystal Structure of Post-isomerized Ertapenem Covalent Adduct with TB B-lactamase
|
Biochemical and Structural Characterization of Mycobacterium
tuberculosis β-Lactamase (BlaC) with the Carbapenems
Ertapenem and Doripenem
Lee W. Tremblay#, Fan Fan#, and John S Blanchard‡,*
Department of Biochemistry, Albert Einstein College of Medicine, 1300 Morris Park Avenue,
Bronx, New York 10461
Abstract
Despite the enormous success of β-lactams as broad-spectrum antibacterials, they have never been
widely used for the treatment of TB due to intrinsic resistance that is caused by the presence of a
chromosomally-encoded gene (blaC) in Mycobacterium tuberculosis. Our previous studies of TB
BlaC revealed that this enzyme is an extremely broad-spectrum β-lactamase hydrolyzing all β-
lactam classes. Carbapenems are slow substrates that acylate the enzyme but are only slowly
deacylated and can therefore act also as potent inhibitors of BlaC. We carried out the in vitro
characterization of doripenem and ertapenem with BlaC. A steady-state kinetic burst was observed
with both compounds with magnitudes proportional to the concentration of BlaC used. The results
show apparent Km and kcat values of 0.18 µM and 0.016 min−1 for doripenem and 0.18 µM and
0.017 min−1 for ertapenem. FTICR mass spectrometry demonstrated that the doripenem and
ertapenem acyl-enzyme complexes remain stable over a time period of 90 min. The BlaC-
doripenem covalent complex obtained after 90 minutes of soaking was solved to 2.2 Å, while the
BlaC-ertapenem complex obtained after a 90 minute soak was solved to 2.0 Å. The 1.3 Å
diffraction data from a 10 minute ertapenem-soaked crystal revealed an isomerization occurring in
the BlaC-ertapenem adduct in which the original Δ2 pyrroline ring was tautomerized to generate
the Δ1 pyrroline ring. The isomerization leads to the flipping of the carbapenem-hydroxyethyl
group to hydrogen bond to the carboxyl O2 of Glu166. The hydroxyethyl flip results in both
decreased basicity of Glu166 and in a significant increase in the distance between the carboxyl O2
of Glu166 and the catalytic water molecule, slowing hydrolysis.
Tuberculosis (TB), caused by Mycobacterium tuberculosis, continues to be a worldwide
health concern (1). There were an estimated 9.3 million new cases of TB in 2007 and
approximately 1.3 million HIV-negative patient fatalities as well as nearly half a million
deaths amongst HIV-positive populations (2). Even fifty years after the introduction of
powerful antibiotics to treat TB, it has been estimated that one person is infected in the
world every few seconds (3). The failure to control TB is due to the emergence of M.
tuberculosis strains that are multiply drug resistant towards the front line antimycobacterial
drugs such as isoniazid and rifampicin.
Phone: (718) 430-3096; Fax: (718) 430-8565.
*AUTHOR EMAIL ADDRESS: blanchar@aecom.yu.edu
#These authors contributed equally to this work.
Supporting Information Available
One figure showing the dependence of kburst on the [ertapenem] and [doripenem]. This material is available free of charge via the
Internet at http://pubs.acs.org
NIH Public Access
Author Manuscript
Biochemistry. Author manuscript; available in PMC 2011 May 4.
Published in final edited form as:
Biochemistry. 2010 May 4; 49(17): 3766–3773. doi:10.1021/bi100232q.
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As one of the most important antibiotic families, β-lactams include a broad range of
molecules including penicillin derivatives, cephalosporins, monobactams, carbapenems, and
β-lactamase inhibitors. The carbapenems exhibit the broadest spectrum of activity among
the β-lactam antimicrobials, providing safe and efficacious therapies in the treatment of
serious infections caused by Gram-positive, Gram-negative, and anaerobic bacterial
pathogens (4,5). Carbapenem antibiotics were originally developed from thienamycin, a
natural product identified in culture filtrates of Streptomyces cattleya (6). There are four
carbapenems approved thus far for human use: imipenem, meropenem, ertapenem, and
doripenem (5). Imipenem was the first carbapenem approved by the US Food and Drug
Administration (FDA) in 1985, and is by far the most widely used carbapenem. The use of
meropenem was approved in 1995, followed by ertapenem and doripenem in 2001 and 2007,
respectively. Except for imipenem, all carbapenems are stable against the mammalian
kidney dehydropeptidase (7). In clinical usage, imipenem and meropenem have to be given
frequently to maintain high circulating levels. Also, weight-dosage adjustment of imipenem
is required to minimize the chance of seizures (8). Ertapenem and doripenem can be given
once per day due to their high target affinity and circulating stability (5,9). The lower
effective doses of these latter drugs reduces potential side effects, as well as the
development of resistance (10). Currently, ertapenem and doripenem are used for
complicated intra-abdominal, and urinary tract infections (11,12).
Despite the general success of β-lactam antibiotics, they have not been widely used for the
treatment of TB due to intrinsic resistance that is caused by the presence of a
chromosomally-encoded gene (blaC) in M. tuberculosis for a Class A Ambler β-lactamase
(BlaC). Like other Class A β-lactamases, BlaC catalyzes the opening of the β- lactam ring
via nucleophilic attack by an active site serine residue to generate the acylenzyme, followed
by the hydrolysis of the ester bond to generate the ring-opened, inactive product. Our
previous studies of TB BlaC revealed that this enzyme is an extremely broad-spectrum β-
lactamase hydrolyzing all β-lactam classes, including the carbapenems meropenem and
imipenem (13). Being slow substrates that exhibit rapid acylation followed by a slow
deacylation step, meropenem and imipenem also act as potent inhibitors of BlaC (14).
FTICR mass spectrometry demonstrated that the acylated intermediate remains stable for
many minutes (14). Such slow turnover rates allowed the determination of three-
dimensional structure of BlaC in complex with meropenem at a resolution of 1.8 Å. In vivo
studies showed that meropenem in combination with the β-lactamase inhibitor, clavulante, is
bactericidal against clinical TB strains that are phenotypically exensively drug resistant
(XDR-TB) (14). As an extension of our prior work, we carried out an in vitro
characterization of doripenem and ertapenem with BlaC.
Materials and Methods
All chromatographic materials were purchased from Pharmacia. Meropenem and faropenem
were from IKT Laboratories. Doripenem (as Doribax) was from Ortho-McNeil
Pharmaceutical Inc (Raritan, NJ). Ertapenem (as Invanz) was from Merck & Co. Inc. The
potassium salt of clavulanic acid was from Sigma Aldrich. All other chemicals were
purchased from Sigma or Aldrich. Nitrocefin was purchased from Beckton Dickinson.
Purification of BlaC
Recombinant and truncated BlaC from M. tuberculosis expressed from plasmid pET28a(+)
and purified to homogeneity as described by Hugonnet and Blanchard (13).
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Kinetics
The steady state rate of hydrolysis of β-lactam ring was monitored as a decrease in the
absorbance in the UV region, as described previously (13). Assays using doripenem,
ertapenem, faropenem and meropenem were performed at 296 nm (ε = 7,540 M−1 cm−1),
295 nm (ε = 9,970 M−1 cm−1), 306 nm (ε = 3,445 M−1 cm−1), and 297 nm (ε = 6,152 M−1
cm−1), respectively. Assays using the chromogenic substrate nitrocefin were performed at
486 nm (ε = 20,500 M−1 cm−1). Assays were performed in 100 mM MES (pH 6.5).
Reactions were initiated by the addition of enzyme at concentrations between 0.1–25 µM
using 100 µM of the carbapenem substrate.
Inhibition Studies
Carbapenems at concentrations ranging from 0.1–10 µM were tested as inhibitors of 1.5 nM
BlaC using 60 µM nitrocefin as substrate. Time courses were followed for 15 min. For slow
onset inhibition, reaction velocities as a function of time were fitted to eq 1:
(1)
where [P] is the concentration of the product, vi and vs are the initial and final reaction
velocities respectively for the reaction in the presence of inhibitor and kiso is the apparent
first order rate constant for the inter-conversion between vi and vs, and t is time.
The general mechanism can be modeled as:
(2)
where k1 and k−1 represent the reversible binding to and dissociation from the carbapenem
to BlaC, k2 represents the irreversible cleavage of the carbapenem β-lactam ring and k3
represents the hydrolysis of the BlaC-carbapenem adduct.
For this model, the rate constant that describes kiso is given by eq 3, where Kd equals k−1/k1.
(3)
In eq 4, the Km value can be expressed as:
(4)
In addition, from the determined k2 and k3 values, kcat is calculated from eq 7, assuming
k2,k3≪k1,k−1.
(5)
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Mass Spectrometry
All mass spectra were acquired on a 9.6 T Fourier Transform Ion Cyclotron Resonance
(FTICR) mass spectrometer (Ionspec, Lake Forest, CA). To avoid salt interference, BlaC
was dialyzed against 20 mM ammonium bicarbonate, pH 6.5. The molecular mass of each
protein sample was determined for the 25+ charge state using the equation m = (m/z x
25)−25 on the isotopic centroid. To monitor the intermediate of steady state turnover or
small molecular mass spectrometry, 51 µM of enzyme was incubated with 25 µM
carbapenem in a total volume of 20 µL. An aliquot of 1 µL was withdrawn at desired time
(0, 30, 60, and 90 min) and mixed with 9 µL of mixing solution (containing 50% acetonitrile
and 0.1% formic acid). The resulting mixture was injected into the FTICR mass
spectrometer.
Crystallization
BlaC was crystallized in the hanging drop vapor diffusion configuration over well
conditions of 0.1 M HEPES, pH 7.5 and 2 M NH4H2PO4. The final pH of the well solution
was 4.1. Protein at a concentration of 10 mg/ml was mixed 1:1 with the well solution and
incubated at 18 °C. Initial crystals grew within a week but were small, sparse and
amorphous. New wells were sealed and allowed to equilibrate overnight. Equilibrated drops
were micro-seeded, which resulted in efficient crystal growth as well as improved
morphology. Iterative seeding resulted in diffraction quality crystals of active enzyme.
Data collection and refinement
Crystals were soaked with either ~ 50 mM ertapenem or doripenem in mother liquor plus
20% glycerol as a cryo-protectant. Data were collected after 10 and 90 minute soaks with
ertapenem and a 90 minute soak with doripenem at Brookhaven National Laboratory on
beamlines X12C and X29, in which various resolutions of diffraction were obtained
dependent on the soaking times and beamline. The data were processed using either
HKL2000 (15) or Mosflm (16). Our previous structure of clavulanate bound M. tuberculosis
β-lactamase (17) (PDB entry 3CG5) was used to phase all the data, using the CCP4 software
suite (18). Iterative rounds of structural refinement and model building were performed in
Refmac5 (19,20) and Coot (21). Table 1 lists the data collection statistics for the structures
as well as the final refinement statistics.
RESULTS and DISCUSSION
Kinetics
The accurate determination of the kinetic parameters for doripenem and ertapenem was
severely hampered by apparent very low Km values, very low kcat values and the modest
extinction coefficients accompanying hydrolysis. At the [BlaC] required to see any
significant rate of reaction (~2 µM), variation of the [doripenem] or [ertapenem] at
concentrations from 2–20 µM showed almost no difference in rate, suggesting their Km
values were less than 2 µM. The steady-state kinetic parameters determined for faropenem, a
structurally distinct penem, were Km = 55 ± 11 µM, and kcat = 0.65 ± 0.04 min−1 (data not
shown). This Km value is ~17 times larger and the kcat value 8 times faster than those of
meropenem (14).
Detailed investigations of the kinetics of carbapenem hydrolysis under near stoichiometric
enzyme concentrations were carried out over 30 minute time periods. As shown in Figure 1,
a steady-state kinetic burst was observed with both compounds where the magnitudes of the
burst are proportional to the concentration of BlaC used. Extrapolation of the rates of
hydrolysis to the y-axis demonstrates that the acylation is stoichiometric with the
concentration of enzyme.
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Due to the extremely feeble turnover rate, we further tested these carbapenems as inhibitors
of the reaction of nitrocefin with BlaC. Nitrocefin is an extremely good substrate for BlaC
and its β-lactam ring-opened form is extremely chromogenic. As shown in Figure 2,
doripenem and ertapenem act as slow-onset, tight binding inhibitors of BlaC when the
hydrolysis of nitrocefin was monitored. In contrast, faropenem exhibited standard,
competitive inhibition with no time dependent component (data not shown). This type of
time-dependent inhibition for a dead-end inhibitor is modeled as being due to the reversible
formation of a non-covalent complex (E–I), followed by the reversible conversion to an
isomerized complex (E–I*). However, in the case of a slow substrate for BlaC, the initially
formed Michaelis complex reacts with the enzyme in an irreversible step to generate the
BlaC-carbapenem covalent intermediate. This is then hydrolyzed slowly to regenerate the
free enzyme that can react with nitrocefin. While the same equation is used to fit the two
models, the kinetic constants that contribute to kiso (Figure S1) and Ki (or Kd) are different.
Using the fits of the slow-onset data and eq 3 to calculate Kd, k2 and k3, we can then used
eqs 4 and 5 to calculate the apparent Km and kcat values for doripenem (0.18 µM and 0.016
min−1, respectively), and for ertapenem (0.18 µM and 0.017 min−1, respectively). We have
not corrected for the concentration of nitrocefin used in these experiments because of the
large standard errors (>40%) associated with these kinetic parameters (the reported Km
values are apparent values). However, the extremely tight binding and extremely low
turnover of these carbapenems is evident from these rather imprecise kinetic data.
Mass Spectrometry
The rapid acylation and slow deacylation of BlaC by the carbapenems allows the
observation of the covalently bound, acyl-enzyme intermediate by Fourier transform ion
cyclotron resonance. A freshly prepared solution containing excess BlaC and doripenem
displayed three peaks: the first peak corresponds to free BlaC with mass/charge ratio (m/z) =
28,785.0, a second peak corresponding to the covalently acylated BlaC-doripenem complex
with mass/charge ratio (m/z) = 29, 204.1 and a third peak whose mass corresponds to the
mass of the covalently acylated BlaC-doripenem complex minus 44 mass unit (m/z = 29,
161.0), as shown in Figure 3. With ertapenem, the two covalent acylated BlaC complex
peaks observed had molecular masses of 29,260.0 and 29,217.1, corresponding to acylated
BlaC-ertapenem complex and acylated BlaC-doripenem complete minus 44 mass units,
respectively. This data demonstrates that both doripenem and ertapenem undergo the same
chemical breakdown in the active site as meropenem (14). Once the acyl-enzyme forms, the
carbapenems partition between hydrolysis and enzyme-catalyzed decomposition of the C6
hydroxyethyl substituent, via a retro-Aldol decomposition, which yields acetaldehyde (14).
Intriguingly, the intensities of the acyl-enzyme complexes remain stable over the time period
of 90 min for doripenem and ertapenem. This is in contrast with previous observations with
meropenem, where the acylated forms of the enzyme started to diminish after several
minutes. These data suggest that doripenem and ertapenem form more stable complexes
with BlaC than meropenem, reinforcing the kinetic data.
X-ray Crystallography
The 2.2 Å data from a 90 min doripenem-soaked crystal were refined to an Rwork of 0.161
and an Rfree of 0.205. The 1.3 Å diffraction data from a 10 minute ertapenem-soaked crystal
refined to an Rwork of 0.147 and an Rfree of 0.176. The 2.0 Å diffraction data from a 90 min
ertapenem-soaked crystal were refined to an Rwork of 0.175 and an Rfree of 0.222. In these
three structures, the active site Ambler residue Ser70 has been covalently linked with the
ring open form of these β-lactams in accordance with the acylation chemistry of the first half
of the enzymatic reaction (Scheme 1). The quality of the electron density is displayed in
Figure 4 and Figure 5 under a Fo-Fc omit calculated map contoured at 2.0 σ.
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The C3 atom of the pyrroline rings of doripenem and ertapenem covalent adducts are sp3
hybridized in the 90 min soaks. These results require an isomerization event occurring in the
BlaC-carbapenem adducts in which the original Δ2 pyrroline ring was tautomerized to
generate the Δ1 pyrroline ring, evidenced by the collinear positioning of the C5, N1, C2, and
C3 atoms. In addition, the BlaC-doripenem and ertapenem covalent adduct densities allow
for the positioning of the thioether sulfur atom in the unambiguous assignment of the S
configuration at C3, requiring protonation at the re face of the C2–C3 double bond. This is
similar to our earlier findings with BlaC crystals soaked with meropenem on similar time
scales (14), and represents the thermodynamically preferred product with the trans
orientation of the C4 methyl and thioether substituents. Interestingly, in the recently reported
structure of the Class D β-lactamase, OXA-1 covalent adduct with doripenem, revealed that
while an identical isomerization had taken place, that the final Δ1 pyrroline ring product was
of the opposite, R, stereochemistry (22).
In the structure determined after the shorter 10-minute soak, the BlaC-ertapenem adduct was
covalently bound in the active site, but in a different geometry. On this shorter time scale,
the BlaC-ertapenem adduct C3 atom is found in its original sp2 hybridization with the
definitive collinear positioning of the thioether sulfer atom in line with the N1, C2, C3, and
C4 bonds indicating the presence of the Δ2 pyrroline ring. This result requires that β-lactam
ring cleavage and isomerization of the methyl pyrroline ring not be concerted.
The active site interactions vary in some subtle ways between the pre and post-isomerization
ertapenem complexes, yet a number of common interactions are observed in all complexes.
Both the BlaC-ertapenem and -doripenem adducts bind as covalent adducts with the active
site Ser70 and position their lactam ring-opened ester carbonyl oxygen atom within the
oxyanion hole formed from hydrogen bonding interactions with the Ser70 and Thr253 amide
nitrogen atoms. All structures contain a hydrophobic interaction between the methyl group
of the pyrroline ring and the sidechain of Ile117 and different forms of hydrogen bonding
interactions between the C6 hydroxyethyl substituent of the carbapenem and Glu166. All
three structures also show a conserved interaction between the sidechain hydroxyl of
Ser130, which consistently hydrogen bonds the pyrroline ring nitrogen atom at a distance of
2.7–2.8 Å. The pyrroline C2 carboxylate group forms hydrogen bonds with the Thr251
hydroxyethyl side chain and an active site water molecule. In structures of other β-
lactamase-carbapenem adducts, this carboxylate electrostatically interacts with a conserved
arginine residue (R244 in TEM-1) (23), but this is not the case for BlaC. In the pre-
isomerized ertapenem structure, there is an additional hydrogen bond between the C2
carboxylate of the pyrroline ring and Thr253, which is broken upon the repositioning of the
meta-amino-benzoate ‘arm’ observed in the post-isomerized ertapenem structure. The
isomerization and stereospecific protonation leads to a reorientation of the terminal portion
of the molecule within the active site, allowing for the formation of the hydrogen bond
between the terminal carboxylate group of the meta-amino-benzoic acid moiety and Ser118.
A final difference between the initially formed Δ2-pyrroline isomer and the final Δ1-
pyrroline form is the orientation of the C6 hydroxyethyl substituent. In the pre-isomerized
complex it is oriented away from Lys73 and hydrogen bonds to the carboxyl O1 of Glu166
as well as the Asn186 nitrogen, but rotates upon isomerization to hydrogen bond with the
carboxyl O2 of Glu166 and the ε-amino group of Lys73 in a manner similar to that observed
for the doripenem complex.
The structures of the pre and post-isomerization states reveal the mechanistic basis for the
relative stability of the carbapenems within the active site of BlaC and their ability to resist
hydrolysis by the enzyme. As seen in Scheme1, deacylation-hydrolysis from the enzyme
requires the activation of the conserved active site water by the side chain carboxyl O2 of
Glu166. The probability of water activation by Glu166 decreases with the increased distance
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between the two. The distance between the carboxyl O2 of Glu166 and the active site water
is significantly increased after isomerization from 2.4 Å to 2.7 Å, making water activation
less probable. The isomerization event does not directly cause these changes but rather alters
the positioning of the adduct such that the adduct-hydroxyethyl flips in the active site,
breaking the hydrogen bond formed between the hydroxyl group with the side chain amide
nitrogen of Asn186 and the adjacent carboxyl O1 of Glu166 oxygen. The hydroxyethyl
substituent then rotates to generate a hydrogen bond network with Lys73 and the carboxyl
O2 of Glu166, effectively ‘pulling’ this essential base away from the conserved active site
water. These residues (Lys 73 and Glu166) are involved as general bases in the acylation
and deacylation reactions, respectively. In addition, by hydrogen bonding carboxyl O2 of
Glu166, the reoriented hydroxyethyl substituent reduced the basicity of Glu166. These two
factors, introduced in the reorientation of the hydroxyethyl substituent in the post-
isomerization complex, reduce water activation and thereby stabilize the acyl-intermediate
in the active site. The studies reported here allow us to directly visualize the changes that
occur between the pre- and post-isomerization adduct structures and are atomic level
observations relevant to the biphasic kinetics previously reported for the reactions between
carbapenems and the RTEM β-lactamase (24).
The Δ2 to Δ1-pyrroline isomerized forms of carbapenems have been known to form within
the active sites of various β-lactamase enzymes (25). In confirmation of this, crystal
structures of carbapenems bound within the active sites of the Class A β-lactamases TEM-1
(PDB entry 1BT5) (26) and SHV-1 (PDB entry 2ZD8) (27) as well as AmpC (PDB entry
1LL5) (28) a Class C β-lactamase all revealed the Δ2 form of the carbapenem bound in the
active sites, while the Class D OXA-1 (PDB entry 3ISG) (22) and the Class A BlaC (PDB
entry 3DWZ) (14) were both bound with carbapenems in the Δ1 isomerized forms with
respective R and S-stereochemistries. Our findings are the first to show the structures of
both the Δ2 and Δ1 forms of a carbapenem bound to a single β-lactamase. Interestingly,
several of the structures of carbapenems bound as the Δ2 isomers show evidence for
alternate conformations for the carbapenem-carbonyl oxygen position. This oxygen is found
buried within the oxyanion-hole as well as bound in a position rotated by 180 degrees,
usually facing an opposing serine residue (Ser130). In these instances it has been proposed
that the flipping of the carbonyl oxygen from the oxyanion-hole blocks formation of the
deacylation tetrahedral intermediate to inhibit the enzyme. In the cases of OXA-1 and BlaC,
the carbapenem-carbonyl oxygen is only found bound tightly within the oxyanion-hole and
no evidence of alternate conformers has been observed. In these cases inhibition by the
carbapenem is likely due to disruption of water activation.
A second possible reason for the observed alternate conformers at the carbapenem-carbonyl
is likely due to the position of the carbapenem-carboxylate moiety within those active sites.
To date, those β-lactamases with alternate conformations for the carbapenem-carbonyl, have
a highly conserved Arg244 reside which electrostatically interacts with the carbapenem-
carboxylate moiety. The OXA-1 and BlaC active sites lack this arginine interaction and
instead use a combination of threonine and/or serine residues coordinated with waters to
bind the carboxylate moiety. These residues are located closer to the oxyanion hole and act
to ‘clamp’ the carboxylate into a proximal position, as opposed to the Arg244 mechanism of
carboxylate binding, where distance introduces flexibility, allowing for the alternate
positioning of the pyrroline ring. This bonding pattern to the carbapenem allows for
alternate “in/out” conformations of the carbapenem carbonyl in the oxyanion hole. In
contrast, the carbapenem carbonyl is tightly bound in the oxyanion hole in BlaC in both the
Δ2 and Δ1 forms reported here.
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Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
This work was supported in part by a grant from the National Institute of Health (AI33696 to J. S. B.) and in part by
the Charles Revson Foundation (to L.W.T.)
ABBREVIATIONS
BlaC
Mycobacterium tuberculosis beta-lactamase
TB
Tuberculosis
XDR-TB
extensively drug resistant
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Figure 1.
Time courses of doripenem (A) and ertapenem (B) hydrolysis with various concentrations of
BlaC.
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Figure 2.
Time courses of nitrocefin hydrolysis by BlaC in the presence of doripenem (upper) and
ertapenem (lower).
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Figure 3.
Mass spectra of enzyme-carbapenem species. The 25+ charge state ions are shown.
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Figure 4.
(A) Overall structure of BlaC displayed in rainbow from N term (blue) to the C term (red),
with the doripenem adduct displayed in red surface mesh. (B) Fo-Fc omit density (green)
contoured at 2.0 σ surrounds the covalent doripenem adduct formed at the Ambler active-
site residue serine 70. All structure figures were produced using Pymol.
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Figure 5.
(A) Fo-Fc omit density (green) contoured at 2.0 σ surrounds the covalent ertapenem adduct
formed at the Ambler active-site residue serine 70 in the pre-isomerization state. (B) Fo-Fc
omit density (green) contoured at 2.0 σ surrounds the covalent ertapenem adduct formed at
the Ambler active-site residue serine 70 in the post-isomerization state. The resolution of the
densities unambiguously demonstrates the shift in stereochemistry with the change from sp2
to sp3 hybridization of the C3 carbapenem carbon atom with the change in the position of
the density associated with the meta-amino-benzoate and the hydoxyethyl ertapenem
moieties.
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Scheme 1.
(A) the structures of doripenem and ertapenem. (B) The chemical mechanism of hydrolysis
of ertapenem by the Mycobacterium tuberculosis BlaC.
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Table 1
Data Collection and Refinement Statistics
Data Collection
Doripenem
Δ1-isomerζ
Ertapenem
Δ2-isomer
Ertapenem
Δ1-isomer
Resolution (Å)
50.0-2.2
(2.32-2.20)
50.0-1.30
(1.33-1.30)
50.0-2.0
(2.07-2.00)
Completeness
100% (100%)
100.0% (100%)
99.5 (99.9)
Redundancy
7.6 (7.4)
7.5 (5.7)
4.4 (4.4)
I/sigma(I)
3.8 (1.6)
21.4 (1.8)
9.8 (4.0)
Rmerge
0.077 (0.47)
0.057 (0.757)
0.158 (0.373)
Space Group
P212121
P212121
P212121
Unit cell (Å)
a =49.989
b =68.068
c =75.792
α = β = γ = 90.0°
a = 49.66
b = 67.92
c = 75.55
α = β = γ = 90.0°
a =49.934
b =67.830
c =75.201
α = β = γ = 90.0°
Reflections
13,695 (1,943)
60,263 (4,388)
17,920 (1,762)
Refinement Statistics
Rwork
0.161 (0.176)
0.147 (0.265)
0.175 (0.191)
Rfree
0.205 (0.237)
0.176 (0.278)
0.222 (0.281)
Average B-factors (Å2)
Protein
6.97
10.49
6.09
Adduct
27.32
18.64
15.50
Solvent
17.51
32.64
14.36
PO4
12.89
14.40
10.53
RMS deviations
bonds (Å)
0.010
0.010
0.012
angles (°)
1.204
1.428
1.386
Ramachandra
Favored= 97.7%
outliers= 0.0%
Favored= 97.7%
outliers= 0.0%
Favored= 98.1%
outliers= 0.0%
PDB accession code
3IQA
3M6B
3M6H
Values in parentheses are for the highest resolution bin.
ζThis data processed using Mosflm
Biochemistry. Author manuscript; available in PMC 2011 May 4.
|
3M6K
|
Crystal Structure of N-terminal 44 kDa fragment of topoisomerase V in the presence of guanidium hydrochloride
|
Structures of minimal catalytic fragments of topoisomerase V
reveals conformational changes relevant for DNA binding
Rakhi Rajan*, Bhupesh Taneja*,†, and Alfonso Mondragón*,‡
* Department of Biochemistry, Molecular Biology and Cell Biology, Northwestern University, 2205
Tech Dr, Evanston, IL 60208
Summary
Topoisomerase V is an archaeal type I topoisomerase that is unique among topoisomerases due to
presence of both topoisomerase and DNA repair activities in the same protein. It is organized as an
N-terminal topoisomerase domain followed by 24 tandem helix hairpin helix (HhH) motifs.
Structural studies have shown that the active site is buried by the (HhH) motifs. Here we show that
the N-terminal domain can relax DNA in the absence of any HhH motifs and that the HhH motifs
are required for stable protein-DNA complex formation. Crystal structures of various
topoisomerase V fragments show changes in the relative orientation of the domains mediated by a
long bent linker helix, and these movements are essential for the DNA to enter the active site.
Phosphate ions bound to the protein near the active site helped model DNA in the topoisomerase
domain and shows how topoisomerase V may interact with DNA.
Introduction
DNA topoisomerases are enzymes found in all forms of life (bacteria, eukarya, and archaea)
and they regulate the topological state of DNA inside the cell. They form a transient break in
a single or double stranded DNA and allow the passage of another single or double DNA
strand through the break, before resealing the break (Champoux, 2001) (Schoeffler and
Berger, 2008). As a result of this, topoisomerases can relax supercoiled DNA, help in the
segregation of DNA strands following replication, and lead to the formation and resolution
of knots and catenates (Gellert, 1981). Topoisomerases participate in many aspects of DNA
metabolism, such as replication, recombination, and transcription (Champoux, 2001). In
addition, they are targets of various anti-cancerous drugs and anti-bacterial agents (Maxwell,
1999; Pommier, 1998; Rothenberg, 1997; Wang et al., 1997).
DNA topoisomerases are broadly classified into two types, type I and type II enzymes. Type
I enzymes cleave a single strand of a DNA molecule and pass another single or double
stranded DNA through the break before resealing the opening. Type II enzymes cleave both
‡Corresponding author: Phone: 847-491-7726, Fax: 847-467-6489, a-mondragon@northwestern.edu.
†Present address: Institute of Genomics and Integrative Biology, CSIR, Delhi, India
Protein data bank accession codes
The final structure factors and coordinates of Topo-31, Topo-44 Form I, Form II, and Form III have been deposited in the Protein Data
Bank with accession codes 3M7G, 3M7D, 3M6K, and 3M6Z respectively.
Supplementary data
Supplementary data are available at Structure Journal Online.
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Author Manuscript
Structure. Author manuscript; available in PMC 2011 July 14.
Published in final edited form as:
Structure. 2010 July 14; 18(7): 829–838. doi:10.1016/j.str.2010.03.006.
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strands of a double stranded DNA in concert and pass another double stranded DNA through
the break. Type I enzymes use the torsional energy stored in the supercoiled DNA to drive
DNA relaxation and hence they do not require high energy cofactors, such as ATP, for their
activity (Baker et al., 2009) Type II enzymes, on the other hand, require ATP and Mg2+ for
their activity. Type I topoisomerases are further subdivided into three subtypes: IA, IB, and
IC (Forterre et al., 2007). Type IA and IB enzymes have been studied extensively (Baker et
al., 2009) and there is ample information available about their general mechanism of DNA
relaxation and the mode of DNA binding. Type IC, on the other hand, is a relatively new
subtype. Currently topoisomerase V is the only member of this family and it has been
identified only in the Methanopyrus genus. Previously, topoisomerase V had been
considered as a type IB enzyme based on its biochemical characteristics (Slesarev et al.,
1993), but the crystal structure of an N-terminal 61 kDa of topoisomerase V (Topo-61)
(Taneja et al., 2006) revealed a completely new fold without similarity to other
topoisomerases or any other known protein. Furthermore, the orientation of the putative
active site residues is also different from other type I topoisomerases, suggesting a different
mechanism of cleavage and religation of DNA. These observations, together with the lack of
sequence similarity, indicated that topoisomerase V defines a new subtype of type I enzymes
(Forterre, 2006).
Topoisomerase V was identified in Methanopyrus kandleri, an extremophile isolated from a
deep-water ‘black smoker’ chimney in the Gulf of California (Huber et al., 1989). The
enzyme is active at very high temperatures (122°C) and high salt concentrations (0.65 M
NaCl and 3.1 M potassium glutamate). The unusual characteristic of topoisomerase V is that
it has both topoisomerase and DNA repair activities in the same polypeptide (Belova et al.,
2001). Based on the sequence analysis of topoisomerase V, it has been predicted that the
protein contains 24 helix-hairpin-helix (HhH) DNA binding motifs arranged as 12 (HhH)2
domains around the N-terminal topoisomerase domain (Belova et al., 2002) (Figure 1A).
Some of these (HhH)2 domains are involved in the apurinic/apyrimidinic (AP) site-
processing activity, but the exact location of the repair active site is not known yet.
Topoisomerase V can relax both positively and negatively supercoiled DNA without the
need for metal cations or high energy cofactors. Single molecule experiments have shown
that topoisomerase V relaxes DNA by a constrained swiveling mechanism, relaxing around
12 turns of DNA per relaxation cycle (Taneja et al., 2007). Type IB enzymes, which also use
a constrained swiveling mechanism for DNA relaxation, relax around 19 turns of DNA per
relaxation cycle (Koster et al., 2005).
The structure of Topo-61 showed that the topoisomerase domain is mainly alpha helical and
that the first four (HhH)2 domains curl around the topoisomerase domain (Taneja et al.,
2006) (Figure 1B). The topoisomerase and (HhH)2 domains are joined by a long bent helix,
termed the “linker helix”. Three of the five putative active site residues are present in a
helix-turn-helix (HTH) domain and the other two are present in an intervening loop and a
helix. The active site residues are buried by the first (HhH)2 domain and it has been
suggested that large conformational changes will be needed for the DNA to access the active
site of topoisomerase V (Taneja et al., 2007). Here we present data that shows that the N-
terminal 31 kDa fragment of topoisomerase V (Topo-31) has topoisomerase activity,
consistent with previous predictions based on the structure. In addition, we show that the
Topo-44 fragment (N-terminal 44 kDa fragment of topoisomerase V) can form a stable
protein-DNA complex, emphasizing the need of the (HhH)2 domains for binding DNA. We
determined a crystal structure of (Topo-31) fragment, which has only the topoisomerase
domain, and three different crystal structures of the Topo-44 fragment, which includes the
topoisomerase domain and three tandem HhH motifs. In all structures, the topoisomerase
domain is very similar. In contrast, the structures of Topo-44 show conformational changes
in the linker helix resulting in variable orientations of the (HhH)2 domains when compared
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to the Topo-61 structure. Phosphate ions are present in the vicinity of the topoisomerase
active site in two of the Topo-44 structures. Some of the catalytic residues interact with the
phosphate ions and may mimic contacts with DNA. These observations suggest that the
movement of the (HhH)2 domains is mediated by the linker helix and helps expose the
topoisomerase active site to facilitate DNA binding. In addition, the location of the
phosphate ions suggests a possible path for the DNA and the way the active site residues
interact with it.
Results
The topoisomerase domain can relax DNA
DNA relaxation assays using different topoisomerase V fragments showed that the
topoisomerase domain alone is capable of relaxing DNA. Topoisomerase V fragments with
different numbers of (HhH)2 domains, Topo-31, Topo-44, and Topo-78, were studied using
relaxation assays. Topo-31 has no (HhH)2 domains, Topo-44 has one full and one partial
(HhH)2 domain, while Topo-78 has eight full (HhH)2 domains, including a putative DNA
repair domain. In addition to standard conditions, the effect of different pH conditions and
presence of magnesium ions were also tested. The experiments show that Topo-31 is
capable of relaxing DNA, despite the absence of the (HhH)2 domains (Figure 2B). A pH
profile analysis for the DNA relaxation assays showed that Topo-78 relaxes DNA over a
wider pH range (pH 5 to 9), while Topo-31 and Topo-44 relax DNA optimally at pH 5
(Figure 2A, 2B, 2C). In addition, magnesium is not required for the reaction, but stimulates
it at all pH values (Figure 2B, 2C). Topo-78 can relax DNA to the same extent with lower
amounts of protein (0.1 μg/reaction) compared to Topo-44 (~1.5 μg/reaction) and Topo-31
(~9 μg/reaction). This could be due to the enhanced DNA binding facilitated by the (HhH)2
domains. Together, these results suggest that, even though the (HhH)2 domains are
dispensable for topoisomerase activity, they enhance DNA relaxation activity. In addition,
the pH dependence of the DNA relaxation activity indicates that the reaction is likely to
involve side chains with ionizable groups in the low pH range, such as glutamates. Finally,
the magnesium independence of the reactions confirms that even the smallest fragments do
not require metals for activity, although magnesium has a stimulatory effect. This may be
due to favorable interactions of the cations with DNA.
The (HhH)2 domains enhance DNA binding affinity
EMSA experiments with different fragments of topoisomerase V and DNA showed that
(HhH)2 domains could help in the formation of a stable protein-DNA complex. Various
topoisomerase V fragments (Topo-31, Topo-44, and Topo-78) and single and double
stranded DNA were analyzed by EMSA experiments. Topo-44 and Topo-78 formed stable
complexes with a 39mer double stranded DNA (Figure 2D), while no DNA binding was
observed for the Topo-31 fragment (data not shown). These observations indicate that
(HhH)2 domains are necessary for a stable protein-DNA complex and that as few as one and
half (HhH)2 domains are enough for formation of a stable protein-DNA complex. EMSA
with single stranded DNA showed that Topo-31 and Topo-44 cannot bind to single stranded
DNA, while Topo-78 can bind to single stranded DNA (data not shown).
Overall Structures
The topoisomerase domain of topoisomerase V is a helical-rich compact domain that has no
structural similarity to any other known protein. The only recognizable structural element is
a HTH that contains some of the active site residues. Not surprisingly, the topoisomerase
domain of the four structures (Topo-31, Topo-44 (Forms I, II, and III)) superimpose very
well on each other and also to that from the Topo-61 structure. In the Topo-31 structure, two
surface loops, residues 39-49 and 120-124, adopt a different conformation compared to the
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Topo-61 and Topo-44 structures. These two loops are not always visible in the Topo-44
structures, suggesting that they are mobile regions. The r.m.s.d. for the superposition of the
topoisomerase core domain of all the new structures on to the Topo-61 structure range from
0.2 Å to 0.7 Å if the two mobile surface loops are not included (Figure 3A). In general, the
topoisomerase domain remains unchanged and is identical in all structures. The (HhH)2
domains also remain largely unchanged, with r.m.s.d. for the superposition of only the
(HhH)2 domains from the three Topo-44 crystal forms and equivalent domains in the
Topo-61 structure ranging from 0.31 Å to 0.56 Å.
The five crystallographically independent structures of Topo-44 (Form I, Form II A and B
monomers, and Form III A and B monomers) were compared with each other and to the two
crystallographically independent Topo-61 monomers to understand the conformational
changes in the protein. The r.m.s.d. for the superposition of all the Topo-44 structures
(residues 3-375) on to the Topo-61 fragment or on each other vary between 0.9 Å and 2.7 Å,
with the majority above 1.5 Å, showing that in general the structures have slightly different
conformations. As mentioned above, the different domains behave as rigid or almost rigid
subunits and the only change in the structure is the relative orientation between the
topoisomerase and the (HhH)2 domains. The change in orientation of the domains starts at
the linker helix (residues 269-295), which acts as a hinge region, and follows into the
(HhH)2 domains. At the start of the linker helix, the structures superimpose very well for all
five Topo-44 and two Topo-61 structures. In the middle of the linker helix there is a kink
after which the linker helix from all the structures shows different orientations (Figure 3B).
The flexibility of the linker helix is also evident by the fact that the linker helix in the B
subunit of Form III crystals appears in two alternate conformations. The change in the
relative orientation of the (HhH)2 and topoisomerase domains (Figure 3C and 3D), suggests
that these domains can adopt different orientations and these movements might be necessary
for the DNA to access the active site.
The topoisomerase domain has a positively charged groove adjacent to the active site
The structure of the Topo-31 as well as the structures of the Topo-44 fragment reveals the
presence of a positively charged groove in the protein that encompasses the active site
region (shown later in Figure 6C). This charged groove had been observed before in the
structure of the Topo-61 fragment, although several (HhH)2 motifs partially obstruct it
(Taneja et al., 2006). The structure of the Topo-31 confirms the presence of the groove even
in the absence of the (HhH)2 motifs. The groove is long and can be deep in some areas. It
includes regions of the HTH motifs and extends all the way to the linker helix. All the
residues forming the active site pentad point towards the groove. The active site tyrosine,
Tyr226, is found near one of the ends of the groove, a region where it widens. The positively
charged character of the groove and its presence by the active site strongly suggest that it
may be involved in DNA binding.
Phosphate ions bind in the groove near the topoisomerase active site
An interesting observation stemming from the Form II and Form III Topo-44 structures is
the presence of phosphate ions near the positively charged DNA binding groove. All three
Topo-44 crystal forms were crystallized in the presence of phosphate-citrate buffer, but only
Form II and Form III structures showed phosphate ions bound to the protein, which were
assigned based on electron density consistent with a tetrahedral phosphate ion (Figure 4A).
Form II and Form III crystals include 1–1.2 M guanidium hydrochloride in the
crystallization solution. The high resolution Form III structure shows clear density for three
guanidium ions bound to the protein, two very well ordered and one with weak density. The
presence of guanidium hydrochloride in the crystals appears to trigger a conformational
change allowing the binding of phosphate ions to the protein. It is interesting to note that
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Form I crystals did not show any bound phosphate albeit its presence in the crystallization
condition. This could be due to the absence of guanidium hydrochloride to trigger the
binding of phosphate ions as observed in Form II and Form III structures. There are three
phosphate ions in the B subunit and none in the A subunit of the Form II Topo-44 structure.
Two of the phosphates are in the topoisomerase active site and one of them forms close
contacts with the putative active site residues in the topoisomerase domain (Figure 4B).
Form III crystal has seven phosphate ions, three in each subunit and one between both the
subunits. In the Form III structure, the phosphate ion near the active site Tyr226 is absent,
but it shows several new locations for phosphate ions, especially in the positively charged
groove containing the topoisomerase active site (Figure 5A). An overlay of the A and B
subunits of the Topo-44 Form III structure with the B subunit of Topo-44 Form II structure
shows eight unique phosphate ions (Figure 5A). It clearly shows that there are more
phosphate ions bound in the positively charged groove compared to other regions of the
protein.
Taking into account all structures, there are five unique phosphate ion binding sites in the
putative DNA binding groove and an additional one near its end and close to the start of the
linker helix. Several pairs of phosphates in the groove are separated by a distance of around
7 Å (Figure 5B), which would be consistent with the phosphate-phosphate distance in
adjacent nucleotides in a DNA double helix. One of the phosphates (P1) is found near the
active site tyrosine and is coordinated by Tyr226 and Arg131, two residues that have been
implicated in cleavage and religation of the DNA (Taneja et al., 2006), and by Glu215,
whose charge may be important for interactions with DNA (R.R. and A.M., unpublished
observations). The side chains of the tyrosine and the glutamate residues are in contact with
Arg144 and His200, the other putative active site residues, and these interactions may help
to orient them for the catalytic reaction. Adjacent to P1, there is a second phosphate (P2) at a
distance of 7.5 Å which is trapped between the topoisomerase domain and an HhH motif. P2
is coordinated by Arg131, an active site residue, in addition to Arg108 from the
topoisomerase domain and Arg293 and serines 322 and 324 from the second HhH motif
(Figure 6C). Three more phosphates are found in the groove (P3, P4, and P5) coordinated
mainly by positively charged residues, such as Arg37, Lys47, Arg108, Lys134, and Arg135
from the topoisomerase domain and also residues from the linker helix such as Tyr289 and
Arg293. In general, some of the side chains can contact more than one phosphate. The
distance between P3 and P4 and P4 and P5 is 6.8 Å and 6.5 Å respectively. A final
phosphate (P6) is located at the start of the linker helix and on the edge of the groove
(Figure 5A).
Discussion
Topoisomerase V is active at very high temperatures (122°C) and high salt concentrations.
DNA relaxation assays with various topoisomerase V fragments (Topo-44 and Topo-61)
show that a temperature above 60° C is required for optimal activity, although longer
fragments of topoisomerase V can relax DNA at lower temperatures (Taneja et al., 2007).
Topo-44 was first identified by limited proteolytic digestion of the full length topoisomerase
V protein (Belova et al., 2002) at 80°C. In contrast, Topo-61 is the shortest fragment
showing topoisomerase activity when the proteolytic reaction is performed at 37°C (Belova
et al., 2002). The N-terminal Topo-31 fragment, which contains neither HhH motifs nor the
linker helix, was identified as the smallest region spanning the topoisomerase domain from
the crystal structure of Topo-61 fragment (Taneja et al., 2006) and it was suggested that it
could represent the minimal domain capable of relaxing DNA. Relaxation experiments with
this minimal domain show that this is indeed the case, although the activity is not as robust
as with longer fragments. As expected, Topo-31 does not require magnesium for activity,
but magnesium enhances it, as is the case for type IB topoisomerases, which also uses a
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swiveling mechanism for DNA relaxation (Stewart et al., 1996). The optimal pH for activity
for the Topo-31 and the Topo-44 fragments is around 5. This pH dependence is not observed
for the Topo-78 fragment. The DNA relaxation by shorter fragments of topoisomerase V at
pH 5 could point to the involvement of some ionizable side chains in the relaxation activity.
It could also be simply due to the effects of various side chains on DNA binding. Further
experiments with different active site mutations in both longer and shorter fragments of
topoisomerase V will be required to probe the pH dependence of the relaxation reaction by
shorter topoisomerase V fragments.
Gel shift experiments show that Topo-44 and also longer fragments (Topo-78) can bind
double stranded DNA. Surprisingly, Topo-31 does not show DNA binding activity in these
assays even though it is still capable of relaxing DNA. It appears that the presence of the
(HhH)2 domains stabilizes the DNA/protein complex. One possibility is that the (HhH)2
domains could play a similar role to the cap domain present in type IB enzymes, which helps
to encircle the DNA during the swiveling reaction (Redinbo et al., 1998). In addition, both
short fragments of topoisomerase V do not bind single stranded DNA, whereas Topo-78 can
form a stable complex with single stranded DNA (data not shown). (HhH)2 domains binding
to single stranded DNA has been observed before. For instance, the N-terminal 8 kDa of
mammalian polymerase β, which contains a single HhH motif, binds to single stranded DNA
through both helices (Kumar et al., 1990; Liu et al., 1994). The exact mode of single
stranded DNA binding by Topo-78 or the possible role in relaxation or repair activities is not
yet clear.
The structure of Topo-61 showed that the topoisomerase active site of topoisomerase V is
buried by one of the (HhH)2 domains suggesting that conformational changes are essential
for the protein to bind DNA. The present structures of Topo-44 reinforce this observation
and show that the (HhH)2 domains can change their position relative to the topoisomerase
domain and that this change is mediated by the movement of the linker helix. The (HhH)2
domains act as rigid individual units, as evidenced by the fact that in different structures
they show the same structure and relative orientation of the two HhH motifs. The
topoisomerase domain also appears to be rigid showing the same structure even in the total
absence of the rest of the protein. The linker helix (residues 269-295), which is a long bent
helix, serves as a hinge for the movement of the (HhH)2 domains away from the rigid
topoisomerase domain, possibly by responding to interactions with double stranded DNA.
This movement has to be quite large. The Topo-44 structures in the absence of DNA capture
the regions that move, but do not show the full extent of the movement or indicate the way
the HhH motifs interact with DNA.
As mentioned before, topoisomerase V binds double stranded DNA and has a groove wide
enough to accommodate double strand DNA (Figure 6C). The presence of an HTH domain
normally associated with DNA binding, the positively charged nature of it, and several
phosphates bound along it suggest that this groove could be involved in DNA binding. In
addition, the active site is found in this groove and some residues form part of the HTH
domain. Previously, DNA was modeled bound to the topoisomerase domain (Taneja et al.,
2006) based on the structures of HTH domains in complex with DNA but there was no
evidence to support it. Using the phosphates present in the groove in the current structures, it
is possible to refine this model. A superposition of the B subunit of Form II and the A and B
subunits of Form III Topo-44 structures shows five different phosphate ions in the positively
charged groove which are separated by a distance of around 7 Å, consistent with the
distance of consecutive phosphates in B DNA of ~6.4 Å. A sixth phosphate ion is found
outside the groove near the linker helix. A double stranded DNA molecule was modeled into
the groove based on the positions of the phosphate ions (Figure 6). Only five out of the six
phosphates could be placed on the DNA molecule, as one of them was inconsistent with a
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double stranded molecule. Phosphate ions P3, P4, and P5 would correspond to three
adjacent phosphates in one DNA strand, while P1, located near the active site, would belong
to the opposite strand. A final phosphate (P6) is away from the groove and near the linker
helix (Figure 6A, 6B). The fit to the four inner phosphates is excellent and the DNA can be
accommodated in the groove of the Topo-31 structure without the need for any major
rearrangements of the protein backbone. The fifth phosphate (P6) does not fit as well and a
better fit would require movement of either the protein or the DNA, but the change would be
relatively modest. Several side chains would need to move, but these changes would also be
minor. The major change needed to accommodate the DNA in the structures with the
(HhH)2 domains present is the movement of the (HhH)2 domains away from the
topoisomerase domain (Figure 6B). The movement of (HhH)2 domains should be feasible as
is evident from the Topo-44 structures showing different orientations of the (HhH)2
domains. The location of the (HhH)2 domains after DNA binding is not evident, but one
possibility is that they would help enclose the DNA to form a clamp around it, similar to the
arrangement in type IB enzymes.
In the model of the topoisomerase domain in complex with DNA, the active site residues are
in close contact with the backbone of DNA. The catalytic Tyr226 is pointing towards the
phosphate of the DNA backbone; Arg131 and Arg144 are positioned to stabilize the protein-
DNA covalent complex. Surprisingly Glu215 also appears to interact directly with the DNA
phosphate backbone. The other active site residues like His200 and Lys 218 are also near the
DNA. The active site is located near the end of the groove, where it widens. At this end, the
DNA fits loosely in the groove, which is spacious to accommodate the movement of the
strands. The ‘constrained swiveling’ mechanism employed by type IB and IC enzymes
necessitates rotation of one strand about the other after forming the covalent protein-DNA
intermediate. The position of the active site at the wider end of the putative DNA binding
groove would facilitate the rotation of the DNA strand at this end, while holding the rest of
the DNA in place through extensive interactions along the groove.
Even though type IB and IC enzymes have a similar overall mechanism of action, the
structures of fragments of topoisomerase V suggest many differences. Type IB enzymes
have two domains which come together to form a C-shaped clamp around the DNA (Perry et
al., 2006; Redinbo et al., 1998; Stewart et al., 1998) The protein has an open stage where
these domains are separate and this helps in the entry and release of the DNA from the
protein active site. A wide DNA binding cavity is not observed in the topoisomerase V
structures. Instead, the structures show a positively charged groove which is always present
in the protein and does not require domain rearrangements to form. DNA can access this
groove after a conformational change involving the movement of the (HhH)2 domains
exposing the active site. The (HhH)2 domains could help enclose DNA during the swiveling
of the DNA, forming a similar enclosure to the one observed for type IB enzymes. It is not
known whether all HhH motifs contact DNA simultaneously, but this appears unlikely
without a major rearrangement of the motifs. It is likely that only some of the HhH motifs
contact DNA at any given time or that some of the motifs do not have the capacity to bind
DNA. Finally, similar to type IB enzymes (Cheng and Shuman, 1998), the putative domain
enclosing the DNA is dispensable for activity, although it enhances the relaxation activity
markedly. Thus, it is likely that type IB and IC enzymes have several overall similarities in
the way that they interact with DNA, but the atomic details are markedly different.
There are still many details of the atomic mechanism of type IC topoisomerases that need to
be understood. The present functional and structural studies provide new information about
topoisomerase V including the observations that the Topo-31 is the minimal fragment
capable of DNA relaxation, the (HhH)2 domains enhance binding of the protein to DNA, the
changes in relative orientation of the domains is mediated by the linker helix, and several
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phosphate ions bind in a positively charged groove. Furthermore, the position of the
phosphate ions in the groove helped in the placement of DNA in the topoisomerase domain
and this provides an initial model of how topoisomerase V interacts with DNA. Thus the
present study helps to establish the role of different domains more clearly, to illustrate a
mechanism to drive the conformational changes needed for activity, and to suggest a
possible manner of binding DNA. Additional work on structures of protein/DNA complexes
and intermediates in the swiveling reaction are needed to understand the way this new type
of topoisomerases interacts with DNA to perform a complex reaction.
Experimental Procedures
Protein purification
The N-terminal 31 kDa (Topo-31: residues 1-269), and 44 kDa (Topo-44: residues 1 to 380)
fragments of topoisomerase V protein were cloned into the pET15b plasmid and
transformed into Escherichia coli BL21 Rosetta (DE3) cells. The N-terminal 78 kDa
(Topo-78: residues 1 to 685) fragment of topoisomerase V protein was cloned into the
pET14b plasmid (Belova et al., 2002) and transformed into Escherichia coli BL21(DE3)
cells. For protein production, cells were grown at 37° C in LB medium containing 100 μg/ml
ampicillin and 100 μg/ml chloramphenicol for Rosetta cells and LB medium with 100 μg/ml
ampicillin for BL21(DE3) cells to an optical density (OD600) of 0.6. The cells were then
cooled down on ice, followed by the addition of isopropyl β-D-1-thiogalactopyranoside
(IPTG) to a final 0.5 mM concentration, and grown overnight at 16° C. Cells were harvested
and resuspended in 50 mM Tris pH 8, 500 mM NaCl, 0.5 mM EDTA, 1 mM DTT, flash
frozen in liquid nitrogen and stored at −80° C. After thawing the pellet, pepstatin (1μg/ml),
benzamidine (1mM), PMSF (1mM), and Brij 58 (0.1%) were added to the cells and the
protein was purified as described earlier (Taneja et al., 2006) The protein was further
purified by anion exchange and gel filtration chromatography. Pure protein was
concentrated and stored in 50 mM Tris pH 8, 250 mM NaCl, and 1 mM DTT. The seleno-
methionine substituted Topo-44 was prepared from cells grown in a minimal medium
supplemented with nutrients and salts (Doublie, 1997); protein purification followed the
same procedure as for the native protein except that 5mM DTT was used in all the
purification steps and for storage.
Relaxation assays
Relaxation assays with the different topoisomerase V fragments were carried out at pH
values ranging from 4 to 10. The pH of the buffers was adjusted at 65 °C to account for the
change in pH at higher temperature. The different buffers used were: sodium acetate for pH
4 and 5, MES for pH 6, HEPES for pH 7, TRIS for pH 8, CHES for pH 9, and CAPS for pH
10. Topoisomerase activity assays were performed by incubating varying amounts of protein
(Topo-31, Topo-44 or Topo-78) with 0.2 μg negatively supercoiled pUC19 DNA in 50 mM
of the required buffer, 30 mM NaCl, 0.2 mM or 5 mM EDTA or 1 mM MgCl2. The
reactions were carried out at 65 °C for 15 min and terminated by cooling and addition of
SDS to a final 1% concentration. The products were resolved on a 1% agarose gel and
visualized by ethidium bromide staining.
Electrophoretic Mobility Shift Assay
For Electrophoretic Mobility Shift Assay (EMSA), 4 μM of a 39mer double stranded DNA
oligonucleotide (5′ GCGACGCGAGGCTGGATGGCCTTCCCCATTATGATTCTT3′) was
incubated with different concentrations of topoisomerase V fragments in 50 mM sodium
acetate pH 5, 30 mM NaCl, 1 mM MgCl2 at 65 °C for 30 minutes. Glycerol was added to
the reaction mixture to a final concentration of 8% and the products were separated on a 4 %
acrylamide native gel. The gel was stained with ethidium bromide to detect the DNA. When
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a stable protein-DNA complex was formed, there was an upward shift in the band indicating
a higher molecular weight complex.
Crystallization
Topo-31 crystals were grown using the sitting drop vapor diffusion method equilibrated
against, 23% PEG 6000, 0.1 M Na citrate pH 5.5, at 22°C. For data collection, the Topo-31
crystals were cryo-protected by adding glycerol to the mother liquor to a final 20%
concentration. Topo-44 was crystallized by the hanging drop vapor diffusion method under
three different crystallization conditions (Forms I, II, and III). Crystal Form I grew under 0.1
M phosphate citrate pH 5, 0.2 M NaCl, 15% PEG 3350 and 8% dioxane. The crystals were
cryo-protected by increasing the PEG concentration to 30%. Form II crystals grew under 0.1
M phosphate citrate pH 5, 0.2 M NaCl, 16% PEG 8000 and 1M guanidium hydrochloride.
For cryo-protection, they were transferred to a solution with 1.5X reservoir solution and
20% 2,3 butanediol or 20% DMSO for 10 seconds and immediately flash frozen under
liquid nitrogen. Form III crystals grew under 0.1 M phosphate citrate pH 5.5, 0.15 M sodium
sulfate, 0.01 M MgCl2, 1 M guanidium hydrochloride, and 28 % PEG 3350. The crystals
were grown at 30°C and were cryo-protected by increasing the PEG concentration to 40%.
Further details of crystallization are presented in the Supplementary Information.
Data collection and structure determination
Diffraction data were collected at the Dupont Northwestern Dow and Life Science
Collaborative Access Team stations (DND and LS CAT) at the Advanced Photon Source in
Argonne National Laboratory. Data collection and refinement statistics are shown in Table I.
All data were processed and integrated using XDS (Kabsch, 1993) and scaled with SCALA
(Collaborative-Computational-Project-4, 1994). Data on the Topo-31 crystals were collected
to 2.4 Å resolution. The structure was solved by Molecular Replacement (McCoy et al.,
2007) using the topoisomerase domain from the Topo-61 structure (residues 1-266) (Taneja
et al., 2006) as the search model. It was refined with refmac5 (Murshudov et al., 1997) and
Phenix (Afonine et al., 2005) to a final Rwork of 20.0 % and Rfree of 24.8 %. Topo-44 Form I
crystals diffract to 1.8 Å. The structure of Form I crystals was solved by Molecular
Replacement (McCoy et al., 2007) using the topoisomerase domain from the Topo-61
structure as the search model. Model rebuilding was performed using coot (Emsley and
Cowtan, 2004), and refinement using refmac5 (Murshudov et al., 1997). The final Rwork and
Rfree are 17.5 % and 22.0 % respectively.
For Topo-44 Form II and Form III crystals, seleno-methionine derivatized crystals were
used for single-wavelength anomalous dispersion (SAD) phasing. AutoSharp (Vonrhein et
al., 2007) was used for locating the selenium atoms; model building was done using coot
(Emsley and Cowtan, 2004), and refinement was carried out using refmac5 (Murshudov et
al., 1997) Three phosphate ions were noticed in the Form II structure; two of which present
in the topoisomerase active site and are separated by a distance of ~7.5 Å. The structure was
refined to a final Rwork of 24.1 % and Rfree of 28.9 %. Topo-44 Form III crystals diffracted
to 1.4 Å. The final Rwork and Rfree are 16.5 % and 18.4%, respectively. An interesting
observation is the presence of both phosphate and guanidium ions in the Form III Topo-44
structure. The linker helix and part of the first HhH motif of the B monomer show alternate
conformations and were built as two separate chains with occupancy of 0.5 each. Further
details on data collection and structure determination are given in the Supplementary
Information.
All figures were made with Pymol (DeLano, 2002) and the electrostatic surfaces were
calculated with APBS (Baker et al., 2001).
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Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We acknowledge staff and instrumentation support from the Keck Biophysics Facility and the Center for Structural
Biology at Northwestern University, and DND and LS-CAT at the Advanced Photon Source (APS) at Argonne
National Laboratory. Support from the R.H. Lurie Comprehensive Cancer Center of Northwestern University to the
Structural Biology Facility is also acknowledged. DND-CAT is supported by Dupont, DOW and the NSF. LS-CAT
was supported by the Michigan Economic Development Corporation and the Michigan Technology Tri-Corridor.
Use of the APS is supported by the Department of Energy (DOE). Research was supported by NIH grant GM51350
(to AM).
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Figure 1. Organization of topoisomerase V
Topoisomerase V is a multi-domain protein consisting of 24 helix-hairpin-helix (HhH) DNA
binding motifs arranged as 12 (HhH)2 domains following the N-terminal topoisomerase
domain. A) Schematic diagram of various topoisomerase V fragments. The topoisomerase
domain is shown in red, the (HhH)2 domains are shown in alternating colors of cyan and
yellow. The (HhH)2 domains with repair activity are shown in green. All fragments shown
have topoisomerase activity, but only the full length protein and the Topo78 fragment have
repair activity. B) Crystal structure of Topo-61 fragment (Taneja et al., 2006). The coloring
scheme is the same as in Figure 1A, except that the linker helix is shown in grey.
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Figure 2. DNA relaxation activity and EMSA for Topo-31, Topo-44 and Topo-78 fragments of
topoisomerase V
A) pH profile of the DNA relaxation activity of Topo-78 and Topo-44 fragments. 0.2 μg of
pUC19 DNA were incubated with 0.1 μg of Topo-78 or 1.5 μg of Topo-44 proteins at 65°C
for 15 minutes for the relaxation reaction. The reaction buffer contained 50 mM of the
appropriate buffer, 30 mM NaCl and 0.2 mM EDTA. Topo-78 relaxes DNA at a wider pH
range (5 to 9) than Topo-44, which relaxes DNA efficiently only at pH 5. DNA relaxation
activity of Topo-31 (B) and Topo-44 (C) fragments in the absence and presence of MgCl2.
0.2 μg of pUC19 DNA were incubated with 9 μg of Topo-31 or 1.5 μg of Topo-44 proteins
at 65°C for 15 minutes for the relaxation reaction. The reaction buffer contained 50 mM of
the appropriate buffer, 30 mM NaCl and 5 mM EDTA or 1 mM MgCl2. Both Topo-31 and
Topo-44 fragments can relax DNA in the absence of MgCl2, but MgCl2 enhances the DNA
relaxation activity of the topoisomerase V fragments. The black triangle in panels A, B and
C represents increasing pH from 4 to 10 by one pH unit. D) EMSA of Topo-44 and Topo-78
fragments with a 39mer double stranded DNA. Both Topo-44 and Topo-78 form stable
complexes with DNA, although Topo-78 seems to saturate DNA binding while Topo-44
does not. In addition, Topo-44 shows some cleavage of the DNA (bottom free DNA band),
while the cleavage is not apparent in Topo-78. The numbers at the bottom represent the
molar ratio of protein to DNA.
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Figure 3. Structure of Topo-44 fragments
A) Overlay of Form I (green), Form II (B subunit: blue), and Form III (B subunit: magenta)
structures onto the Topo-61 structure (B subunit: orange). The topoisomerase domains
superimpose very well for all the structures, while the linker helix and (HhH)2 domains
show differences in orientation. B) Overlay of the linker helices of Form I, II, and III
structures with that of Topo-61. The color scheme is same for all the figures unless
mentioned otherwise. Note that the linker helices have the same orientation at the start and
they change as they move further down the helix. C) Superposition of Form I, II, and III
Topo-44 structures with that of Topo-61. Only the (HhH)2 domains are colored while the
remaining parts are shown in gray for clarity. The active site residues are shown as orange
sticks. Note that the (HhH)2 domains adopt different orientations in all the structures. D)
Orientation of the (HhH)2 domains of Form I, II and Topo-61 structures. In Form I and II
structures, the (HhH)2 domains are moved away from the topoisomerase domain. For clarity,
the (HhH)2 domains of Form III are not shown. In panels C and D, the topoisomerase
domains were superposed to emphasize the different orientation of the other domains.
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Figure 4. Phosphate ions present near the active site of the Topo-44 structure
A) Stereo view of a Form III difference electron density map calculated with a model not
including the phosphates. The electron density is contoured at 3.7σ and shows the
tetrahedral shape of the phosphate ions. The active site residues are shown in stick. B)
Stereo view of the interaction of the phosphate ions with the putative active site residues.
The B subunit of Form II structure was superimposed onto the B subunit of Form III
structure and the phosphates ions from both structures are shown together with the Form II
B subunit protein backbone. The interactions made by the phosphate ion with the active site
residues and the corresponding distances in Å are represented as black dotted lines.
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Figure 5. Representation of the unique phosphate ions from Form II and Form III Topo-44
structures
A) An overlay of the A (magenta) and B (brown) subunits of the Form III structure and B
(blue) subunit of Form II Topo-44 structures. The positions of eight unique phosphate ions
(orange spheres) are shown. Note that most phosphate ions are found along the DNA
binding groove of the topoisomerase domain. B) The phosphate ions in the DNA binding
groove are separated by distances of around 7 Å. The protein backbone is that of the B
subunit of Form III structure. The active site residues are represented as sticks and distances
in Å between adjacent phosphate ions are shown as black dotted lines.
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Figure 6. Model showing DNA bound to the topoisomerase domain
A) Model of a 17-mer double stranded DNA bound to the Topo-31 structure (teal). The
DNA is represented as green sticks, where as phosphate ions are represented as orange
sticks. DNA binds along the DNA binding groove and five of the eight phosphate ions noted
in the Topo-44 structures coincide with the DNA backbone. B) Model of Topo-44 (Form II,
B subunit: blue) binding to 17-mer double stranded DNA. Note that the linker helix and the
(HhH)2 domains interfere with DNA binding to the topoisomerase domain and are likely to
move away to allow binding. C) Electrostatic surface representation of the Topo-31
structure. The positively charged DNA binding groove is clearly visible and the phosphate
ions are bound in this groove. The orientation corresponds to a 90° rotation of the one shown
in Figure 6A in the direction of the arrow. Note that the DNA binding groove goes from one
end of the molecule to the other and it is narrower at one end (start of the linker helix) and
wider at the other end. The putative active site residues (green sticks) are located at the
wider end of the groove. Other residues lining the groove and interacting with the phosphate
ions are shown as cyan sticks. D) Electrostatic surface representation of Topo-31 with
phosphate ions (orange) and DNA (green). Three phosphate ions (P3, P4, and P5) coincide
with the phosphates of one of the DNA strands, where as P1 coincides with a phosphate of
the opposite DNA strand. The model shows that the DNA binding groove of topoisomerase
V is wide enough to bind DNA and that the movement of linker helix and (HhH)2 domains
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are required to accommodate the DNA. The electrostatic potential was calculated with a
dielectric constant of 80 for solvent and 2 for protein. The surface is colored with a blue to
red gradient from +10 to −10 KbT/ec.
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Table 1
Data collection and refinement statistics
Topo-31
Topo-44 Form I
Topo-44 Form II
Topo-44 Form III
Data Collection
Space group
C2221
C121
P41212
P212121
Cell dimensions
a=106.7 Å, b=119.4
Å, c=63.7 Å
a=104.2 Å, b=47.7 Å,
c=81.2 Å (β=112.48)
a=b=70.1 Å, c=349.6 Å
a=63.6 Å, b=80.1 Å,
c=137.2 Å
Resolution (Å)a
79.56 – 2.4 (2.53 –
2.4)
75.05 – 1.82 (1.91 –
1.82)
29.5- 2.6 (2.72-2.6)
28.9-1.4 (1.46-1.4)
Number of observed
reflections
78,729 (11,538
134,411 (13,220)
227,408 (19,917)
1,157,917 (126,319)
Number of unique reflections
16,259 (2,346)
32,998 (4,301)
28,151 (3,331)
136,662 (15,986)
Completeness (%)
99.8 (99.8)
98.3 (88.6)
99.9 (100.0)
98.8 (95.5)
Multiplicity
4.8 (4.9)
4.1 (3.1)
8.1 (6.0)
8.5 (7.9)
Rmerge (%)b
4.7 (71.1)
4.0 (16.3)
7.4 (52.2)
4.5 (37.9)
Rmeas (%)c
5.3 (79.6)
4.6 (19.4)
7.9 (57.2)
4.8 (40.5)
≪I>/σ(<I>)>d
20.5 (2.5)
23.0 (6.8)
19 (3.2)
27.5 (5.3)
Refinement
Resolution (Å)
79.56 - 2.4 (2.46 -
2.4)
28.06 -1.82 (1.87 –
1.82)
29.14 – 2.6 (2.67 – 2.6)
28.9 - 1.4 (1.44 - 1.4)
Number of reflections
working/test
15,419/821
31,317/1,673
26,710/1,438
129,802/6,859
Rwork (%)e
20.0(24.3)
17.5 (17.9)
24.1(36.6)
16.5 (19.3)
Rfree(%)f
24.8 (31.1)
22.0 (24.8)
28.9 (45.1)
18.4 (22.1)
Protein residues/atomsg
269/2,203
376/3212
727/5,970
738/7,511
Atoms in alternate
conformations
0
258 (20 protein
residues)
8 (1 protein residue)
2846 (157 protein
residues)
Water molecules
29
238
30
573
Other atoms
-
-
3 PO4
7 PO4, 3 Gmh, 3 Mg++, 2
Cl−
B-factor (Å2)
Protein atoms (chain)
68.4
22.8
A:53.8; B:58.2
A:13.4; B:14.9
Water molecules
59.1
29.3
40.0
23.7
r.m.s. deviations
bond lengths (Å)
0.015
0.006
0.01
0.009
bond angles (°)
1.42
0.920
1.2
1.2
Ramachandran ploti
Favored regions (%)
94.3
98.9
96.2
98.5
Outliers (%)
0.0
0.0
0.3
0
aNumbers in parenthesis correspond to highest resolution shell.
bRmerge= Σ|I − <I>|/ΣI, where I is the observed intensity and <I> the average intensity obtained from multiple measurements.
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cRmeas as described in Diederichs and Karplus (Diederichs and Karplus, 1997).
d≪I>/σ(<I>)> = Mean Ih over the standard deviation of the mean Ih averaged over all reflections in a resolution shell.
eRwork= Σ ||Fo| − |Fc||/Σ|Fo|, where |Fo| is the observed structure factor amplitude and |Fc| the calculated structure factor amplitude.
fRfree: Rfactor based on 5% of the data excluded from refinement.
gTotal number of protein atoms, including those in alternate conformations.
hGm: guanidinum ion.
iAs reported by Molprobity (Davis et al., 2004).
Structure. Author manuscript; available in PMC 2011 July 14.
|
3M6O
|
Crystal structure of Arabidopsis thaliana peptide deformylase 1B (AtPDF1B)
|
Trapping Conformational States Along Ligand-Binding
Dynamics of Peptide Deformylase: The Impact of
Induced Fit on Enzyme Catalysis
Sonia Fieulaine1, Adrien Boularot1, Isabelle Artaud2,3, Michel Desmadril4,5, Fre´de´ric Dardel6,7, Thierry
Meinnel1*, Carmela Giglione1*
1 CNRS, ISV, UPR2355, Gif-sur-Yvette, France, 2 Universite´ Paris Descartes, UMR8601, Paris, France, 3 CNRS, UMR8601, Paris, France, 4 Universite´ Paris-Sud, IBBMC,
UMR8619, Orsay, France, 5 CNRS, IBBMC, UMR8619, Orsay, France, 6 Universite´ Paris Descartes, UMR8015, Paris, France, 7 CNRS, UMR8015, Paris, France
Abstract
For several decades, molecular recognition has been considered one of the most fundamental processes in biochemistry.
For enzymes, substrate binding is often coupled to conformational changes that alter the local environment of the active
site to align the reactive groups for efficient catalysis and to reach the transition state. Adaptive substrate recognition is a
well-known concept; however, it has been poorly characterized at a structural level because of its dynamic nature. Here, we
provide a detailed mechanism for an induced-fit process at atomic resolution. We take advantage of a slow, tight binding
inhibitor-enzyme system, actinonin-peptide deformylase. Crystal structures of the initial open state and final closed state
were solved, as well as those of several intermediate mimics captured during the process. Ligand-induced reshaping of a
hydrophobic pocket drives closure of the active site, which is finally ‘‘zipped up’’ by additional binding interactions.
Together with biochemical analyses, these data allow a coherent reconstruction of the sequence of events leading from the
encounter complex to the key-lock binding state of the enzyme. A ‘‘movie’’ that reconstructs this entire process can be
further extrapolated to catalysis.
Citation: Fieulaine S, Boularot A, Artaud I, Desmadril M, Dardel F, et al. (2011) Trapping Conformational States Along Ligand-Binding Dynamics of Peptide
Deformylase: The Impact of Induced Fit on Enzyme Catalysis. PLoS Biol 9(5): e1001066. doi:10.1371/journal.pbio.1001066
Academic Editor: Gregory A. Petsko, Brandeis University, United States of America
Received January 7, 2011; Accepted April 14, 2011; Published May 24, 2011
Copyright: 2011 Fieulaine et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by the Centre National de la Recherche Scientifique (CNRS, France), grant ANR-06-MIME-010-01 (Agence Nationale de la
Recherche, France), and grant #4920 from the Association pour la Recherche sur le Cancer (Villejuif, France). SF was partly supported by a postdoctoral fellowship
from the Fondation pour la Recherche Me´dicale (France). AB was supported by CNRS, France. The funders had no role in study design, data collection and
analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
Abbreviations: DSC, differential scanning calorimetry; Fo, formyl; PDF, peptide deformylase; r.m.s.d., root mean square deviation.
* E-mail: Thierry.Meinnel@isv.cnrs-gif.fr (TM); Carmela.Giglione@isv.cnrs-gif.fr (CG)
Introduction
Flexibility of proteins around their active site is a central feature
of molecular biochemistry [1–5]. Although this has been a central
concept
in
biochemistry
for
half
a
century,
the
detailed
mechanisms describing how the active enzyme conformation is
achieved have remained largely elusive, as a consequence of their
transient
nature.
Direct
structural
evidence
and/or
kinetic
analyses have only recently emerged [6–10]. Three classic
‘‘textbook’’ models are used to describe the formation of the
ligand-enzyme complex: (i) the Fischer’s ‘‘lock-and key’’ model, (ii)
the Koshland’s induced-fit model, and (iii) the selected-shift model
or conformational selection mechanism [6–8,11–13]. In the
Fischer’s ‘‘lock-and key’’ model, the conformations of free and
ligand-bound proteins are essentially the same. In the induced-fit
model, ligand binding induces a conformational change in the
protein, leading to the precise orientation of the catalytic groups
and implying the existence of initial molecular matches that
provide sufficient affinity prior to conformational adaptation [14].
In contrast, the selected-fit model assumes an equilibrium between
multiple conformational states, in which the ligand is able to select
and stabilize a complementary protein conformation. In this case,
the conformational change precedes ligand binding, in contrast to
the induced-fit model in which binding occurs first. The
conformational selection and/or induced-fit processes have been
shown to be involved in a number of enzymes [12,13,15,16]. For
several of these studies, conformational selection is proposed
because the experimental data support that, even in the absence of
the ligand, the enzyme samples multiple conformational states,
including the ligand-bound (active) state [6]. Although direct
structural evidence and/or kinetic analyses have provided clues
[6–8,12,13,16], how we can distinguish whether a protein binds its
ligand in an induced- or selected-fit mechanism remains critical
and often controversial.
The enzyme-inhibitor interaction is a form of molecular
recognition that is more amenable to investigation than the
enzyme-substrate interaction as there is no chemical transforma-
tion of the ligand during this process. In this context, slow, tight-
binding inhibition is an interesting interaction process, as it closely
mimics the substrate recognition process and has been shown to be
commonly involved in adaptive conformational changes [12,
17,18]. In slow, tight-binding inhibition, the degree of inhibition at
a fixed concentration of compound varies over time, leading to a
curvature of the reaction progress curve over time during which
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the uninhibited reaction progress curve is linear [19]. Indeed, the
slow, tight-binding inhibition is a two-step mechanism that
depends on the rate and strength of inhibitor interactions with
the enzyme. Binding of the inhibitor (I) to the enzyme (E) leads to
the rapid formation of a non-covalent enzyme-inhibitor complex
(E:I) followed by monomolecular slower step (k5) in which the E:I is
transformed into a more stable complex (E:I*) that relaxes and
dissociates at a very slow rate, mainly inferred by the k6 value
when k6,,k5,,k4, (Figure 1A; see also footnote f in Table 1).
Although only a few studies have investigated the mechanisms
of slow, tight-binding inhibitors, such molecules are favored for use
as
therapeutics,
as
they
usually
exhibit
unique
inhibitory
properties, including selective potency and long-lasting effects
[20–26]. Here, we explore the precise structural inhibitory
mechanism of actinonin (Figure 1A; [27]), which is a slow, tight-
binding inhibitor of peptide deformylase (PDF), a metal cation-
dependent enzyme [28,29]. The function of the active-site metal is
to activate the reactive water molecule involved in peptide
hydrolysis [30]. PDF is the first enzyme in the N-terminal
methionine excision pathway, an essential and ubiquitous process
that contributes to the diversity of N-terminal amino acids [31,32].
Actinonin is a natural product with antibiotic activity that inhibits
PDF by mimicking the structure of its natural substrates (nascent
peptide chains starting with Fo-Met-Aaa, where Fo is a formyl
group and Aaa is any amino acid) in their transition state
(Figure 1B). The transition state inhibitor actinonin, as well as
other structurally related inhibitors, has been shown to systemat-
ically exhibit a ‘‘slow-binding’’ inhibition behavior (Figure 1A),
regardless of the organism of origin of the PDF [29,33].
Using structural, biocomputing, and enzymatic analyses, we
were able to (i) reveal that the free enzyme is in an open
conformation and that actinonin induces transition of the enzyme
into a closed conformation; (ii) show that there is no evidence for
the occurrence of a closed conformation in the apostructure of the
open enzyme, which, together with detailed kinetic analyses,
makes the closed form fully compatible with an induced-fit model;
and (iii) identify the sequence of molecular events leading to the
final, bound, closed complex (E:I*). Moreover, using several
rationally designed point mutants of the enzyme, ligand-induced
intermediates, which mimic conformational states that normally
would not be expected to accumulate with the wild-type (WT)
enzyme, were trapped. These conformations recapitulate physical
states that the WT enzyme must pass through during its overall
transition from the apo-enzyme to the E:I* complex. ‘‘Freezing’’ of
ligand-induced intermediate states provides direct evidence for an
induced-fit mechanism and allows the reconstruction of a virtual
‘‘movie’’ that recapitulates this mechanism. Since PDF is one
example of an enzyme remaining active in the crystalline state and
because actinonin closely mimics the natural substrates bound to
PDF in the transition state as shown previously with the Escherichia
coli form (EcPDF; see Figure 1B) [34,35], we propose a model
suggesting that induced fit also contributes to efficient catalysis.
Results
Slow, Tight Binding of the Transition-State Analog
Actinonin to Peptide Deformylase
In the present study, at the atomic level we explored the precise
inhibitory mechanism of actinonin on Arabidopsis thaliana PDF1B
(AtPDF), a close eukaryotic homologue of EcPDF (Figure S1)
[36,37]. Measurements of the kinetic parameters of the second step
of the binding mechanism (k5) revealed a timescale in the 10-s range
(Table 1), which is consistent with the collective motion of a large
domain [4,5]. This finding is supported by NMR studies [38,39],
which showed that actinonin binding induces drastic changes in the
heteronuclear single quantum coherence (HSQC) spectrum of
EcPDF, since most resonances undergo significant shifts that affect a
large part of the structure [40,41]. The existence of alternative
conformational states of EcPDF is further supported by recent
biophysical studies [42]. Previously reported snapshots of a series of
different conformations of the enlarged and mobile loop—the so-
called CD loop—of the dimeric PDF from Leptospira interrogans PDF
(LiPDF) in the presence or absence of inhibitor led to the hypothesis
of the existence of an equilibrium between a closed and open form
of the CD-loop of PDF enzymes, suggesting a selected-shift model to
the authors [43]. Taken together, these data suggest that the binding
of actinonin to PDF is accompanied or preceded by conformational
changes within the enzyme. Paradoxically, this proposal has not
been currently supported by the available structural data. Indeed,
free and complexed crystal structures have provided no evidence for
any significant conformational change in PDF structure induced by
the binding of ligand [35,43–47].
Tight inhibition in the closed state is associated with the KI*
apparent equilibrium constant (Figure 1A). A KI* value (see Table 1
and Materials and Methods for the biochemical definition of KI*)
of 0.9 nM for actinonin could be measured for AtPDF; that is, a
value very similar to that obtained for bacterial PDFs, including
EcPDF and Bacillus stearothermophilus PDF2 (BsPDF2, Table 1).
Tightening of the initial encounter complex (E:I) resulted in a final
complex (E:I*) in which the potency of actinonin (KI/KI*) was
enhanced by more than two orders of magnitude and exhibited a
very slow off-rate (k6, Table 1). The dissociation constant value of
AtPDF for actinonin was also assessed using isothermal titration
calorimetry (ITC) experiments (Table S1 and Figure S2A). The
corresponding ITC titration curves (Figure S2A) are consistent
with a very strong affinity of the ligand for the enzyme [48],
enabling us to determine an accurate Kd. Moreover, these studies
generated values similar to those measured by other means for
AtPDF and EcPDF [42,49].
Author Summary
The notion of induced fit when a protein binds its ligand—
like a glove adapting to the shape of a hand—is a central
concept of structural biochemistry introduced over 50
years ago. A detailed molecular demonstration of this
phenomenon has eluded biochemists, however, largely
due to the difficulty of capturing the steps of this very
transient process: the ‘‘conformational change.’’ In this
study, we were able to see this process by using X-ray
diffraction to determine more than 10 distinct structures
adopted by a single enzyme when it binds a ligand. To do
this, we took advantage of the ‘‘slow, tight-binding’’ of a
potent inhibitor to its specific target enzyme to trap
intermediates in the binding process, which allowed us to
monitor the action of an enzyme in real-time at atomic
resolution. We showed the kinetics of the conformational
change from an initial open state, including the encounter
complex, to the final closed state of the enzyme. From
these
data
and
other
biochemical
and
biophysical
analyses, we make a coherent causal reconstruction of
the sequence of events leading to inhibition of the
enzyme’s activity. We also generated a movie that
reconstructs the sequence of events during the encounter.
Our data provide new insights into how enzymes achieve a
catalytically competent conformation in which the reactive
groups are brought into close proximity, resulting in
catalysis.
The Dynamics of Induced Fit at High Resolution
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Ligand-Induced Conformational Closure of AtPDF in the
Crystalline State
Occurrence of a conformational change induced by drug binding
was visualized via the resolution of several crystal structure forms of
AtPDF, the free form and/or in a complex with actinonin (Table
S2). The data reveal a structural switch between the two forms that
can account for both the thermodynamic and kinetic data. The
enzyme was observed in two states, a novel open apo-form and a
closed, induced, actinonin-bound complex (Figure 1C). Binding of
actinonin resulted in a tightening of the active site through the
collective closure of the entire N-terminal portion of the protein
(strands b1, b2, and b3; helix a1; and CD-loop, see Movies S1 and
S2, Figure 1C, and Figure S1). The amplitude of the structural
change was maximal for Pro60 (Figure S1), the Ca of which was
shifted 4 A˚ upon actinonin binding. This collective movement
involved the formation of a ‘‘super b-sheet’’ as the result of the large
rearrangement of b-strands 4 and 5 relative to the rest of the
structure in which actinonin forms an additional strand bridging the
two b-sheets (b1 andb2) on either side of the active site (Figure 1D
and Figure S1B). As actinonin is a peptide-like compound (see
Introduction and Figure 1B), this behavior closely mimics what
occurs in the natural protein substrates of PDF, which also form this
strand-bridging interaction. This phenomenon also accounts for the
strong stabilization of the protein by actinonin, which was also
challenged by differential scanning calorimetry (DSC) experiments:
the Tm of AtPDF increased from 61uC to 81uC upon binding of the
inhibitor (Figure 1D, see also below).
Thus far, this closure of the enzyme induced by actinonin is part
of the rare structural evidence for the slow, tight-binding
mechanism at an atomic scale. The open state, which has never
been observed, was captured not only in the two molecules of the
asymmetric subunit but also in different crystals and under two
distinct crystallization conditions (Table S2 and Figure 2). All
r.m.s.d. values were smaller than 0.25 A˚ . The closure is very
unlikely to result from crystal packing constraints, as soaking the
apo-AtPDF crystals in a solution containing actinonin induced the
Figure 1. Slow, tight-binding inhibition of PDF by actinonin induces conformational change in the protein. (A) Inhibition by a two-step
mechanism, involving a tightening of the initial enzyme-inhibitor complex (E?I) to form a more stable complex (E?I*), with the chemical structure of
actinonin (I), the natural inhibitor of PDF enzymes (E). (B) Structures of EcPDF bound to actinonin (left) and to the transition state resulting from the
cleavage of its substrate, Fo-Met-Ala-Ser (right) [34,35]. (C) Superimposition of free and actinonin-bound AtPDF indicated in green and purple,
respectively. The three conserved motifs of the PDF enzymes family are indicated in orange and numbered I, II, and III. Molecules A of both models
were superimposed, resulting in an r.m.s.d. of 0.9 A˚ for 100% of the Ca. Left inset, close-up comparison of the open and closed forms figured in the
ribbon representation. (D) Baseline-corrected DSC thermograms of free and actinonin-bound WT AtPDF recorded under the same experimental
conditions.
doi:10.1371/journal.pbio.1001066.g001
The Dynamics of Induced Fit at High Resolution
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structural transition from the open to the closed state within the
crystals without cracking them or altering their diffracting power.
Thus, crystal packing is compatible with both states of the enzyme
(Figure S3). Therefore, the open structure most likely corresponds
to a stable state in solution.
The closed final conformation was identical to that previously
reported for PDF complexes obtained either with actinonin or with a
product of the reaction [34,35,44,50], indicating that this structure is
common for the ligands (compare Figures 1B and 2A, and Figure S4).
Hydrogen bonding was also conserved, especially the bond between
the backbone nitrogen of Ile42 (corresponding to Ile44 in EcPDF, see
Figure 1B and Figure S5A) and the alkyl carbonyl chain of actinonin,
which potently contributes to the formation of the super b-sheet
(Movie S2 and Figure S1B, see also below). Between the open and
closed states, the side chains of Ile42, Phe58, and Ile130 underwent
significant structural changes (Figure 3A and D and Figure S6),
corresponding to a hydophobic pocket rearrangement, with Ile42
being the most affected (Figure 3). Interestingly, Ile42 is the second
residue of the conserved active-site motif G41IGLAAXG (motif 1)
that was previously shown to be essential for activity [51].
To assess and visualize the differences between the two states, two
independent structural parameters were measured: the r.m.s.d.
value with respect to the open form and the aperture angle (dap),
which measures the angle made between the N- and C-domains
through three fixed-points, corresponding to the Ca of three
conserved residues, each sitting in one of the three conserved motifs
(Figure 2A). The bi-dimensional graph of these two parameters is a
good representation of the closing motion snapshots (Figure 2B)
shown in Movie S1. With this tool at this stage, two states could be
defined: the closed (C) and open (O) states (Figure 2B).
Evidence for a Pure Induced-Fit Mechanism in the
Binding of Actinonin to AtPDF
Recent quantitative analyses of both conformational selection
and induced fit have led to an integrated continuum—a so-called
‘‘flux-description’’—of
these
two
limiting
mechanisms
[16].
According to this model, conformation selection tends to be
preferred at low ligand concentrations (mM range)—that is, using
detailed kinetic studies—whereas induced fit dominates at high
ligand and enzyme concentrations (mM range) obtained, for
instance, in NMR or crystallographic approaches. Structural
studies are most useful to reveal subpopulations of biological
significance.
We investigated the existence of lowly populated, alternative
conformations of apoPDF. To probe the occurrence of alternate
conformers in the crystalline state of PDF, the new Ringer
program is the most suitable investigation tool [52,53]. Ringer
searches for evidence of alternate rotamers by systematically
sampling electron density maps—free of model bias—around the
dihedral angles of protein side chains. Two independent WT open
datasets of the apoenzyme, including a high-resolution set (1.3 A˚ ),
were used in the analysis. Ringer analysis revealed the existence of
only one rotamer of most side chains of either molecule in the
asymmetric unit, including the three main residues primarily
involved in conformation change—that is, Ile 42, Phe58, and
Ile130
(Figure
4A).
Ringer
analysis
showed
evidence
for
unmodeled alternate conformers for very few residues, including
Ile121 and Phe87, or Phe119 to a much lesser extent (Figure S7).
There is therefore no evidence for the occurrence of a closed
conformation in the apostructure of AtPDF, supporting the
hypothesis that the conformational change was essentially induced
by the binding of actinonin rather than from conformational
selection among multiple states occurring in the crystalline state.
To further investigate the mechanism involved, we followed a
kinetic approach aimed at discriminating between induced fit and
population shift at low ligand concentrations (sub-mM range) [12].
The experimentally observed pseudo-first-order rate constant for
the approach to equilibrium between the free components and the
binary AtPDF-actinonin complex (kobs) was measured and plotted
as a function of actinonin concentration. This plot yielded a
hyperbolic saturation curve with a positive slope, as fully expected
for a pure induced-fit mechanism (Figure 4B and C). In contrast, if
the enzyme sampled two or more conformational states, the curve
would imply that the value of kobs decreases with increasing ligand
concentration (see, for instance, curve C in Figure 1 in [12]). The
same conclusion can be reached for EcPDF and BsPDF2
(Figure 4B and C) and was already reported by others for S.
aureus PDF [29].
Together, these data indicate that a pure induced-fit mechanism
triggered by the binding of actinonin appears to direct the
conformational change both in solution and in the crystalline state.
Single Variants at Gly41 Exhibit Strongly Reduced
Actinonin-Binding Potency and Catalytic Efficiency
When dealing with an induced-fit mechanism, knowledge of the
initial O and final C state is crucial but does not provide direct
information on the position of actinonin in the encounter complex
or on the sequential mechanism of the transition process. We
suspected that the conserved glycine-rich motif 1 (G41IGLAAXQ)
could contribute to the flexibility required for the observed
structural transition. Evidence for such flexibility comes from
NMR analysis of EcPDF in which a few residues show exchange
cross-peaks of an additional, alternative form [38]. The most
strongly affected residues are Cys90, one of the metal ligands, its
neighbor Leu91, and both of the alanines within the above
conserved glycine-rich motif (Figure S1B), suggesting that EcPDF
undergoes conformational dynamics in a similar region.
To unravel the dynamics of the recognition process, we
surmised that it should be possible to freeze the conformational
Table 1. Comparison of the main kinetic and thermodynamic
parameters describing the inhibition of PDF by actinonin.
Parameter
AtPDFa
EcPDFa
BsPDF2a,b
KI (nM)d
140610
112610
185615
KI* (nM)c
0.960.5
1.360.2
2.960.8
KI/KI*
155615
86610
6467
k5 (s21) 6103d
6369
170620
7268
k6 (s21) 6104d
461
1962
1163
k4 (s21)e
140610
112610
185615
t1/2 (min)f
2965
661
1.160.2
aThe enzyme concentrations used in the assay were 100, 50, and 25 nM for
AtPDF, EcPDF, and BsPDF2, respectively.
bData from [49].
cPrior to kinetic analysis for determination of the KI* value, actinonin was
incubated at the final concentration in the presence of the studied enzyme set
for 10 min at 37uC. The kinetic assay was initiated by the addition of a small
volume of the substrate.
dFor determination of KI, k5, and k6 values, actinonin was not preincubated with
the enzyme. The kinetic assay was initiated by the addition of the enzyme.
ek4 corresponds to the kinetic constant of the dissociation of the primary
enzyme-actinonin complex. It is assumed that the rate of complex association
is diffusion-limited (see Table 7.3 in [19]), that is, k3—the kinetic constant of the
association of the primary enzyme-actinonin complex—is 109 M21.s21.
ft1/2 is 0.693(k4+k5+k6)/k4k6 (see case of induced fit and calculation in Table 1 of
[12]). In this case, t1/2,0.693/k6 because k6,,k5,,k4.
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Figure 2. Four distinct conformational states of PDF enzymes. (A) AtPDF and several other representative bacterial PDFs are superimposed. A
zoom is displayed on the right of the panel. Superimpositions were realized using ‘‘module superpose’’ in the CCP4i package and the ‘‘secondary
structure matching’’ tool. The extent of aperture/closure of PDF enzymes was assessed primarily by the measurement of the aperture angle (dap), the
angle made between the Ca of three strictly conserved residues (C, H, and I) of all PDFs, each characterizing a secondary crucial structure module of
the active site crevice, namely b4, a2, and b1 (see Figure S1C). Each single residue belongs to one of the three conserved motifs (motifs 2, 3, and 1,
respectively) and corresponds respectively to Cys91, His137, and Ile42 in AtPDF. The dap was measured in each case (see B). (B) The dap values
combined with those of the r.m.s.d. associated with the superimposition of the open structure of AtPDF allows the identification of four
conformational states: open (O), intermediate (I), closed (C), and super-closed (S). We compared AtPDF1B (this work and PDB CODE 3CPM; brown,
orange, and yellow in A and B; black in C), EcPDF (1BS7, free enzyme; 1BS6, with Met-Ala-Ser; 1G2A, with actinonin; magenta), BsPDF2 (1LQY, with
actinonin; green), LiPDF (1SV2, free; 1SZZ, with actinonin; red), and PfPDF (1JYM, free; blue).
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Figure 3. Effect of actinonin binding on the conformation of key residues in PDF. Conformation of key residues Ile42, Phe58, and Ile130 in
the different complexes: (A) in unbound WT AtPDF, (B and C) in the structure of G41Q and G41M actinonin-bound variants, respectively, and (D) of
actinonin-bound WT protein. In the final complex (D), a hydrogen bond is formed between actinonin and the peptidic bond, which links Gly41 and
Ile42. During the deformylation reaction, which is catalyzed by the PDF enzyme, the N-terminal formyl-methionine fits into the S1’ pocket. The
solvent-accessible surface of this pocket is represented here, and only the aliphatic chain of actinonin is shown, mimicking the N-terminal
methionine. (E) Free WT enzyme with the S1’ pocket shown open in two orientations (top and bottom). (F and G) S1’ pocket in the G41Q and G41M
variant structures, respectively, shown in two orientations (top and bottom). (H) After the complete conformational modifications of actinonin-bound
WT protein induced by actinonin binding, the S1’ pocket is shown closed in two orientations (top and bottom). (I) The four models are superimposed;
the ligand-binding site is magnified: unbound WT AtPDF; G41Q and G41M actinonin-bound enzyme; and WT actinonin-bound enzyme are indicated
in brown, red, orange, and yellow, respectively. Actinonin is indicated by lines. (J) A detailed view of the AtPDF ligand-binding site for all the
complexes, which are superimposed, as indicated in the same colors. Arrows indicate the direction of the closing movement within the enzyme, from
the open, unbound state to the closed, bound state.
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change along the pathway by introducing selected, minor
variations within the above-mentioned crucial residues involved
in the collective motion. In this respect, site-directed mutagenesis
of AtPDF was performed on Gly41, Ile42, and Ile130. Single
substitutions were made at Gly41 (G41A/Q/M), Ile42 (I42A/F/
N/W), and Ile130 (I130A/F), and the variants were purified and
characterized. These mutant proteins showed no change in overall
stability, as evidenced by DSC experiments (unpublished data).
However, two variants of G41, G41Q and G41M, showed
dramatic effects; the kcat/Km values were reduced by three orders
of magnitude due to large decreases in the kcat values compared to
the WT enzyme (Figure 5A and Table S1). The reduced kcat/Km
values suggest an altered ability of these variants to attain the final
enzyme-transition state complex and, as a result, to give rise to
possible states different from the final E:I* complex. Substitutions
at positions 42 and 130 only caused small reductions in the kcat
values (Figure 5A, Figure S2C, and Table S1). The actinonin-
binding potency of both G41 variants was also greatly reduced
(Table S1 and Figure S2B). The time-dependent inhibition by
actinonin of the most active variants was then studied (Table S3).
Figure 4. Evidence for an induced fit in crystalline and solution states of AtPDF. (A) Absence of evidence for alternative conformers in the
crystalline state of AtPDF. Ringer plots of electron density (r) versus x1 angle for representative residues of the 3D apo-structure of AtPDF. Data were
obtained with the 3M6O, 3PN2, and 3PN3 datasets (2.0 and 1.3 A˚ resolution, respectively, see Table S1). The secondary peaks in the Ile residues are
observed because Ile is a branched amino acid. To reveal an alternative conformation with Ile, three peaks should be observed. (B) kobs is a saturable
function of actinonin with various PDFs, including AtPDF. Data obtained for kobs, the experimentally observed pseudo-first-order rate constant for the
approach to equilibrium between the free components and the binary PDF-actinonin complex, were obtained at various concentrations of actinonin
in the presence of EcPDF, AtPDF, and BsPDF2. A direct plot is shown. Inset, time-course measurement of deformylation as a function of varying
actinonin concentrations. (C) Inverted plot of the data in panel B, which is expected to be a straight line if the kobs is ..k6 in the case of induced fit
[19]. The correlation coefficient of each line is 1.00, 0.99, and 1.00 for AtPDF, BsPDF2, and EcPDF, respectively, indicative of the accuracy of the
conclusion.
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The half-lives of the final complexes—as assessed by comparison
of the 1/k6 values—were always significantly smaller (Table S3),
suggesting that the conformational change induced by actinonin
binding still occurred, but the C state is destabilized relative to the
O state in the mutants compared to the WT. Accordingly,
actinonin strongly stabilized almost all of the variants; Tm was
increased by more than 20uC. This differs from the G41M and
G41Q variants, which both showed increases in the Tm of only
12uC, consistent with reduced binding potency (Table S1).
Conformational Changes of Gly41 Variants Are Affected
On-Pathway
The two most interesting variants, G41Q and G41M, could be
crystallized under the same conditions as the WT protein. In the
case of G41Q, the structure of the apo-protein did not show any
modifications compared to the WT structure and remained in an
O conformation (Figure 2B; ‘‘O’’ zone). In contrast, the 3D
structure of the G41M variant showed that the asymmetric unit
was composed of two molecules with distinct structures. One
molecule (chain A) is in the O state and is similar to the structures
of the WT and the G41Q variant (Figure 2B; zone ‘‘O’’). The
second molecule (chain B) is in a C state, closer to that observed
for the WT chain in the presence of actinonin (‘‘C’’), a so-called
‘‘superclosed’’ state (Figure 2B; zone ‘‘S’’), suggesting that the
substitution modified the equilibrium between the two states in
solution either (i) at the step of protein synthesis by providing two
conformers, the inter-conversions of which are blocked due to
steric hindrance brought by the new bulkier side-chain at position
41, or (ii) by dramatically unbalancing the free inter-conversion
between the O and S conformers towards the S state. Ringer
analysis indicates that in the free G41M variant, many residues
show evidence for unmodeled alternate conformers—including
positions 58, 42, and 130—in keeping with the second hypothesis.
For all variants of position G41, addition of actinonin to the
crystal (Figure 3 and Figure S6) induced a closure of the protein
within the crystal. Nevertheless, as expected from in silico graphic
modeling followed by energy minimization, the occurrence of a
bulky side chain at position 41 prevented the completion of the
closure in the presence of the ligand and, hence, the formation of
the hydrogen bond between the backbone nitrogen of Ile42 and
actinonin. This finding is consistent with the strongly reduced Tm
of the complex of the variants with actinonin compared to WT as
measured by DSC. Remarkably, both S and O forms of the G41M
apo-structures in the asymmetric unit of the crystal yielded a
unique intermediary structure (‘‘I’’ state) upon actinonin binding
(r.m.s.d. between the molecules is ,0.25 A˚ ; see also Figure 2B,
zone ‘‘I’’). In this case, it is likely that the induced-fit mechanism
drives the equilibrium by capturing only the O population and
closing it to an intermediary step, thus depleting the pool of O
conformers that is shifted sequentially back from the remaining
pool of S conformers and allows the complete binding of actinonin
to the enzyme.
In line with the rational design of the PDF mutants, the extent
of the structural differences suggests that the underlying motions
are dependent on the length of the side chain (Figure S8).
Together, these data account for the reduced catalytic rate, as the
hydrogen bond is strictly required for the substrate to be efficiently
cleaved by PDFs (Figure S8A) [54]. Therefore, from both
structural and kinetic analyses, each substitution most likely
reproduces intermediates along the pathway that lead to the
closure of PDF around its substrate (Figure S2B).
Conformational Changes of Gly41 Variants Recapitulate
Closing Intermediates
Analysis of the structures allows us to propose the following
sequence of atomic events (Figures 3 and 2B and Figure S6). To
name the various sites of the ligand and subsites of PDF, we will
use the usual nomenclature found in [55], which defines the
various binding pockets of a protease, where P1’ is the first side
chain at the C-terminal side of the cleavage site and its binding
pocket is S1’, also referred to as the hydrophobic pocket in the case
of PDF. First, actinonin aligns along the S1’ pocket to form the
encounter complex, which shifts the Ile130 side chain to avoid
steric hindrance in the S1’ pocket, promotes rotation of the Ile42
side chain, and finally rearranges the phenyl group of Phe58.
These
events
achieve an
optimal
hydrophobic
S1’
pocket
conformation (Figure 3), and the concomitant closure leads to
the formation of a hydrogen bond between the first carbonyl
group of actinonin and the backbone nitrogen of Ile42. The initial
N-O distance is reduced from 5 A˚ to 2.8 A˚ , which is an optimal
value for hydrogen bonding (Movie S2 and Figure S8B). Thus, the
primary driving force for the active site closure appears to be the
P1’:S1’ hydrophobic interaction. The C state is ultimately locked
by the super-b-sheet hydrogen bonds extending across the ligand,
including those involving Ile42. The DDGbinding value (2.2–
2.4 kcal/mol, Figure S8B), as calculated from the Kd values for
actinonin binding to wild-type (WT) and G41M and G41Q, is
consistent with the loss of a hydrogen bond that also contributes to
the conformational stability of the protein [56,57]. Thus, this bond
contributes to the major binding free energy difference between
the two complexes (3.1 kcal/mol; Figure S8B, Tables S1 and S3,
and [29]). Interestingly, the above DDGbinding values also correlate
with the DDGES values derived from the kcat/Km and kcat
measurements [19]. This dataset strongly correlates with the
Figure 5. Inhibition and enzymatic reactions progress through an induced fit pathway. (A) The catalytic parameters Km and kcat, for all
AtPDF variants are provided as a percentage of the wild-type values (WT). Detailed values are presented in Table S2. (B) Schematic model for
actinonin binding to AtPDF in favor of an induced-fit pathway. PDF might exist in at least two conformational states, open (O) or closed (C). The
relative abundance of each conformation would vary, depending on the enzyme type. With AtPDF, it is likely that the most abundant form is the O
one, which is the only form leading to a productive complex. The superclosed form (S) is likely to show reduced affinity for the ligand because of
steric occlusion of the active site. At the initial stage, the inhibitor (shown in red) binds to AtPDF (indicated in brown) in the O conformation. To reach
the final key-lock state (productive closed conformation, C), two major and extreme pathways can be used. According to the conformational selection
pathway, the inhibitor selects the C conformation. This pathway, which is represented by the dashed arrow, does not occur within the crystal. In
contrast, the G41Q and G41M mutants, by providing the structure of the enzyme in intermediate conformations (I), prove the existence of the so-
called encounter complex and confirm that the inhibitor binds to the enzyme when it is in the O conformation. The ligand-binding site is then
reorganized to yield the C enzyme conformation, that is, the key-lock state. Indeed, the inhibitor binds to the enzyme through the induced-fit
pathway. Each timescale was calculated using the data available in the text and corresponds to t1/2 values deduced from the calculation of 0.693/
(kinetic constant of interest). The kcat value (k2) was used to assess the timescale of catalysis in panel C, whereas, in (B), k4 assesses the first step of
inhibition, and k6 is used in the case of the slow step. For the SO conversion (left, B), the lifetime of the minor form of EcPDF was used to assess the
order of magnitude (see text and [38]). (C) Schematic model for the deformylation reaction catalyzed by PDF. Since actinonin is a pseudo-peptidic
inhibitor, it is likely that a peptidic substrate can bind to the PDF enzyme through an induced-fit pathway, as described in (B). The key-lock state
represents a transition state in which the N-formylated substrate is deformylated to yield the final reaction product.
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gyration and van der Waals radii of the side chain at position 41 as
well as the N-O distance between the first carbonyl group of
actinonin and the backbone nitrogen of Ile42 (Figure S8). These
results suggest that the capacity of both G41M and G41Q variants
to form the transition state is a consequence of their inability to
reach the fully closed state.
Thus, our study of the designed Gly41 mutant enzymes reveals
that, in addition to the initial and final states observed for the WT
enzyme, the conformations of the Gly41 variants correspond
indeed to on-pathway intermediates, thus providing snapshots
along the trajectory from the O to the C state of the enzyme
(Figures 2B and 3). The 3D structure of the variants in the absence
of ligand is similar to that of WT, and a strict correlation exists
between the completeness of the conformational change and both
binding potency and catalytic efficiency. This suggests that both
events require complete protein closure to generate a productive
complex.
The
strong
stabilization
of
AtPDF
by
actinonin
(Figure 1D) closely mimics what occurs with its natural substrates
when it reaches the transition state [34,58]. Indeed, as expected,
the enzyme facilitates the final C conformation by lowering its
final energy [6]. Optimal arrangement of the S1’ pocket (Figure 3)
proceeds
along
the
reaction
process
towards
the
final
C
conformation, triggering the alignment of reactive groups in an
optimal arrangement for ligand recognition. Upon binding,
actinonin alters the thermodynamic landscape for the structural
transition between the O and C states. This ligand is a potent
inhibitor because it can trigger the above sequence of events
similar to the substrate, but unlike the substrate, it is non-
hydrolyzable. Thus, by mimicking the transition state and being
non-hydrolyzable (Figure 1B), the final C complex is long lasting.
Ligand-Induced Conformational Closure Is Initially
Triggered by the Binding of the P1’ Group in the S1’
Pocket
Given the similarity between actinonin and natural substrate
binding, the very slow kinetics of inhibitor binding (10-s time-scale)
remains puzzling compared to the 10 ms required for catalysis
(deduced from the kcat). This finding could be explained as a
conformational effect during the formation of the hydrogen bond,
aligning the substrate as an additional beta-sheet and eventually
stabilizing the entire enzyme-ligand complex. The significantly
longer time needed to reach the most stable state compared to the
substrate would most likely be due to the presence of the flexible
and one carbon longer metal-binding group in actinonin (i.e.,
hydroxamate versus formyl, Figure 1B). This suggestion is in line
with the overall data obtained when we investigated more deeply
the role of the first carbonyl group of the ligand. This group is well
known
to
exert
a
crucial
effect
in
both
productive
and
unproductive ligand binding (i.e., substrate and inhibitor) [54].
In this respect, we studied the binding of compound 6b (Figure
S5B), a PDF ligand that does not exhibit a reactive group at this
position [49]. We observed that this compound binds strongly to
both EcPDF (KI* = 6366 nM) and AtPDF (KI* = 400635 nM)
but, unlike actinonin, does not display slow, tight binding as KI* =
KI. This impact on binding is consistent with the absence of the
hydrogen bond involving the first carbonyl group of the ligand.
The 3D structure of AtPDF was determined after soaking the
compound in crystals of the free, open AtPDF form. Upon binding,
6b induced a complete conformational change, identical to that
observed with actinonin (Figures 2B and 6A; ‘‘O’’ state). This
result further suggests that the conformational change is not
induced initially by the formation of this hydrogen bond and that
the encounter complex is primarily driven by the fit within the S1’
pocket. This also reveals that the timescale of the large
conformational change is several orders of magnitude faster than
the kinetics of slow binding and fully compatible with both the first
step of actinonin binding (k4 = 140 s21; see Table 1) and the
catalytic rate of the substrate (kcat = 37 s21; see Table 1 and Table
S3). The 3D structure also revealed that both the P1’ and the
hydroxamate groups are bound similarly to the corresponding
groups of actinonin (Figure 6B). As expected, no additional
bonding occurs, especially around the backbone nitrogen of Ile42
(Figure 6C).
Taken together, these data allow us to conclude that the
conformational change observed upon ligand binding is triggered
primarily by binding in the S1’ pocket. As revealed by the binding of
6b, the one carbon longer metal-binding group fits, immediately
upon recognition of the P1’ group, in the S1’ pocket and forms a
bidentate complex with the metal cation, mimicking the transition
state as a result. Thus, the active site is very confined and rigid due
to the presence and length of the hydroxamate group (compare
right and left panels in Figure 1B). As a result, compared to the
complex made with the substrate, it is likely that the formation of the
hydrogen bond involving the carbonyl of actinonin and the
backbone nitrogen of Ile42 becomes strongly rate-limiting (k5
= 0.044 s21; Table 1). Once this hydrogen link is locked, the
uncleavable bond, mimicking the labile formyl group at the
transition state, stabilizes the enzyme-inhibitor complex, making it
long-lasting (k6 = 0.0006 s21; Table 1) and providing a mechanistic
explanation for the slow-binding effect that involves both large and
fine conformational changes. The large conformational change is
similar to the one occurring with the substrate, whereas the second is
more subtle and locks the hydrogen bond involving the backbone
nitrogen of Ile42. The second step is rate-limiting with some
transition state analogs such as actinonin (Figure 5B and C).
Proper Positioning of the Carbonyl Group Is Required to
Stabilize the Complex at S1’
Compound 21 corresponds to another interesting derivative
designed to probe the impact of the peptide bond in PDF binding
[49]. In addition to the hydroxamate group, this compound
features both a hydrophobic benzyl group at P1’ and a reverse
peptide bond. Compound 21 shows modest but significant
inhibitory potency to AtPDF1B (KI* = 400637 nM), confirming
the crucial role of the peptide bond in PDF binding. After soaking
with crystals of apo-AtPDF, compound 21 could be detected in
high-resolution electron density maps (Figure S9A). Unlike 6b, 21
did not bind the active site of the enzyme but an alternative pocket
at the surface of the protein (Figure S9B). A docking study
performed with EcPDF had previously revealed this alternative
binding pocket (Figure S9C; [59]).
The aforementioned data indicate that the occurrence of a S1’-
binding group placed in the unfavorable context of a reverse
peptide bond does not stably promote binding at the active site of
AtPDF. Upon binding of 21, the 3D structure of both molecules of
the asymmetric unit remain in an O conformation (r.m.s.d.
,0.2 A˚ with respect to the apo-structures in the ‘‘O’’ state). This
finding suggests that only the binding of compounds entering the
S1’ pocket, such as actinonin or 6b, induces conformational
change, in keeping with the crucial role of the P1’ group if located
in the frame of a classic peptide bond. Moreover, we noticed that
the binding pocket of 21 was located on the rear side of the true
S1’ pocket and induced a weak modification of the P1’ hosting
platform (Figure S9D). Indeed, when crystals of the 21:AtPDF
complex were soaked in actinonin, the final 3D structure no longer
showed evidence of compound 21 occupancy greater than 5%.
Instead, this structure revealed both actinonin and closing of the
protein (Table S2). The r.m.s.d. between this structure and that
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obtained directly with actinonin was less than 0.2 A˚ ; the actinonin
position was virtually identical, indicating that the protein had
retained full capacity for binding actinonin and closing despite the
presence of compound 21. We conclude that actinonin does
compete with 21 because of the overlap at P1’ of AtPDF1B (Figure
S9C). As the actinonin S1’ subsite strongly mimics that of a true
substrate, this result also explains the inhibitory behavior of 21
towards AtPDF.
Discussion
Although PDF catalysis has been extensively studied and the
mechanism has been elucidated [34], how the enzyme achieves the
catalytically competent state remains unknown. Here, we provide
insight on how the enzyme might reach a catalytically competent
conformation, demonstrating that the reactive groups move into
proximity to promote catalysis (Figures 2B and 5C). We suggest
Figure 6. Effect of 6b binding on the conformation of key residues of PDF. Superimposition of free, 6b-, and actinonin-bound AtPDF
indicated in brown, red, and yellow, respectively. (A) Molecule A in the three models was superimposed, resulting in an r.m.s.d. of 0.9 A˚ for 100% of
the Ca. Actinonin is shown in yellow and 6b in red. (B) Conformation of key residues Ile42, Phe58, and Ile130 in the different complexes and in
unbound WT AtPDF. Actinonin is shown in yellow and 6b in red. (C) A detailed view of the AtPDF ligand-binding site for both actinonin and 6b
complexes, which are indicated by sticks and are superimposed. The two ligands are colored in pale and dark grey, respectively. The hydrogen bond
made by actinonin only is shown.
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that the motions of the catalytic centre starting with free ligand-
PDF favor a final configuration that is optimal for binding and/or
catalysis (Figures 1B, 2B, and 5B and C). In our model, we propose
that free PDF might exist in at least two conformational states, that
is, open (O) or super-closed (S). The relative abundance of each
conformation varies by enzyme type and incubation conditions,
explaining why both conformations have not been trapped thus
far. In the case of AtPDF, it is likely that the most abundant form
corresponds to an O state, which is the form that leads to a
productive complex. Indeed, in the NMR spectra for EcPDF, a
few residues show exchange cross-peaks from an additional,
alternative form [38]. The most strongly affected residues are
Cys90, one of the metal ligands, its neighbor Leu91, as well as
Ala47 and Ala48 on the facing strand. This suggests that EcPDF
exists in at least two conformations (‘‘S’’ and ‘‘C’’; see Figure 2B),
which undergo slow interconversion on the NMR timescale. The
3D structure of the major conformation (75%, lifetime 300 ms)
could be solved at high resolution, but the structure of the minor
form (25%, lifetime 100 ms), which exhibits very weak signals,
could
not
be
solved
[38].
This
conformation
appears
to
correspond to that of the complex obtained with the product of
the reaction (Met-Ala-Ser). A very similar situation—although
more balanced between the two states—appears to occur in the
case of variant G41M, suggesting that a mechanism involving
conformational selection followed by induced fit is a general model
for PDF and that AtPDF is a specific case where population shift
virtually does not occur as the free enzyme is completely in the O
conformation. This is also in line with data obtained with L.
interrogans PDF (LiPDF), which reveal conformers in both the S and
C states (see Figure 2B) and suggest a population-shift mechanism
[43]. It is interesting to note that LiPDF is a poorly active PDF
[60]. According to the representation shown in Figure 2B,
Plasmodium falciparum PDF (PfPDF), a poorly active PDF [61],
was retrieved only in the S state. Finally, weak decompaction of
the structure of Bacillus cereus and Staphylococcus aureus PDFs in the
presence of actinonin have been described [45,46]. These
examples suggest that the enzyme is trapped in the S conformer
in the free state and converts to the C conformer when bound to
actinonin, suggesting that the S conformer is overrepresented in
solution compared to the O state, unlike AtPDF.
This study of AtPDF—including 10 different crystal structures of
apo- and complexed enzyme variants—reveals the 3D structure of
a PDF in at least four distinct states. This includes the O form, the
occurrence of which is crucial for catalysis, as it is the active form.
Here, we propose that the transition from the O to the C state is
directly induced by the ligand. Indeed, the O form, which is
captured in the crystal, undergoes closure directly upon ligand
binding in our soaking experiments. Progression to this closure
involves intermediary states (‘‘I’’) similar to those observed with
variants G41Q and G41M in the presence of actinonin (see
Figure 2B). Extrapolating the situation to catalysis, which occurs in
the crystalline states of PDF, it is likely that hydrolysis of the
substrate frees the enzyme in its S state, which in turn needs to
open to accommodate a new substrate (Figures 2B and 5C). This is
well illustrated in the 3D structure of EcPDF complexed with a
product of the reaction, obtained after co-crystallization of the
enzyme with the substrate in a closed conformation [34]. The S
free form is likely to exhibit a slower on-rate for the ligand (k3)
compared to the O form because of steric occlusion of the active
site (Figure S10). In support of this hypothesis, recent data show
that the 3D structure of a C-terminally truncated, poorly active
version of AtPDF is in the C conformation in the unbound state,
although crystallized under conditions identical to ours [62,63].
This structure is similar to that of chain B, one of the two
molecules of the asymmetric subunit of variant G41M (Figure 2B).
This suggests that alterations remote from the active site
significantly unbalance the equilibrium between the two conform-
ers, thus altering the efficiency of the reaction (Figure 5C). As the S
version corresponds to a significantly less active version of AtPDF
compared to that reported in our present work, this further
confirms that, compared to the O state, the S state has a
significantly weaker propensity to bind substrate or a close mimic
ligand, such as actinonin. Comparison of the 3D structures of the
free-closed and the ligand-bound-closed forms reveals some
differences responsible for the slight steric reduction of the active
site of free-closed AtPDF1B with respect to that of the actinonin-
AtPDF1B complex (Figure S10A), including the side chain of Ile42
burying the S1’ binding pocket (Figure S10B). Overall, these data
suggest that an S form might exist under the free state but that it
would feature a k3 value with respect to the ligand that is
significantly weaker than that of the O form, which would strongly
slow down the reaction or the binding as a result.
With the interaction scheme proposed in our model (Figure 5B
and C), the ligand/substrate binds more easily to the O form and
induces the optimal conformation of the enzyme to reach the
transition state, thus allowing the reaction to be efficiently
catalyzed.
In
the
final
model
(Figure
5C),
there
is
both
conformational selection and induced fit subsequently involved
in line with the recently proposed existence of such mixed
mechanisms for other enzymes [15,16]. Nevertheless, in our model
(Figure 5C), we suggest that induced fit is the primary mechanism,
as it provides energy input from the ligand, which eventually drives
the enzyme towards the productive key-lock complex. Unambig-
uous distinction between the relative contributions of the two
mechanisms is deduced from the observation that kobs is a saturable
function of actinonin with various PDF, including EcPDF, BsPDF,
AtPDF (Figure 4B and C), and S. aureus PDF [12,16,29,49].
Using crystallographic reconstruction analysis involving enzyme
variants, motions of small mobile loops and movie reconstructions
of snapshots of catalytic events have been previously documented
[1–3,64–66], often by visualizing the binding of unnatural
inhibitors and not necessarily mimicking closely the substrate
and transition state as actinonin does [67,68]. However, only a few
examples make use of soaking conditions of a crystal to promote
the motion and show the importance of induced fit [1,69]. None of
these data show a motion of the amplitude revealed here with PDF
and a large stabilization of the complex involving the formation of
the four-stranded b-sheet superstructure and the entire N-domain
of the enzyme. Compared to previous crystallographic analyses,
our work integrates biophysical, computational, and kinetic
analyses to reconstruct the whole picture, allowing a better
understanding of the slow-binding mechanism.
While our work primarily focused on an induced-fit mechanism
of enzyme inhibition and catalysis, it should be emphasized that
this phenomenon is also applicable to the broader area of
receptor-ligand interactions. For example, in all cases where
conformational change mechanisms have been proposed for
kinase inhibitors without supporting experimental data [12,26],
further experimental work must be provided to clarify the precise
mechanism. We expect this will have important implications on
how one conducts future drug-discovery efforts against such
enzymes [70].
Materials and Methods
Protein Expression and Purification
Expression and purification of mature Arabidopsis thaliana PDF1B
and all variants (i.e., AtPDF) were derived from the previously
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May 2011 | Volume 9 | Issue 5 | e1001066
described protocol [37]: the lysis supernatant after sonication was
applied on a Q-Sepharose column (GE Healthcare; buffers A and
B as described containing 5 mM NiCl2) followed by Superdex-75
chromatography (GE Healthcare) using buffer C consisting of
buffer A supplemented with 0.1 M NaCl. For crystallization
experiments, the protein was purified further. The sample was
concentrated on an Amicon Ultra-15 centrifugal filter unit
(Millipore Corp.) with a 5-kDa cut-off and applied to a MonoQ
HR5/5 column (GE Healthcare) previously equilibrated in buffer
A (50 mM Hepes, pH 7.5, and 5 mM NiCl2). Elution was
performed with a 50-mL gradient from 0% to 100% buffer B.
The buffer of the pooled purified AtPDF1B was exchanged using a
PD-10 desalting column (GE Healthcare) to yield a protein
solution in 50 mM Hepes, pH 7.5, 0.1 M NaCl, and 5 mM NiCl2
(buffer C). The protein was concentrated on an Amicon Ultra-15
centrifugal filter unit. The resulting AtPDF1B preparation was
frozen in aliquots and stored at 280uC (for crystallization
purposes) or diluted 2-fold in 100% glycerol and stored at
220uC (for enzymatic purposes). The typical yield was 5–10 mg
AtPDF per liter of culture. All purification procedures were
performed at 4uC. Samples of the collected fractions were
analyzed by SDS-PAGE on 12% acrylamide gels, and protein
concentrations were estimated from the calculated extinction
coefficients for each variant.
Site-directed mutagenesis of AtPDF sequence in plasmid
pQdef1bDN [36] was carried out using the QuickChange Site-
Directed Mutagenesis Kit (Stratagene).
Enzymology
Assay of PDF activity was coupled to formate dehydrogenase,
where the absorbance of NADH at 340 nm was measured at 37uC
as previously described [71]. For measurements of classical kinetic
parameters (i.e., Km and kcat), the reaction was initiated by addition
of the substrate Fo-Met-Ala-Ser to the mixture containing purified
enzyme in the presence of 1 mM NiCl2. The kinetics parameters
were derived from iterative non-linear least square calculations
using the Michaelis-Menten equation based on the experimental
data (Sigma-Plot; Kinetics module). For determination of kinetic
parameters related to actinonin, the reaction mixture contained
750 mM NiCl2. In some cases, the mixture containing PDF and
actinonin was incubated for 15 min at 37uC before kinetic
analysis, which was initiated by the addition of substrate. The
same protocol was used to determine the dissociation constant of
actinonin [KI* = k4/(k3+k3k5/k6)], but the initial reaction velocities
were measured with varying concentrations of Fo-Met-Ala-Ser and
actinonin. The data were then calculated according to the method
of Henderson, which can be used to determine the dissociation
constant
of
the tight-binding
competitive enzyme
inhibitor
[28,49,72] by varying both the inhibitor and substrate concentra-
tions. To determine KI, k5, and k6, the reaction was initiated by the
addition of enzyme as previously described [29,49]. KI*app
measurements were used for comparative studies of AtPDF
variants (Table S3) at a concentration of 2 mM substrate by
varying the concentration of actinonin. KI*app is the slope of the
v[Actinonin]/v0 line curve. kobs was fitted from the kinetic data without
preincubation with vI = vs + (v0 2 vs)e2kobst where vI is the observed
velocity at a given concentration of inhibitor I, v0 is the velocity,
and vs is the steady-state velocity [18]. From the set of values
obtained at various concentrations of I, k5 and k6 could be derived
using kobs = k6 + k5[I]/(KI + [I]). By choosing a set of values with
kobs..k6, 1/kobs
= 1/k5(KI/[I] +1) and 1/kobs
=
f(1/[I]) is
expected to be a straight line in case of induced fit whose positive
slope corresponds to 1/k5. k6 was derived from equation k6 = k5/
(KI/KI*21) [18,19].
Microcalorimetry
ITC experiments were performed using a VP-ITC isothermal
titration calorimeter (Microcal Corp.). Experiments were per-
formed at 37uC. For each experiment, injections of 10 mL
actinonin (180 mM) were added using a computer-controlled
300 mL microsyringe at intervals of 240 s into the Ni-AtPDF
variant solution (5 to 10 mM, cell volume = 2.1 mL) dissolved in
buffer C with stirring at 310 rpm. A theoretical titration curve was
fitted to the experimental data using the ORIGIN software
(Microcal). This software uses the relationship between the heat
generated after each injection and DHu (enthalpy change in kcal/
mol), KA (the association binding constant in M21), n (number of
binding sites per monomer), total protein concentration, and free
and total ligand concentrations. The thermal stability of the WT
and variants of Ni-AtPDF1B was studied by DSC using VP-DSC
calorimetry (Microcal Corp.). DSC measurements were made with
10 mM protein solutions in buffer C. The actinonin concentration
was 20 mM. The same buffer was used as a reference. All solutions
were degassed just before loading into the calorimeter. Scanning
was performed at 1uC/min. The temperature dependence of the
partial molar capacity (Cp) was expressed in kcal/K after
subtracting the buffer signal using Origin(R) software.
Crystallization and Soaking Experiments
Crystallization conditions were screened by a robot using the
sitting drop vapor diffusion method. Crystals were obtained and
optimized at 20uC with 15%–20% PEG-3350 and either 0.1 or
0.2 M zinc acetate. The drops were formed by mixing 2 mL of a
solution containing 2 to 4 mg/mL protein and 2 mL of the
crystallization solution. Crystals were soaked for 24 h by adding
actinonin to the crystallization drops at a final concentration of
5 mM. Cryoprotection was achieved by placing crystals for 30 s in
a solution that was composed of 20% PEG-3350 and 0.2 M zinc
acetate, supplemented with 5%, 10%, and 15% glycerol. Crystals
were then directly flash frozen in liquid nitrogen using cryoloops
(Hampton Research). Crystals were also grown under conditions
described for the C-terminally deleted, weakly active version of
AtPDF [63].
X-Ray Diffraction Data Collection
Data collections were performed at 100 K at the European
Synchrotron Radiation Facility (Grenoble, France) on station
ID29, FIP-BM30A, ID14-1, and ID23-2, and at SOLEIL (Gif-sur-
Yvette, France) on station PROXIMA1. In each case, a single
crystal was used to collect a complete dataset. Data were processed
and scaled using XDS software [73]. Two crystal forms were
encountered with different cell parameters. In each case, b
parameter was nearly equal to a, and data could be indexed into
two space groups, P212121 or P43212. The data are shown in Table
S2.
Structure Determination and Refinement
The
structure
of
free
AtPDF
was
solved
by
molecular
replacement with Phaser [74] followed by a rigid-body refinement
by CNS [75] using coordinates from the Plasmodium falciparum PDF
(PDB code 1RL4) [76] as a search model. The structures of
actinonin-bound proteins—that is, WT and mutants—were solved
using rigid-body refinement by CNS of the free AtPDF structure.
The ten final models were obtained by manual rebuilding using
TURBO-FRODO [77] and combined with refinement of only
calculated phases using CNS and Refmac [78] software. No non-
crystallographic symmetries were used. Quality control of the
three models was performed using the PROCHECK program
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13
May 2011 | Volume 9 | Issue 5 | e1001066
[79]. To probe for alternative conformers, Ringer was used [53].
Ringer is a program to detect molecular motions by automatic X-
ray electron density sampling, and can be accessed at http://
ucxray.berkeley.edu/ringer.htm.
Accession Numbers
PDB codes for the PDF structures presented within this
manuscript are as follows: 3M6O, 3PN2, 3M6P, 3O3J, 3PN3,
3PN4, 3PN5, 3M6Q, 3PN6, and 3M6R. UniProtKB accession
numbers for other PDF studied are P0A6K3 (EcPDF) and O31410
(BsPDF).
Supporting Information
Figure S1
Alignment of PDF sequences and secondary structures.
(A) PDF1B from Arabidopsis thaliana (AtPDF1B) is compared with
bacterial type 1B (EcPDF and LiPDF), pathogenic protozoa
(PfPDF1B), eukaryotic mitochondrial PDF1A from A. thaliana
(AtPDF1A), and bacterial type 2 (BsPDF2). This figure was created
with ENDscript [80]. The sequence alignment was realized with the
algorithm muscle included in ENDscript, and modified according to
the superimposition of structures. The blue frames indicate
conserved residues, white characters in red boxes indicate strict
identity, and red characters in yellow boxes indicate homology. The
secondary structures at the top (a-helices, 310 helices, b-strands, and
b-turns are shown by medium squiggles, small squiggles, arrows,
and TT letters, respectively) were predicted by DSSP [81]. Relative
accessibility (acc) of subunit A is shown by a blue-colored bar below
sequence. White is buried, cyan is intermediate, and blue with red
borders is highly exposed. A red box means that relative accessibility
is not calculated for the residue, because it is truncated. Hydropathy
(hyd) is calculated from the sequence according to [82]. It is shown
by a second bar below accessibility: pink is hydrophobic, grey is
intermediate, and cyan is hydrophilic. Motifs 1 (41GwGwAAXQ48),
2 (89EGCLS93), and 3 (133HEwDH137), where w is a hydrophobic
amino acid, are labeled by red stars below the sequence alignment.
To simplify the nomenclature, AtPDF1B is referred to as AtPDF
throughout the text. (B) Topology cartoon of AtPDF, free (left) or
actinonin bound (right), in the same color code as (A). Actinonin
(represented by the yellow arrow) binding to the ligand binding site
allows the linkage of the two distinct b-sheets into one single b-sheet,
by mimicking an additional b-strand. PDB sum (http://www.ebi.ac.
uk/thornton-srv/databases/pdbsum/) was used. (C) 3-D structure
of AtPDF is represented showing the position of the residues
discussed in the text, indicated in red.
(EPS)
Figure S2
Microcalorimetric titration of AtPDF with actinonin.
Data were obtained at 37uC by an automated sequence of 28
injections of 180 mM actinonin from a 300 ml syringe into the
reaction cell, which contain 9.85 mM AtPDF. The volume of each
reaction was 10 ml, and injections were made at 240 s intervals.
Top, raw data from the titration. Each peak corresponds to the
injection. Bottom, the peaks in the upper panel were integrated
with ORIGIN software and the values were plotted versus
injection number. Each point corresponds to the heat in mcal
generated by the reaction upon each injection. The solid line is the
curve fit to the data by the Origin program. This fit yields values
for Kd. Experiments were done with wild type protein and others
variants, and gave similar raw data and curve fit. (A) WT; (B)
variant G41M; (C) variant I42W.
(EPS)
Figure S3
Binding of actinonin to AtPDF does barely modify the
crystal packing. (A) Crystal pack of the two complexes: open, free
complex (left) and bound to actinonin (right) (B). Non-crystallo-
graphic contacts into asymmetric unit are not modified by closing
movement of the protein due to actinonin binding, except for zinc
atom number 6. This metal ion is coordinated by side chains of
Asp40 and Glu63, and water molecules, Asp40 and Glu63 being
hydrogen bonded by side chain of Lys38 of the other subunit of
the asymmetric unit. With the closing movement of the protein
into the crystal, Ca of Asp40 shifted by 3.1 A˚ and its side chain
flipped by 90u. Therefore, it does no longer participate to the
coordination shell of this Zn2+ ion. However, it is still hydrogen
bonded by Lys38 from chain B.
(EPS)
Figure S4
Binding of actinonin to AtPDF closely mimics both
actinonin and product binding to EcPDF. Superimposition of
EcPDF and AtPDF bound to either actinonin (1LRU PDB code,
panel A) or Met-Ala-Ser (1BS6 PDB code, panel B), the product of
the
reaction.
The
r.m.s.d.
value
is
1.11
A˚
for
151
Ca
superimposed.
(EPS)
Figure S5
The ligand binding site of AtPDF. This picture shows
the residues of AtPDF that are in contact with actinonin (left) and
6b (right) according to the 3-D structure; this should be compared
to the similar scheme shown in Figure 1B for EcPDF.
(EPS)
Figure S6
Electronic densities of the moving side-chains and of
actinonin at the binding site in some variants of AtPDF. Actinonin
and selected residues (G/Q/M41, I42, F58, and I130) are drawn
in stick and are shown in their FO–FC electron density omit maps
contoured at 2s, in free wild-type AtPDF (two crystallization
conditions, WT1 and WT2), and ligand-bound WT (actinonin, 6b
and 21), G41Q, and G41M variants.
(EPS)
Figure S7
Only few residues show alternative conformation in
AtPDF. Alternative conformers in the crystalline state of AtPDF.
Ringer
plots
of
electron
density
(r)
versus
x1
angle
for
representative residues of the 3-D apostructure of AtPDF. Data
were obtained with the 3M6O dataset (see Table S1). The
secondary peaks in the Ile residues are observed because Ile is a
branched amino acid. To evidence an alternative conformation
with Ile, three peaks should be observed.
(EPS)
Figure S8
Impact of induced fit on the binding free energy of
actinonin depends on the capacity to stabilize a hydrogen bond
with PDF. (A) The gyration radii [83] of the side chain occurring
at position 41 is displayed with black squares and compared to the
kcat/Km values (grey bars). (B) The distance between the NH of I42
and the CO of actinonin was measured in each case. The
percentage of the distance required to make a hydrogen bond (2.8
A˚ ) is reported (dark squares). The difference of binding free energy
(DDGbinding) between the open, free state and the variants closed
complexes of the G41 variants are displayed as grey bars. The
values were calculated as follows. For the WT, it corresponds to
the RT ln(KI*/KI) value [29], where R is the ideal gas constant and
T is the temperature in Kelvin. RT is 0.616 kcal.mol21 at 37uC.
For the G41M and G41Q variants, the DDGbinding corresponds to
RT ln(KI-G41variant/KD-WT). The obtained values are similar to that
obtained if the kcat/Km substitutes the KD value in the calculation
(DDGbinding = RT ln(kcat/Km –G41variant/kcat/Km –WT).
(EPS)
Figure S9
Compound 21 does not bind AtPDF1B at S1’. (A) 21
is shown in ball-and-stick format in its FO–FC electron density omit
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May 2011 | Volume 9 | Issue 5 | e1001066
map contoured at 2s. (B) Binding site of 21 into AtPDF1B is
detailed. Red and blue residues indicate residues that accommo-
date
the
‘‘phenylalanine’’
and
‘‘trimethyl’’
groups
of
21,
respectively. (C) Overall view of 21 binding site (left). Molecular
surface of AtPDF is represented, as well as 21 in ball-and-stick
format. Residues belonging to the 21 binding pocket are colored in
orange. For comparison, molecular surface of EcPDF (PDB code
1G2A) in the same orientation is also represented, with residues
forming the new ligand binding pocket colored in orange.
Actinonin is represented in ball-and-stick format and is seen
through the molecular surface of each PDF. (D) Ball-and-stick
representation of the interaction network around compound 21.
The metal cation is shown as a grey sphere.
(EPS)
Figure S10
Poorly active versions of AtPDF are in a closed
conformation incompatible with actinonin binding. (A) Free and
close AtPDF were superimposed as in Figure 1C and are figured in
brown and yellow, respectively. Both the G41M (chain B, shown
in orange) and the free C-deleted weakly active AtPDF versions
([63], colored in purple, PDB entry code 3CPM) were superim-
posed, to the two structures, showing that they both fit better to the
ligand-bound full-length close form than to the free open form, but
that the closure is further pronounced, burying the entrance to a
ligand. (B) Close-up showing that the shape of the S1’ pocket of the
poorly active closed versions make it poorly available to P1’
recognition (see circled Ile142 and Ile130 side chains).
(EPS)
Table S1
Catalytic properties of AtPDF. Nm, not measurable;
ND, not determined; WT, is wild-type. aKinetic constants were
determined using the coupled assay as indicated in Materials and
Methods with substrate Fo-Met-Ala-Ser, in the presence of 100
nM enzyme variant and 750 mM NiCl2, at 37uC. The relative
value of kcat/Km for wild-type AtPDF was set at 100%. bData
correspond to the binding constant of actinonin as obtained either
from ITC or from enzymatic analysis when indicated with an
asterisk. cData from Table S3. dGyration radii are from [83].
(DOC)
Table S2
Crystallographic data and refinement statistics. Values
in parentheses are for the outer resolution shell. aRsym (I) =
ShklSi|Ihkl,i 2 ,Ihkl.|/ShklSi|Ihkl,i|, where ,Ihkl. is the mean
intensity of the multiple Ihkl,i observations for symmetry-related
reflections. bRwork = 1006(Shkl|Fobs 2 Fcalc|/Shkl|Fobs|). Rfree
is a test set including ,5% of the data. cPercentage of residues in
most-favored/additionally
allowed/generously
allowed/disal-
lowed regions of the Ramachandran plot. dCompound 21 was
added first, and actinonin afterwards.
(DOC)
Table S3
Kinetic parameters for inhibition of some AtPDF
variants by actinonin. The enzyme concentration used in the assay
was 100 nM. Prior to kinetic analysis for determination of KI*app
values, actinonin was incubated in the presence of each variant set
at the final concentration for 10 min at 37uC; kinetic assay was
started
by
adding
a
small
volume
of
the
substrate.
For
determination of KI, k5, and k6 values, actinonin was not pre-
incubated with enzyme and kinetic assay was started by adding the
enzyme.
(DOCX)
Movie S1
Dynamics of actinonin binding to peptide deformylase
and closure of the active site.
(WMV)
Movie S2
Progressive motions of the main side chains at the
active site and final locking of the hydrogen bond.
(WMV)
Acknowledgments
We are strongly indebted to James Fraser and Tom Alber (University of
California, Berkeley, USA) for introducing us to Ringer before the release
of the freely available downloadable version. We thank Benoıˆt Gigant,
Virginie Gueguen-Chaignon, Solange Morera, and Philippe Peynot
(CNRS, Gif-sur-Yvette, France) for help with data processing and access
to the crystallization facilities. We also thank Magali Nicaise-Aumont
(IBBMC, Orsay, France), who performed the microcalorimetry experi-
ments. We are grateful to the staff of the European Synchrotron Radiation
Facility (ESRF) and SOLEIL beamlines for their help during data
collection.
Author Contributions
The author(s) have made the following declarations about their
contributions: Conceived and designed the experiments: SF CG TM.
Performed the experiments: AB SF. Analyzed the data: FD MD SF CG
TM. Contributed reagents/materials/analysis tools: IA MD CG TM.
Wrote the paper: CG TM.
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|
3M6P
|
Crystal structure of Arabidopsis thaliana peptide deformylase 1B (AtPDF1B) in complex with actinonin
|
Trapping Conformational States Along Ligand-Binding
Dynamics of Peptide Deformylase: The Impact of
Induced Fit on Enzyme Catalysis
Sonia Fieulaine1, Adrien Boularot1, Isabelle Artaud2,3, Michel Desmadril4,5, Fre´de´ric Dardel6,7, Thierry
Meinnel1*, Carmela Giglione1*
1 CNRS, ISV, UPR2355, Gif-sur-Yvette, France, 2 Universite´ Paris Descartes, UMR8601, Paris, France, 3 CNRS, UMR8601, Paris, France, 4 Universite´ Paris-Sud, IBBMC,
UMR8619, Orsay, France, 5 CNRS, IBBMC, UMR8619, Orsay, France, 6 Universite´ Paris Descartes, UMR8015, Paris, France, 7 CNRS, UMR8015, Paris, France
Abstract
For several decades, molecular recognition has been considered one of the most fundamental processes in biochemistry.
For enzymes, substrate binding is often coupled to conformational changes that alter the local environment of the active
site to align the reactive groups for efficient catalysis and to reach the transition state. Adaptive substrate recognition is a
well-known concept; however, it has been poorly characterized at a structural level because of its dynamic nature. Here, we
provide a detailed mechanism for an induced-fit process at atomic resolution. We take advantage of a slow, tight binding
inhibitor-enzyme system, actinonin-peptide deformylase. Crystal structures of the initial open state and final closed state
were solved, as well as those of several intermediate mimics captured during the process. Ligand-induced reshaping of a
hydrophobic pocket drives closure of the active site, which is finally ‘‘zipped up’’ by additional binding interactions.
Together with biochemical analyses, these data allow a coherent reconstruction of the sequence of events leading from the
encounter complex to the key-lock binding state of the enzyme. A ‘‘movie’’ that reconstructs this entire process can be
further extrapolated to catalysis.
Citation: Fieulaine S, Boularot A, Artaud I, Desmadril M, Dardel F, et al. (2011) Trapping Conformational States Along Ligand-Binding Dynamics of Peptide
Deformylase: The Impact of Induced Fit on Enzyme Catalysis. PLoS Biol 9(5): e1001066. doi:10.1371/journal.pbio.1001066
Academic Editor: Gregory A. Petsko, Brandeis University, United States of America
Received January 7, 2011; Accepted April 14, 2011; Published May 24, 2011
Copyright: 2011 Fieulaine et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by the Centre National de la Recherche Scientifique (CNRS, France), grant ANR-06-MIME-010-01 (Agence Nationale de la
Recherche, France), and grant #4920 from the Association pour la Recherche sur le Cancer (Villejuif, France). SF was partly supported by a postdoctoral fellowship
from the Fondation pour la Recherche Me´dicale (France). AB was supported by CNRS, France. The funders had no role in study design, data collection and
analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
Abbreviations: DSC, differential scanning calorimetry; Fo, formyl; PDF, peptide deformylase; r.m.s.d., root mean square deviation.
* E-mail: Thierry.Meinnel@isv.cnrs-gif.fr (TM); Carmela.Giglione@isv.cnrs-gif.fr (CG)
Introduction
Flexibility of proteins around their active site is a central feature
of molecular biochemistry [1–5]. Although this has been a central
concept
in
biochemistry
for
half
a
century,
the
detailed
mechanisms describing how the active enzyme conformation is
achieved have remained largely elusive, as a consequence of their
transient
nature.
Direct
structural
evidence
and/or
kinetic
analyses have only recently emerged [6–10]. Three classic
‘‘textbook’’ models are used to describe the formation of the
ligand-enzyme complex: (i) the Fischer’s ‘‘lock-and key’’ model, (ii)
the Koshland’s induced-fit model, and (iii) the selected-shift model
or conformational selection mechanism [6–8,11–13]. In the
Fischer’s ‘‘lock-and key’’ model, the conformations of free and
ligand-bound proteins are essentially the same. In the induced-fit
model, ligand binding induces a conformational change in the
protein, leading to the precise orientation of the catalytic groups
and implying the existence of initial molecular matches that
provide sufficient affinity prior to conformational adaptation [14].
In contrast, the selected-fit model assumes an equilibrium between
multiple conformational states, in which the ligand is able to select
and stabilize a complementary protein conformation. In this case,
the conformational change precedes ligand binding, in contrast to
the induced-fit model in which binding occurs first. The
conformational selection and/or induced-fit processes have been
shown to be involved in a number of enzymes [12,13,15,16]. For
several of these studies, conformational selection is proposed
because the experimental data support that, even in the absence of
the ligand, the enzyme samples multiple conformational states,
including the ligand-bound (active) state [6]. Although direct
structural evidence and/or kinetic analyses have provided clues
[6–8,12,13,16], how we can distinguish whether a protein binds its
ligand in an induced- or selected-fit mechanism remains critical
and often controversial.
The enzyme-inhibitor interaction is a form of molecular
recognition that is more amenable to investigation than the
enzyme-substrate interaction as there is no chemical transforma-
tion of the ligand during this process. In this context, slow, tight-
binding inhibition is an interesting interaction process, as it closely
mimics the substrate recognition process and has been shown to be
commonly involved in adaptive conformational changes [12,
17,18]. In slow, tight-binding inhibition, the degree of inhibition at
a fixed concentration of compound varies over time, leading to a
curvature of the reaction progress curve over time during which
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the uninhibited reaction progress curve is linear [19]. Indeed, the
slow, tight-binding inhibition is a two-step mechanism that
depends on the rate and strength of inhibitor interactions with
the enzyme. Binding of the inhibitor (I) to the enzyme (E) leads to
the rapid formation of a non-covalent enzyme-inhibitor complex
(E:I) followed by monomolecular slower step (k5) in which the E:I is
transformed into a more stable complex (E:I*) that relaxes and
dissociates at a very slow rate, mainly inferred by the k6 value
when k6,,k5,,k4, (Figure 1A; see also footnote f in Table 1).
Although only a few studies have investigated the mechanisms
of slow, tight-binding inhibitors, such molecules are favored for use
as
therapeutics,
as
they
usually
exhibit
unique
inhibitory
properties, including selective potency and long-lasting effects
[20–26]. Here, we explore the precise structural inhibitory
mechanism of actinonin (Figure 1A; [27]), which is a slow, tight-
binding inhibitor of peptide deformylase (PDF), a metal cation-
dependent enzyme [28,29]. The function of the active-site metal is
to activate the reactive water molecule involved in peptide
hydrolysis [30]. PDF is the first enzyme in the N-terminal
methionine excision pathway, an essential and ubiquitous process
that contributes to the diversity of N-terminal amino acids [31,32].
Actinonin is a natural product with antibiotic activity that inhibits
PDF by mimicking the structure of its natural substrates (nascent
peptide chains starting with Fo-Met-Aaa, where Fo is a formyl
group and Aaa is any amino acid) in their transition state
(Figure 1B). The transition state inhibitor actinonin, as well as
other structurally related inhibitors, has been shown to systemat-
ically exhibit a ‘‘slow-binding’’ inhibition behavior (Figure 1A),
regardless of the organism of origin of the PDF [29,33].
Using structural, biocomputing, and enzymatic analyses, we
were able to (i) reveal that the free enzyme is in an open
conformation and that actinonin induces transition of the enzyme
into a closed conformation; (ii) show that there is no evidence for
the occurrence of a closed conformation in the apostructure of the
open enzyme, which, together with detailed kinetic analyses,
makes the closed form fully compatible with an induced-fit model;
and (iii) identify the sequence of molecular events leading to the
final, bound, closed complex (E:I*). Moreover, using several
rationally designed point mutants of the enzyme, ligand-induced
intermediates, which mimic conformational states that normally
would not be expected to accumulate with the wild-type (WT)
enzyme, were trapped. These conformations recapitulate physical
states that the WT enzyme must pass through during its overall
transition from the apo-enzyme to the E:I* complex. ‘‘Freezing’’ of
ligand-induced intermediate states provides direct evidence for an
induced-fit mechanism and allows the reconstruction of a virtual
‘‘movie’’ that recapitulates this mechanism. Since PDF is one
example of an enzyme remaining active in the crystalline state and
because actinonin closely mimics the natural substrates bound to
PDF in the transition state as shown previously with the Escherichia
coli form (EcPDF; see Figure 1B) [34,35], we propose a model
suggesting that induced fit also contributes to efficient catalysis.
Results
Slow, Tight Binding of the Transition-State Analog
Actinonin to Peptide Deformylase
In the present study, at the atomic level we explored the precise
inhibitory mechanism of actinonin on Arabidopsis thaliana PDF1B
(AtPDF), a close eukaryotic homologue of EcPDF (Figure S1)
[36,37]. Measurements of the kinetic parameters of the second step
of the binding mechanism (k5) revealed a timescale in the 10-s range
(Table 1), which is consistent with the collective motion of a large
domain [4,5]. This finding is supported by NMR studies [38,39],
which showed that actinonin binding induces drastic changes in the
heteronuclear single quantum coherence (HSQC) spectrum of
EcPDF, since most resonances undergo significant shifts that affect a
large part of the structure [40,41]. The existence of alternative
conformational states of EcPDF is further supported by recent
biophysical studies [42]. Previously reported snapshots of a series of
different conformations of the enlarged and mobile loop—the so-
called CD loop—of the dimeric PDF from Leptospira interrogans PDF
(LiPDF) in the presence or absence of inhibitor led to the hypothesis
of the existence of an equilibrium between a closed and open form
of the CD-loop of PDF enzymes, suggesting a selected-shift model to
the authors [43]. Taken together, these data suggest that the binding
of actinonin to PDF is accompanied or preceded by conformational
changes within the enzyme. Paradoxically, this proposal has not
been currently supported by the available structural data. Indeed,
free and complexed crystal structures have provided no evidence for
any significant conformational change in PDF structure induced by
the binding of ligand [35,43–47].
Tight inhibition in the closed state is associated with the KI*
apparent equilibrium constant (Figure 1A). A KI* value (see Table 1
and Materials and Methods for the biochemical definition of KI*)
of 0.9 nM for actinonin could be measured for AtPDF; that is, a
value very similar to that obtained for bacterial PDFs, including
EcPDF and Bacillus stearothermophilus PDF2 (BsPDF2, Table 1).
Tightening of the initial encounter complex (E:I) resulted in a final
complex (E:I*) in which the potency of actinonin (KI/KI*) was
enhanced by more than two orders of magnitude and exhibited a
very slow off-rate (k6, Table 1). The dissociation constant value of
AtPDF for actinonin was also assessed using isothermal titration
calorimetry (ITC) experiments (Table S1 and Figure S2A). The
corresponding ITC titration curves (Figure S2A) are consistent
with a very strong affinity of the ligand for the enzyme [48],
enabling us to determine an accurate Kd. Moreover, these studies
generated values similar to those measured by other means for
AtPDF and EcPDF [42,49].
Author Summary
The notion of induced fit when a protein binds its ligand—
like a glove adapting to the shape of a hand—is a central
concept of structural biochemistry introduced over 50
years ago. A detailed molecular demonstration of this
phenomenon has eluded biochemists, however, largely
due to the difficulty of capturing the steps of this very
transient process: the ‘‘conformational change.’’ In this
study, we were able to see this process by using X-ray
diffraction to determine more than 10 distinct structures
adopted by a single enzyme when it binds a ligand. To do
this, we took advantage of the ‘‘slow, tight-binding’’ of a
potent inhibitor to its specific target enzyme to trap
intermediates in the binding process, which allowed us to
monitor the action of an enzyme in real-time at atomic
resolution. We showed the kinetics of the conformational
change from an initial open state, including the encounter
complex, to the final closed state of the enzyme. From
these
data
and
other
biochemical
and
biophysical
analyses, we make a coherent causal reconstruction of
the sequence of events leading to inhibition of the
enzyme’s activity. We also generated a movie that
reconstructs the sequence of events during the encounter.
Our data provide new insights into how enzymes achieve a
catalytically competent conformation in which the reactive
groups are brought into close proximity, resulting in
catalysis.
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Ligand-Induced Conformational Closure of AtPDF in the
Crystalline State
Occurrence of a conformational change induced by drug binding
was visualized via the resolution of several crystal structure forms of
AtPDF, the free form and/or in a complex with actinonin (Table
S2). The data reveal a structural switch between the two forms that
can account for both the thermodynamic and kinetic data. The
enzyme was observed in two states, a novel open apo-form and a
closed, induced, actinonin-bound complex (Figure 1C). Binding of
actinonin resulted in a tightening of the active site through the
collective closure of the entire N-terminal portion of the protein
(strands b1, b2, and b3; helix a1; and CD-loop, see Movies S1 and
S2, Figure 1C, and Figure S1). The amplitude of the structural
change was maximal for Pro60 (Figure S1), the Ca of which was
shifted 4 A˚ upon actinonin binding. This collective movement
involved the formation of a ‘‘super b-sheet’’ as the result of the large
rearrangement of b-strands 4 and 5 relative to the rest of the
structure in which actinonin forms an additional strand bridging the
two b-sheets (b1 andb2) on either side of the active site (Figure 1D
and Figure S1B). As actinonin is a peptide-like compound (see
Introduction and Figure 1B), this behavior closely mimics what
occurs in the natural protein substrates of PDF, which also form this
strand-bridging interaction. This phenomenon also accounts for the
strong stabilization of the protein by actinonin, which was also
challenged by differential scanning calorimetry (DSC) experiments:
the Tm of AtPDF increased from 61uC to 81uC upon binding of the
inhibitor (Figure 1D, see also below).
Thus far, this closure of the enzyme induced by actinonin is part
of the rare structural evidence for the slow, tight-binding
mechanism at an atomic scale. The open state, which has never
been observed, was captured not only in the two molecules of the
asymmetric subunit but also in different crystals and under two
distinct crystallization conditions (Table S2 and Figure 2). All
r.m.s.d. values were smaller than 0.25 A˚ . The closure is very
unlikely to result from crystal packing constraints, as soaking the
apo-AtPDF crystals in a solution containing actinonin induced the
Figure 1. Slow, tight-binding inhibition of PDF by actinonin induces conformational change in the protein. (A) Inhibition by a two-step
mechanism, involving a tightening of the initial enzyme-inhibitor complex (E?I) to form a more stable complex (E?I*), with the chemical structure of
actinonin (I), the natural inhibitor of PDF enzymes (E). (B) Structures of EcPDF bound to actinonin (left) and to the transition state resulting from the
cleavage of its substrate, Fo-Met-Ala-Ser (right) [34,35]. (C) Superimposition of free and actinonin-bound AtPDF indicated in green and purple,
respectively. The three conserved motifs of the PDF enzymes family are indicated in orange and numbered I, II, and III. Molecules A of both models
were superimposed, resulting in an r.m.s.d. of 0.9 A˚ for 100% of the Ca. Left inset, close-up comparison of the open and closed forms figured in the
ribbon representation. (D) Baseline-corrected DSC thermograms of free and actinonin-bound WT AtPDF recorded under the same experimental
conditions.
doi:10.1371/journal.pbio.1001066.g001
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structural transition from the open to the closed state within the
crystals without cracking them or altering their diffracting power.
Thus, crystal packing is compatible with both states of the enzyme
(Figure S3). Therefore, the open structure most likely corresponds
to a stable state in solution.
The closed final conformation was identical to that previously
reported for PDF complexes obtained either with actinonin or with a
product of the reaction [34,35,44,50], indicating that this structure is
common for the ligands (compare Figures 1B and 2A, and Figure S4).
Hydrogen bonding was also conserved, especially the bond between
the backbone nitrogen of Ile42 (corresponding to Ile44 in EcPDF, see
Figure 1B and Figure S5A) and the alkyl carbonyl chain of actinonin,
which potently contributes to the formation of the super b-sheet
(Movie S2 and Figure S1B, see also below). Between the open and
closed states, the side chains of Ile42, Phe58, and Ile130 underwent
significant structural changes (Figure 3A and D and Figure S6),
corresponding to a hydophobic pocket rearrangement, with Ile42
being the most affected (Figure 3). Interestingly, Ile42 is the second
residue of the conserved active-site motif G41IGLAAXG (motif 1)
that was previously shown to be essential for activity [51].
To assess and visualize the differences between the two states, two
independent structural parameters were measured: the r.m.s.d.
value with respect to the open form and the aperture angle (dap),
which measures the angle made between the N- and C-domains
through three fixed-points, corresponding to the Ca of three
conserved residues, each sitting in one of the three conserved motifs
(Figure 2A). The bi-dimensional graph of these two parameters is a
good representation of the closing motion snapshots (Figure 2B)
shown in Movie S1. With this tool at this stage, two states could be
defined: the closed (C) and open (O) states (Figure 2B).
Evidence for a Pure Induced-Fit Mechanism in the
Binding of Actinonin to AtPDF
Recent quantitative analyses of both conformational selection
and induced fit have led to an integrated continuum—a so-called
‘‘flux-description’’—of
these
two
limiting
mechanisms
[16].
According to this model, conformation selection tends to be
preferred at low ligand concentrations (mM range)—that is, using
detailed kinetic studies—whereas induced fit dominates at high
ligand and enzyme concentrations (mM range) obtained, for
instance, in NMR or crystallographic approaches. Structural
studies are most useful to reveal subpopulations of biological
significance.
We investigated the existence of lowly populated, alternative
conformations of apoPDF. To probe the occurrence of alternate
conformers in the crystalline state of PDF, the new Ringer
program is the most suitable investigation tool [52,53]. Ringer
searches for evidence of alternate rotamers by systematically
sampling electron density maps—free of model bias—around the
dihedral angles of protein side chains. Two independent WT open
datasets of the apoenzyme, including a high-resolution set (1.3 A˚ ),
were used in the analysis. Ringer analysis revealed the existence of
only one rotamer of most side chains of either molecule in the
asymmetric unit, including the three main residues primarily
involved in conformation change—that is, Ile 42, Phe58, and
Ile130
(Figure
4A).
Ringer
analysis
showed
evidence
for
unmodeled alternate conformers for very few residues, including
Ile121 and Phe87, or Phe119 to a much lesser extent (Figure S7).
There is therefore no evidence for the occurrence of a closed
conformation in the apostructure of AtPDF, supporting the
hypothesis that the conformational change was essentially induced
by the binding of actinonin rather than from conformational
selection among multiple states occurring in the crystalline state.
To further investigate the mechanism involved, we followed a
kinetic approach aimed at discriminating between induced fit and
population shift at low ligand concentrations (sub-mM range) [12].
The experimentally observed pseudo-first-order rate constant for
the approach to equilibrium between the free components and the
binary AtPDF-actinonin complex (kobs) was measured and plotted
as a function of actinonin concentration. This plot yielded a
hyperbolic saturation curve with a positive slope, as fully expected
for a pure induced-fit mechanism (Figure 4B and C). In contrast, if
the enzyme sampled two or more conformational states, the curve
would imply that the value of kobs decreases with increasing ligand
concentration (see, for instance, curve C in Figure 1 in [12]). The
same conclusion can be reached for EcPDF and BsPDF2
(Figure 4B and C) and was already reported by others for S.
aureus PDF [29].
Together, these data indicate that a pure induced-fit mechanism
triggered by the binding of actinonin appears to direct the
conformational change both in solution and in the crystalline state.
Single Variants at Gly41 Exhibit Strongly Reduced
Actinonin-Binding Potency and Catalytic Efficiency
When dealing with an induced-fit mechanism, knowledge of the
initial O and final C state is crucial but does not provide direct
information on the position of actinonin in the encounter complex
or on the sequential mechanism of the transition process. We
suspected that the conserved glycine-rich motif 1 (G41IGLAAXQ)
could contribute to the flexibility required for the observed
structural transition. Evidence for such flexibility comes from
NMR analysis of EcPDF in which a few residues show exchange
cross-peaks of an additional, alternative form [38]. The most
strongly affected residues are Cys90, one of the metal ligands, its
neighbor Leu91, and both of the alanines within the above
conserved glycine-rich motif (Figure S1B), suggesting that EcPDF
undergoes conformational dynamics in a similar region.
To unravel the dynamics of the recognition process, we
surmised that it should be possible to freeze the conformational
Table 1. Comparison of the main kinetic and thermodynamic
parameters describing the inhibition of PDF by actinonin.
Parameter
AtPDFa
EcPDFa
BsPDF2a,b
KI (nM)d
140610
112610
185615
KI* (nM)c
0.960.5
1.360.2
2.960.8
KI/KI*
155615
86610
6467
k5 (s21) 6103d
6369
170620
7268
k6 (s21) 6104d
461
1962
1163
k4 (s21)e
140610
112610
185615
t1/2 (min)f
2965
661
1.160.2
aThe enzyme concentrations used in the assay were 100, 50, and 25 nM for
AtPDF, EcPDF, and BsPDF2, respectively.
bData from [49].
cPrior to kinetic analysis for determination of the KI* value, actinonin was
incubated at the final concentration in the presence of the studied enzyme set
for 10 min at 37uC. The kinetic assay was initiated by the addition of a small
volume of the substrate.
dFor determination of KI, k5, and k6 values, actinonin was not preincubated with
the enzyme. The kinetic assay was initiated by the addition of the enzyme.
ek4 corresponds to the kinetic constant of the dissociation of the primary
enzyme-actinonin complex. It is assumed that the rate of complex association
is diffusion-limited (see Table 7.3 in [19]), that is, k3—the kinetic constant of the
association of the primary enzyme-actinonin complex—is 109 M21.s21.
ft1/2 is 0.693(k4+k5+k6)/k4k6 (see case of induced fit and calculation in Table 1 of
[12]). In this case, t1/2,0.693/k6 because k6,,k5,,k4.
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Figure 2. Four distinct conformational states of PDF enzymes. (A) AtPDF and several other representative bacterial PDFs are superimposed. A
zoom is displayed on the right of the panel. Superimpositions were realized using ‘‘module superpose’’ in the CCP4i package and the ‘‘secondary
structure matching’’ tool. The extent of aperture/closure of PDF enzymes was assessed primarily by the measurement of the aperture angle (dap), the
angle made between the Ca of three strictly conserved residues (C, H, and I) of all PDFs, each characterizing a secondary crucial structure module of
the active site crevice, namely b4, a2, and b1 (see Figure S1C). Each single residue belongs to one of the three conserved motifs (motifs 2, 3, and 1,
respectively) and corresponds respectively to Cys91, His137, and Ile42 in AtPDF. The dap was measured in each case (see B). (B) The dap values
combined with those of the r.m.s.d. associated with the superimposition of the open structure of AtPDF allows the identification of four
conformational states: open (O), intermediate (I), closed (C), and super-closed (S). We compared AtPDF1B (this work and PDB CODE 3CPM; brown,
orange, and yellow in A and B; black in C), EcPDF (1BS7, free enzyme; 1BS6, with Met-Ala-Ser; 1G2A, with actinonin; magenta), BsPDF2 (1LQY, with
actinonin; green), LiPDF (1SV2, free; 1SZZ, with actinonin; red), and PfPDF (1JYM, free; blue).
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Figure 3. Effect of actinonin binding on the conformation of key residues in PDF. Conformation of key residues Ile42, Phe58, and Ile130 in
the different complexes: (A) in unbound WT AtPDF, (B and C) in the structure of G41Q and G41M actinonin-bound variants, respectively, and (D) of
actinonin-bound WT protein. In the final complex (D), a hydrogen bond is formed between actinonin and the peptidic bond, which links Gly41 and
Ile42. During the deformylation reaction, which is catalyzed by the PDF enzyme, the N-terminal formyl-methionine fits into the S1’ pocket. The
solvent-accessible surface of this pocket is represented here, and only the aliphatic chain of actinonin is shown, mimicking the N-terminal
methionine. (E) Free WT enzyme with the S1’ pocket shown open in two orientations (top and bottom). (F and G) S1’ pocket in the G41Q and G41M
variant structures, respectively, shown in two orientations (top and bottom). (H) After the complete conformational modifications of actinonin-bound
WT protein induced by actinonin binding, the S1’ pocket is shown closed in two orientations (top and bottom). (I) The four models are superimposed;
the ligand-binding site is magnified: unbound WT AtPDF; G41Q and G41M actinonin-bound enzyme; and WT actinonin-bound enzyme are indicated
in brown, red, orange, and yellow, respectively. Actinonin is indicated by lines. (J) A detailed view of the AtPDF ligand-binding site for all the
complexes, which are superimposed, as indicated in the same colors. Arrows indicate the direction of the closing movement within the enzyme, from
the open, unbound state to the closed, bound state.
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change along the pathway by introducing selected, minor
variations within the above-mentioned crucial residues involved
in the collective motion. In this respect, site-directed mutagenesis
of AtPDF was performed on Gly41, Ile42, and Ile130. Single
substitutions were made at Gly41 (G41A/Q/M), Ile42 (I42A/F/
N/W), and Ile130 (I130A/F), and the variants were purified and
characterized. These mutant proteins showed no change in overall
stability, as evidenced by DSC experiments (unpublished data).
However, two variants of G41, G41Q and G41M, showed
dramatic effects; the kcat/Km values were reduced by three orders
of magnitude due to large decreases in the kcat values compared to
the WT enzyme (Figure 5A and Table S1). The reduced kcat/Km
values suggest an altered ability of these variants to attain the final
enzyme-transition state complex and, as a result, to give rise to
possible states different from the final E:I* complex. Substitutions
at positions 42 and 130 only caused small reductions in the kcat
values (Figure 5A, Figure S2C, and Table S1). The actinonin-
binding potency of both G41 variants was also greatly reduced
(Table S1 and Figure S2B). The time-dependent inhibition by
actinonin of the most active variants was then studied (Table S3).
Figure 4. Evidence for an induced fit in crystalline and solution states of AtPDF. (A) Absence of evidence for alternative conformers in the
crystalline state of AtPDF. Ringer plots of electron density (r) versus x1 angle for representative residues of the 3D apo-structure of AtPDF. Data were
obtained with the 3M6O, 3PN2, and 3PN3 datasets (2.0 and 1.3 A˚ resolution, respectively, see Table S1). The secondary peaks in the Ile residues are
observed because Ile is a branched amino acid. To reveal an alternative conformation with Ile, three peaks should be observed. (B) kobs is a saturable
function of actinonin with various PDFs, including AtPDF. Data obtained for kobs, the experimentally observed pseudo-first-order rate constant for the
approach to equilibrium between the free components and the binary PDF-actinonin complex, were obtained at various concentrations of actinonin
in the presence of EcPDF, AtPDF, and BsPDF2. A direct plot is shown. Inset, time-course measurement of deformylation as a function of varying
actinonin concentrations. (C) Inverted plot of the data in panel B, which is expected to be a straight line if the kobs is ..k6 in the case of induced fit
[19]. The correlation coefficient of each line is 1.00, 0.99, and 1.00 for AtPDF, BsPDF2, and EcPDF, respectively, indicative of the accuracy of the
conclusion.
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The half-lives of the final complexes—as assessed by comparison
of the 1/k6 values—were always significantly smaller (Table S3),
suggesting that the conformational change induced by actinonin
binding still occurred, but the C state is destabilized relative to the
O state in the mutants compared to the WT. Accordingly,
actinonin strongly stabilized almost all of the variants; Tm was
increased by more than 20uC. This differs from the G41M and
G41Q variants, which both showed increases in the Tm of only
12uC, consistent with reduced binding potency (Table S1).
Conformational Changes of Gly41 Variants Are Affected
On-Pathway
The two most interesting variants, G41Q and G41M, could be
crystallized under the same conditions as the WT protein. In the
case of G41Q, the structure of the apo-protein did not show any
modifications compared to the WT structure and remained in an
O conformation (Figure 2B; ‘‘O’’ zone). In contrast, the 3D
structure of the G41M variant showed that the asymmetric unit
was composed of two molecules with distinct structures. One
molecule (chain A) is in the O state and is similar to the structures
of the WT and the G41Q variant (Figure 2B; zone ‘‘O’’). The
second molecule (chain B) is in a C state, closer to that observed
for the WT chain in the presence of actinonin (‘‘C’’), a so-called
‘‘superclosed’’ state (Figure 2B; zone ‘‘S’’), suggesting that the
substitution modified the equilibrium between the two states in
solution either (i) at the step of protein synthesis by providing two
conformers, the inter-conversions of which are blocked due to
steric hindrance brought by the new bulkier side-chain at position
41, or (ii) by dramatically unbalancing the free inter-conversion
between the O and S conformers towards the S state. Ringer
analysis indicates that in the free G41M variant, many residues
show evidence for unmodeled alternate conformers—including
positions 58, 42, and 130—in keeping with the second hypothesis.
For all variants of position G41, addition of actinonin to the
crystal (Figure 3 and Figure S6) induced a closure of the protein
within the crystal. Nevertheless, as expected from in silico graphic
modeling followed by energy minimization, the occurrence of a
bulky side chain at position 41 prevented the completion of the
closure in the presence of the ligand and, hence, the formation of
the hydrogen bond between the backbone nitrogen of Ile42 and
actinonin. This finding is consistent with the strongly reduced Tm
of the complex of the variants with actinonin compared to WT as
measured by DSC. Remarkably, both S and O forms of the G41M
apo-structures in the asymmetric unit of the crystal yielded a
unique intermediary structure (‘‘I’’ state) upon actinonin binding
(r.m.s.d. between the molecules is ,0.25 A˚ ; see also Figure 2B,
zone ‘‘I’’). In this case, it is likely that the induced-fit mechanism
drives the equilibrium by capturing only the O population and
closing it to an intermediary step, thus depleting the pool of O
conformers that is shifted sequentially back from the remaining
pool of S conformers and allows the complete binding of actinonin
to the enzyme.
In line with the rational design of the PDF mutants, the extent
of the structural differences suggests that the underlying motions
are dependent on the length of the side chain (Figure S8).
Together, these data account for the reduced catalytic rate, as the
hydrogen bond is strictly required for the substrate to be efficiently
cleaved by PDFs (Figure S8A) [54]. Therefore, from both
structural and kinetic analyses, each substitution most likely
reproduces intermediates along the pathway that lead to the
closure of PDF around its substrate (Figure S2B).
Conformational Changes of Gly41 Variants Recapitulate
Closing Intermediates
Analysis of the structures allows us to propose the following
sequence of atomic events (Figures 3 and 2B and Figure S6). To
name the various sites of the ligand and subsites of PDF, we will
use the usual nomenclature found in [55], which defines the
various binding pockets of a protease, where P1’ is the first side
chain at the C-terminal side of the cleavage site and its binding
pocket is S1’, also referred to as the hydrophobic pocket in the case
of PDF. First, actinonin aligns along the S1’ pocket to form the
encounter complex, which shifts the Ile130 side chain to avoid
steric hindrance in the S1’ pocket, promotes rotation of the Ile42
side chain, and finally rearranges the phenyl group of Phe58.
These
events
achieve an
optimal
hydrophobic
S1’
pocket
conformation (Figure 3), and the concomitant closure leads to
the formation of a hydrogen bond between the first carbonyl
group of actinonin and the backbone nitrogen of Ile42. The initial
N-O distance is reduced from 5 A˚ to 2.8 A˚ , which is an optimal
value for hydrogen bonding (Movie S2 and Figure S8B). Thus, the
primary driving force for the active site closure appears to be the
P1’:S1’ hydrophobic interaction. The C state is ultimately locked
by the super-b-sheet hydrogen bonds extending across the ligand,
including those involving Ile42. The DDGbinding value (2.2–
2.4 kcal/mol, Figure S8B), as calculated from the Kd values for
actinonin binding to wild-type (WT) and G41M and G41Q, is
consistent with the loss of a hydrogen bond that also contributes to
the conformational stability of the protein [56,57]. Thus, this bond
contributes to the major binding free energy difference between
the two complexes (3.1 kcal/mol; Figure S8B, Tables S1 and S3,
and [29]). Interestingly, the above DDGbinding values also correlate
with the DDGES values derived from the kcat/Km and kcat
measurements [19]. This dataset strongly correlates with the
Figure 5. Inhibition and enzymatic reactions progress through an induced fit pathway. (A) The catalytic parameters Km and kcat, for all
AtPDF variants are provided as a percentage of the wild-type values (WT). Detailed values are presented in Table S2. (B) Schematic model for
actinonin binding to AtPDF in favor of an induced-fit pathway. PDF might exist in at least two conformational states, open (O) or closed (C). The
relative abundance of each conformation would vary, depending on the enzyme type. With AtPDF, it is likely that the most abundant form is the O
one, which is the only form leading to a productive complex. The superclosed form (S) is likely to show reduced affinity for the ligand because of
steric occlusion of the active site. At the initial stage, the inhibitor (shown in red) binds to AtPDF (indicated in brown) in the O conformation. To reach
the final key-lock state (productive closed conformation, C), two major and extreme pathways can be used. According to the conformational selection
pathway, the inhibitor selects the C conformation. This pathway, which is represented by the dashed arrow, does not occur within the crystal. In
contrast, the G41Q and G41M mutants, by providing the structure of the enzyme in intermediate conformations (I), prove the existence of the so-
called encounter complex and confirm that the inhibitor binds to the enzyme when it is in the O conformation. The ligand-binding site is then
reorganized to yield the C enzyme conformation, that is, the key-lock state. Indeed, the inhibitor binds to the enzyme through the induced-fit
pathway. Each timescale was calculated using the data available in the text and corresponds to t1/2 values deduced from the calculation of 0.693/
(kinetic constant of interest). The kcat value (k2) was used to assess the timescale of catalysis in panel C, whereas, in (B), k4 assesses the first step of
inhibition, and k6 is used in the case of the slow step. For the SO conversion (left, B), the lifetime of the minor form of EcPDF was used to assess the
order of magnitude (see text and [38]). (C) Schematic model for the deformylation reaction catalyzed by PDF. Since actinonin is a pseudo-peptidic
inhibitor, it is likely that a peptidic substrate can bind to the PDF enzyme through an induced-fit pathway, as described in (B). The key-lock state
represents a transition state in which the N-formylated substrate is deformylated to yield the final reaction product.
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gyration and van der Waals radii of the side chain at position 41 as
well as the N-O distance between the first carbonyl group of
actinonin and the backbone nitrogen of Ile42 (Figure S8). These
results suggest that the capacity of both G41M and G41Q variants
to form the transition state is a consequence of their inability to
reach the fully closed state.
Thus, our study of the designed Gly41 mutant enzymes reveals
that, in addition to the initial and final states observed for the WT
enzyme, the conformations of the Gly41 variants correspond
indeed to on-pathway intermediates, thus providing snapshots
along the trajectory from the O to the C state of the enzyme
(Figures 2B and 3). The 3D structure of the variants in the absence
of ligand is similar to that of WT, and a strict correlation exists
between the completeness of the conformational change and both
binding potency and catalytic efficiency. This suggests that both
events require complete protein closure to generate a productive
complex.
The
strong
stabilization
of
AtPDF
by
actinonin
(Figure 1D) closely mimics what occurs with its natural substrates
when it reaches the transition state [34,58]. Indeed, as expected,
the enzyme facilitates the final C conformation by lowering its
final energy [6]. Optimal arrangement of the S1’ pocket (Figure 3)
proceeds
along
the
reaction
process
towards
the
final
C
conformation, triggering the alignment of reactive groups in an
optimal arrangement for ligand recognition. Upon binding,
actinonin alters the thermodynamic landscape for the structural
transition between the O and C states. This ligand is a potent
inhibitor because it can trigger the above sequence of events
similar to the substrate, but unlike the substrate, it is non-
hydrolyzable. Thus, by mimicking the transition state and being
non-hydrolyzable (Figure 1B), the final C complex is long lasting.
Ligand-Induced Conformational Closure Is Initially
Triggered by the Binding of the P1’ Group in the S1’
Pocket
Given the similarity between actinonin and natural substrate
binding, the very slow kinetics of inhibitor binding (10-s time-scale)
remains puzzling compared to the 10 ms required for catalysis
(deduced from the kcat). This finding could be explained as a
conformational effect during the formation of the hydrogen bond,
aligning the substrate as an additional beta-sheet and eventually
stabilizing the entire enzyme-ligand complex. The significantly
longer time needed to reach the most stable state compared to the
substrate would most likely be due to the presence of the flexible
and one carbon longer metal-binding group in actinonin (i.e.,
hydroxamate versus formyl, Figure 1B). This suggestion is in line
with the overall data obtained when we investigated more deeply
the role of the first carbonyl group of the ligand. This group is well
known
to
exert
a
crucial
effect
in
both
productive
and
unproductive ligand binding (i.e., substrate and inhibitor) [54].
In this respect, we studied the binding of compound 6b (Figure
S5B), a PDF ligand that does not exhibit a reactive group at this
position [49]. We observed that this compound binds strongly to
both EcPDF (KI* = 6366 nM) and AtPDF (KI* = 400635 nM)
but, unlike actinonin, does not display slow, tight binding as KI* =
KI. This impact on binding is consistent with the absence of the
hydrogen bond involving the first carbonyl group of the ligand.
The 3D structure of AtPDF was determined after soaking the
compound in crystals of the free, open AtPDF form. Upon binding,
6b induced a complete conformational change, identical to that
observed with actinonin (Figures 2B and 6A; ‘‘O’’ state). This
result further suggests that the conformational change is not
induced initially by the formation of this hydrogen bond and that
the encounter complex is primarily driven by the fit within the S1’
pocket. This also reveals that the timescale of the large
conformational change is several orders of magnitude faster than
the kinetics of slow binding and fully compatible with both the first
step of actinonin binding (k4 = 140 s21; see Table 1) and the
catalytic rate of the substrate (kcat = 37 s21; see Table 1 and Table
S3). The 3D structure also revealed that both the P1’ and the
hydroxamate groups are bound similarly to the corresponding
groups of actinonin (Figure 6B). As expected, no additional
bonding occurs, especially around the backbone nitrogen of Ile42
(Figure 6C).
Taken together, these data allow us to conclude that the
conformational change observed upon ligand binding is triggered
primarily by binding in the S1’ pocket. As revealed by the binding of
6b, the one carbon longer metal-binding group fits, immediately
upon recognition of the P1’ group, in the S1’ pocket and forms a
bidentate complex with the metal cation, mimicking the transition
state as a result. Thus, the active site is very confined and rigid due
to the presence and length of the hydroxamate group (compare
right and left panels in Figure 1B). As a result, compared to the
complex made with the substrate, it is likely that the formation of the
hydrogen bond involving the carbonyl of actinonin and the
backbone nitrogen of Ile42 becomes strongly rate-limiting (k5
= 0.044 s21; Table 1). Once this hydrogen link is locked, the
uncleavable bond, mimicking the labile formyl group at the
transition state, stabilizes the enzyme-inhibitor complex, making it
long-lasting (k6 = 0.0006 s21; Table 1) and providing a mechanistic
explanation for the slow-binding effect that involves both large and
fine conformational changes. The large conformational change is
similar to the one occurring with the substrate, whereas the second is
more subtle and locks the hydrogen bond involving the backbone
nitrogen of Ile42. The second step is rate-limiting with some
transition state analogs such as actinonin (Figure 5B and C).
Proper Positioning of the Carbonyl Group Is Required to
Stabilize the Complex at S1’
Compound 21 corresponds to another interesting derivative
designed to probe the impact of the peptide bond in PDF binding
[49]. In addition to the hydroxamate group, this compound
features both a hydrophobic benzyl group at P1’ and a reverse
peptide bond. Compound 21 shows modest but significant
inhibitory potency to AtPDF1B (KI* = 400637 nM), confirming
the crucial role of the peptide bond in PDF binding. After soaking
with crystals of apo-AtPDF, compound 21 could be detected in
high-resolution electron density maps (Figure S9A). Unlike 6b, 21
did not bind the active site of the enzyme but an alternative pocket
at the surface of the protein (Figure S9B). A docking study
performed with EcPDF had previously revealed this alternative
binding pocket (Figure S9C; [59]).
The aforementioned data indicate that the occurrence of a S1’-
binding group placed in the unfavorable context of a reverse
peptide bond does not stably promote binding at the active site of
AtPDF. Upon binding of 21, the 3D structure of both molecules of
the asymmetric unit remain in an O conformation (r.m.s.d.
,0.2 A˚ with respect to the apo-structures in the ‘‘O’’ state). This
finding suggests that only the binding of compounds entering the
S1’ pocket, such as actinonin or 6b, induces conformational
change, in keeping with the crucial role of the P1’ group if located
in the frame of a classic peptide bond. Moreover, we noticed that
the binding pocket of 21 was located on the rear side of the true
S1’ pocket and induced a weak modification of the P1’ hosting
platform (Figure S9D). Indeed, when crystals of the 21:AtPDF
complex were soaked in actinonin, the final 3D structure no longer
showed evidence of compound 21 occupancy greater than 5%.
Instead, this structure revealed both actinonin and closing of the
protein (Table S2). The r.m.s.d. between this structure and that
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obtained directly with actinonin was less than 0.2 A˚ ; the actinonin
position was virtually identical, indicating that the protein had
retained full capacity for binding actinonin and closing despite the
presence of compound 21. We conclude that actinonin does
compete with 21 because of the overlap at P1’ of AtPDF1B (Figure
S9C). As the actinonin S1’ subsite strongly mimics that of a true
substrate, this result also explains the inhibitory behavior of 21
towards AtPDF.
Discussion
Although PDF catalysis has been extensively studied and the
mechanism has been elucidated [34], how the enzyme achieves the
catalytically competent state remains unknown. Here, we provide
insight on how the enzyme might reach a catalytically competent
conformation, demonstrating that the reactive groups move into
proximity to promote catalysis (Figures 2B and 5C). We suggest
Figure 6. Effect of 6b binding on the conformation of key residues of PDF. Superimposition of free, 6b-, and actinonin-bound AtPDF
indicated in brown, red, and yellow, respectively. (A) Molecule A in the three models was superimposed, resulting in an r.m.s.d. of 0.9 A˚ for 100% of
the Ca. Actinonin is shown in yellow and 6b in red. (B) Conformation of key residues Ile42, Phe58, and Ile130 in the different complexes and in
unbound WT AtPDF. Actinonin is shown in yellow and 6b in red. (C) A detailed view of the AtPDF ligand-binding site for both actinonin and 6b
complexes, which are indicated by sticks and are superimposed. The two ligands are colored in pale and dark grey, respectively. The hydrogen bond
made by actinonin only is shown.
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that the motions of the catalytic centre starting with free ligand-
PDF favor a final configuration that is optimal for binding and/or
catalysis (Figures 1B, 2B, and 5B and C). In our model, we propose
that free PDF might exist in at least two conformational states, that
is, open (O) or super-closed (S). The relative abundance of each
conformation varies by enzyme type and incubation conditions,
explaining why both conformations have not been trapped thus
far. In the case of AtPDF, it is likely that the most abundant form
corresponds to an O state, which is the form that leads to a
productive complex. Indeed, in the NMR spectra for EcPDF, a
few residues show exchange cross-peaks from an additional,
alternative form [38]. The most strongly affected residues are
Cys90, one of the metal ligands, its neighbor Leu91, as well as
Ala47 and Ala48 on the facing strand. This suggests that EcPDF
exists in at least two conformations (‘‘S’’ and ‘‘C’’; see Figure 2B),
which undergo slow interconversion on the NMR timescale. The
3D structure of the major conformation (75%, lifetime 300 ms)
could be solved at high resolution, but the structure of the minor
form (25%, lifetime 100 ms), which exhibits very weak signals,
could
not
be
solved
[38].
This
conformation
appears
to
correspond to that of the complex obtained with the product of
the reaction (Met-Ala-Ser). A very similar situation—although
more balanced between the two states—appears to occur in the
case of variant G41M, suggesting that a mechanism involving
conformational selection followed by induced fit is a general model
for PDF and that AtPDF is a specific case where population shift
virtually does not occur as the free enzyme is completely in the O
conformation. This is also in line with data obtained with L.
interrogans PDF (LiPDF), which reveal conformers in both the S and
C states (see Figure 2B) and suggest a population-shift mechanism
[43]. It is interesting to note that LiPDF is a poorly active PDF
[60]. According to the representation shown in Figure 2B,
Plasmodium falciparum PDF (PfPDF), a poorly active PDF [61],
was retrieved only in the S state. Finally, weak decompaction of
the structure of Bacillus cereus and Staphylococcus aureus PDFs in the
presence of actinonin have been described [45,46]. These
examples suggest that the enzyme is trapped in the S conformer
in the free state and converts to the C conformer when bound to
actinonin, suggesting that the S conformer is overrepresented in
solution compared to the O state, unlike AtPDF.
This study of AtPDF—including 10 different crystal structures of
apo- and complexed enzyme variants—reveals the 3D structure of
a PDF in at least four distinct states. This includes the O form, the
occurrence of which is crucial for catalysis, as it is the active form.
Here, we propose that the transition from the O to the C state is
directly induced by the ligand. Indeed, the O form, which is
captured in the crystal, undergoes closure directly upon ligand
binding in our soaking experiments. Progression to this closure
involves intermediary states (‘‘I’’) similar to those observed with
variants G41Q and G41M in the presence of actinonin (see
Figure 2B). Extrapolating the situation to catalysis, which occurs in
the crystalline states of PDF, it is likely that hydrolysis of the
substrate frees the enzyme in its S state, which in turn needs to
open to accommodate a new substrate (Figures 2B and 5C). This is
well illustrated in the 3D structure of EcPDF complexed with a
product of the reaction, obtained after co-crystallization of the
enzyme with the substrate in a closed conformation [34]. The S
free form is likely to exhibit a slower on-rate for the ligand (k3)
compared to the O form because of steric occlusion of the active
site (Figure S10). In support of this hypothesis, recent data show
that the 3D structure of a C-terminally truncated, poorly active
version of AtPDF is in the C conformation in the unbound state,
although crystallized under conditions identical to ours [62,63].
This structure is similar to that of chain B, one of the two
molecules of the asymmetric subunit of variant G41M (Figure 2B).
This suggests that alterations remote from the active site
significantly unbalance the equilibrium between the two conform-
ers, thus altering the efficiency of the reaction (Figure 5C). As the S
version corresponds to a significantly less active version of AtPDF
compared to that reported in our present work, this further
confirms that, compared to the O state, the S state has a
significantly weaker propensity to bind substrate or a close mimic
ligand, such as actinonin. Comparison of the 3D structures of the
free-closed and the ligand-bound-closed forms reveals some
differences responsible for the slight steric reduction of the active
site of free-closed AtPDF1B with respect to that of the actinonin-
AtPDF1B complex (Figure S10A), including the side chain of Ile42
burying the S1’ binding pocket (Figure S10B). Overall, these data
suggest that an S form might exist under the free state but that it
would feature a k3 value with respect to the ligand that is
significantly weaker than that of the O form, which would strongly
slow down the reaction or the binding as a result.
With the interaction scheme proposed in our model (Figure 5B
and C), the ligand/substrate binds more easily to the O form and
induces the optimal conformation of the enzyme to reach the
transition state, thus allowing the reaction to be efficiently
catalyzed.
In
the
final
model
(Figure
5C),
there
is
both
conformational selection and induced fit subsequently involved
in line with the recently proposed existence of such mixed
mechanisms for other enzymes [15,16]. Nevertheless, in our model
(Figure 5C), we suggest that induced fit is the primary mechanism,
as it provides energy input from the ligand, which eventually drives
the enzyme towards the productive key-lock complex. Unambig-
uous distinction between the relative contributions of the two
mechanisms is deduced from the observation that kobs is a saturable
function of actinonin with various PDF, including EcPDF, BsPDF,
AtPDF (Figure 4B and C), and S. aureus PDF [12,16,29,49].
Using crystallographic reconstruction analysis involving enzyme
variants, motions of small mobile loops and movie reconstructions
of snapshots of catalytic events have been previously documented
[1–3,64–66], often by visualizing the binding of unnatural
inhibitors and not necessarily mimicking closely the substrate
and transition state as actinonin does [67,68]. However, only a few
examples make use of soaking conditions of a crystal to promote
the motion and show the importance of induced fit [1,69]. None of
these data show a motion of the amplitude revealed here with PDF
and a large stabilization of the complex involving the formation of
the four-stranded b-sheet superstructure and the entire N-domain
of the enzyme. Compared to previous crystallographic analyses,
our work integrates biophysical, computational, and kinetic
analyses to reconstruct the whole picture, allowing a better
understanding of the slow-binding mechanism.
While our work primarily focused on an induced-fit mechanism
of enzyme inhibition and catalysis, it should be emphasized that
this phenomenon is also applicable to the broader area of
receptor-ligand interactions. For example, in all cases where
conformational change mechanisms have been proposed for
kinase inhibitors without supporting experimental data [12,26],
further experimental work must be provided to clarify the precise
mechanism. We expect this will have important implications on
how one conducts future drug-discovery efforts against such
enzymes [70].
Materials and Methods
Protein Expression and Purification
Expression and purification of mature Arabidopsis thaliana PDF1B
and all variants (i.e., AtPDF) were derived from the previously
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12
May 2011 | Volume 9 | Issue 5 | e1001066
described protocol [37]: the lysis supernatant after sonication was
applied on a Q-Sepharose column (GE Healthcare; buffers A and
B as described containing 5 mM NiCl2) followed by Superdex-75
chromatography (GE Healthcare) using buffer C consisting of
buffer A supplemented with 0.1 M NaCl. For crystallization
experiments, the protein was purified further. The sample was
concentrated on an Amicon Ultra-15 centrifugal filter unit
(Millipore Corp.) with a 5-kDa cut-off and applied to a MonoQ
HR5/5 column (GE Healthcare) previously equilibrated in buffer
A (50 mM Hepes, pH 7.5, and 5 mM NiCl2). Elution was
performed with a 50-mL gradient from 0% to 100% buffer B.
The buffer of the pooled purified AtPDF1B was exchanged using a
PD-10 desalting column (GE Healthcare) to yield a protein
solution in 50 mM Hepes, pH 7.5, 0.1 M NaCl, and 5 mM NiCl2
(buffer C). The protein was concentrated on an Amicon Ultra-15
centrifugal filter unit. The resulting AtPDF1B preparation was
frozen in aliquots and stored at 280uC (for crystallization
purposes) or diluted 2-fold in 100% glycerol and stored at
220uC (for enzymatic purposes). The typical yield was 5–10 mg
AtPDF per liter of culture. All purification procedures were
performed at 4uC. Samples of the collected fractions were
analyzed by SDS-PAGE on 12% acrylamide gels, and protein
concentrations were estimated from the calculated extinction
coefficients for each variant.
Site-directed mutagenesis of AtPDF sequence in plasmid
pQdef1bDN [36] was carried out using the QuickChange Site-
Directed Mutagenesis Kit (Stratagene).
Enzymology
Assay of PDF activity was coupled to formate dehydrogenase,
where the absorbance of NADH at 340 nm was measured at 37uC
as previously described [71]. For measurements of classical kinetic
parameters (i.e., Km and kcat), the reaction was initiated by addition
of the substrate Fo-Met-Ala-Ser to the mixture containing purified
enzyme in the presence of 1 mM NiCl2. The kinetics parameters
were derived from iterative non-linear least square calculations
using the Michaelis-Menten equation based on the experimental
data (Sigma-Plot; Kinetics module). For determination of kinetic
parameters related to actinonin, the reaction mixture contained
750 mM NiCl2. In some cases, the mixture containing PDF and
actinonin was incubated for 15 min at 37uC before kinetic
analysis, which was initiated by the addition of substrate. The
same protocol was used to determine the dissociation constant of
actinonin [KI* = k4/(k3+k3k5/k6)], but the initial reaction velocities
were measured with varying concentrations of Fo-Met-Ala-Ser and
actinonin. The data were then calculated according to the method
of Henderson, which can be used to determine the dissociation
constant
of
the tight-binding
competitive enzyme
inhibitor
[28,49,72] by varying both the inhibitor and substrate concentra-
tions. To determine KI, k5, and k6, the reaction was initiated by the
addition of enzyme as previously described [29,49]. KI*app
measurements were used for comparative studies of AtPDF
variants (Table S3) at a concentration of 2 mM substrate by
varying the concentration of actinonin. KI*app is the slope of the
v[Actinonin]/v0 line curve. kobs was fitted from the kinetic data without
preincubation with vI = vs + (v0 2 vs)e2kobst where vI is the observed
velocity at a given concentration of inhibitor I, v0 is the velocity,
and vs is the steady-state velocity [18]. From the set of values
obtained at various concentrations of I, k5 and k6 could be derived
using kobs = k6 + k5[I]/(KI + [I]). By choosing a set of values with
kobs..k6, 1/kobs
= 1/k5(KI/[I] +1) and 1/kobs
=
f(1/[I]) is
expected to be a straight line in case of induced fit whose positive
slope corresponds to 1/k5. k6 was derived from equation k6 = k5/
(KI/KI*21) [18,19].
Microcalorimetry
ITC experiments were performed using a VP-ITC isothermal
titration calorimeter (Microcal Corp.). Experiments were per-
formed at 37uC. For each experiment, injections of 10 mL
actinonin (180 mM) were added using a computer-controlled
300 mL microsyringe at intervals of 240 s into the Ni-AtPDF
variant solution (5 to 10 mM, cell volume = 2.1 mL) dissolved in
buffer C with stirring at 310 rpm. A theoretical titration curve was
fitted to the experimental data using the ORIGIN software
(Microcal). This software uses the relationship between the heat
generated after each injection and DHu (enthalpy change in kcal/
mol), KA (the association binding constant in M21), n (number of
binding sites per monomer), total protein concentration, and free
and total ligand concentrations. The thermal stability of the WT
and variants of Ni-AtPDF1B was studied by DSC using VP-DSC
calorimetry (Microcal Corp.). DSC measurements were made with
10 mM protein solutions in buffer C. The actinonin concentration
was 20 mM. The same buffer was used as a reference. All solutions
were degassed just before loading into the calorimeter. Scanning
was performed at 1uC/min. The temperature dependence of the
partial molar capacity (Cp) was expressed in kcal/K after
subtracting the buffer signal using Origin(R) software.
Crystallization and Soaking Experiments
Crystallization conditions were screened by a robot using the
sitting drop vapor diffusion method. Crystals were obtained and
optimized at 20uC with 15%–20% PEG-3350 and either 0.1 or
0.2 M zinc acetate. The drops were formed by mixing 2 mL of a
solution containing 2 to 4 mg/mL protein and 2 mL of the
crystallization solution. Crystals were soaked for 24 h by adding
actinonin to the crystallization drops at a final concentration of
5 mM. Cryoprotection was achieved by placing crystals for 30 s in
a solution that was composed of 20% PEG-3350 and 0.2 M zinc
acetate, supplemented with 5%, 10%, and 15% glycerol. Crystals
were then directly flash frozen in liquid nitrogen using cryoloops
(Hampton Research). Crystals were also grown under conditions
described for the C-terminally deleted, weakly active version of
AtPDF [63].
X-Ray Diffraction Data Collection
Data collections were performed at 100 K at the European
Synchrotron Radiation Facility (Grenoble, France) on station
ID29, FIP-BM30A, ID14-1, and ID23-2, and at SOLEIL (Gif-sur-
Yvette, France) on station PROXIMA1. In each case, a single
crystal was used to collect a complete dataset. Data were processed
and scaled using XDS software [73]. Two crystal forms were
encountered with different cell parameters. In each case, b
parameter was nearly equal to a, and data could be indexed into
two space groups, P212121 or P43212. The data are shown in Table
S2.
Structure Determination and Refinement
The
structure
of
free
AtPDF
was
solved
by
molecular
replacement with Phaser [74] followed by a rigid-body refinement
by CNS [75] using coordinates from the Plasmodium falciparum PDF
(PDB code 1RL4) [76] as a search model. The structures of
actinonin-bound proteins—that is, WT and mutants—were solved
using rigid-body refinement by CNS of the free AtPDF structure.
The ten final models were obtained by manual rebuilding using
TURBO-FRODO [77] and combined with refinement of only
calculated phases using CNS and Refmac [78] software. No non-
crystallographic symmetries were used. Quality control of the
three models was performed using the PROCHECK program
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13
May 2011 | Volume 9 | Issue 5 | e1001066
[79]. To probe for alternative conformers, Ringer was used [53].
Ringer is a program to detect molecular motions by automatic X-
ray electron density sampling, and can be accessed at http://
ucxray.berkeley.edu/ringer.htm.
Accession Numbers
PDB codes for the PDF structures presented within this
manuscript are as follows: 3M6O, 3PN2, 3M6P, 3O3J, 3PN3,
3PN4, 3PN5, 3M6Q, 3PN6, and 3M6R. UniProtKB accession
numbers for other PDF studied are P0A6K3 (EcPDF) and O31410
(BsPDF).
Supporting Information
Figure S1
Alignment of PDF sequences and secondary structures.
(A) PDF1B from Arabidopsis thaliana (AtPDF1B) is compared with
bacterial type 1B (EcPDF and LiPDF), pathogenic protozoa
(PfPDF1B), eukaryotic mitochondrial PDF1A from A. thaliana
(AtPDF1A), and bacterial type 2 (BsPDF2). This figure was created
with ENDscript [80]. The sequence alignment was realized with the
algorithm muscle included in ENDscript, and modified according to
the superimposition of structures. The blue frames indicate
conserved residues, white characters in red boxes indicate strict
identity, and red characters in yellow boxes indicate homology. The
secondary structures at the top (a-helices, 310 helices, b-strands, and
b-turns are shown by medium squiggles, small squiggles, arrows,
and TT letters, respectively) were predicted by DSSP [81]. Relative
accessibility (acc) of subunit A is shown by a blue-colored bar below
sequence. White is buried, cyan is intermediate, and blue with red
borders is highly exposed. A red box means that relative accessibility
is not calculated for the residue, because it is truncated. Hydropathy
(hyd) is calculated from the sequence according to [82]. It is shown
by a second bar below accessibility: pink is hydrophobic, grey is
intermediate, and cyan is hydrophilic. Motifs 1 (41GwGwAAXQ48),
2 (89EGCLS93), and 3 (133HEwDH137), where w is a hydrophobic
amino acid, are labeled by red stars below the sequence alignment.
To simplify the nomenclature, AtPDF1B is referred to as AtPDF
throughout the text. (B) Topology cartoon of AtPDF, free (left) or
actinonin bound (right), in the same color code as (A). Actinonin
(represented by the yellow arrow) binding to the ligand binding site
allows the linkage of the two distinct b-sheets into one single b-sheet,
by mimicking an additional b-strand. PDB sum (http://www.ebi.ac.
uk/thornton-srv/databases/pdbsum/) was used. (C) 3-D structure
of AtPDF is represented showing the position of the residues
discussed in the text, indicated in red.
(EPS)
Figure S2
Microcalorimetric titration of AtPDF with actinonin.
Data were obtained at 37uC by an automated sequence of 28
injections of 180 mM actinonin from a 300 ml syringe into the
reaction cell, which contain 9.85 mM AtPDF. The volume of each
reaction was 10 ml, and injections were made at 240 s intervals.
Top, raw data from the titration. Each peak corresponds to the
injection. Bottom, the peaks in the upper panel were integrated
with ORIGIN software and the values were plotted versus
injection number. Each point corresponds to the heat in mcal
generated by the reaction upon each injection. The solid line is the
curve fit to the data by the Origin program. This fit yields values
for Kd. Experiments were done with wild type protein and others
variants, and gave similar raw data and curve fit. (A) WT; (B)
variant G41M; (C) variant I42W.
(EPS)
Figure S3
Binding of actinonin to AtPDF does barely modify the
crystal packing. (A) Crystal pack of the two complexes: open, free
complex (left) and bound to actinonin (right) (B). Non-crystallo-
graphic contacts into asymmetric unit are not modified by closing
movement of the protein due to actinonin binding, except for zinc
atom number 6. This metal ion is coordinated by side chains of
Asp40 and Glu63, and water molecules, Asp40 and Glu63 being
hydrogen bonded by side chain of Lys38 of the other subunit of
the asymmetric unit. With the closing movement of the protein
into the crystal, Ca of Asp40 shifted by 3.1 A˚ and its side chain
flipped by 90u. Therefore, it does no longer participate to the
coordination shell of this Zn2+ ion. However, it is still hydrogen
bonded by Lys38 from chain B.
(EPS)
Figure S4
Binding of actinonin to AtPDF closely mimics both
actinonin and product binding to EcPDF. Superimposition of
EcPDF and AtPDF bound to either actinonin (1LRU PDB code,
panel A) or Met-Ala-Ser (1BS6 PDB code, panel B), the product of
the
reaction.
The
r.m.s.d.
value
is
1.11
A˚
for
151
Ca
superimposed.
(EPS)
Figure S5
The ligand binding site of AtPDF. This picture shows
the residues of AtPDF that are in contact with actinonin (left) and
6b (right) according to the 3-D structure; this should be compared
to the similar scheme shown in Figure 1B for EcPDF.
(EPS)
Figure S6
Electronic densities of the moving side-chains and of
actinonin at the binding site in some variants of AtPDF. Actinonin
and selected residues (G/Q/M41, I42, F58, and I130) are drawn
in stick and are shown in their FO–FC electron density omit maps
contoured at 2s, in free wild-type AtPDF (two crystallization
conditions, WT1 and WT2), and ligand-bound WT (actinonin, 6b
and 21), G41Q, and G41M variants.
(EPS)
Figure S7
Only few residues show alternative conformation in
AtPDF. Alternative conformers in the crystalline state of AtPDF.
Ringer
plots
of
electron
density
(r)
versus
x1
angle
for
representative residues of the 3-D apostructure of AtPDF. Data
were obtained with the 3M6O dataset (see Table S1). The
secondary peaks in the Ile residues are observed because Ile is a
branched amino acid. To evidence an alternative conformation
with Ile, three peaks should be observed.
(EPS)
Figure S8
Impact of induced fit on the binding free energy of
actinonin depends on the capacity to stabilize a hydrogen bond
with PDF. (A) The gyration radii [83] of the side chain occurring
at position 41 is displayed with black squares and compared to the
kcat/Km values (grey bars). (B) The distance between the NH of I42
and the CO of actinonin was measured in each case. The
percentage of the distance required to make a hydrogen bond (2.8
A˚ ) is reported (dark squares). The difference of binding free energy
(DDGbinding) between the open, free state and the variants closed
complexes of the G41 variants are displayed as grey bars. The
values were calculated as follows. For the WT, it corresponds to
the RT ln(KI*/KI) value [29], where R is the ideal gas constant and
T is the temperature in Kelvin. RT is 0.616 kcal.mol21 at 37uC.
For the G41M and G41Q variants, the DDGbinding corresponds to
RT ln(KI-G41variant/KD-WT). The obtained values are similar to that
obtained if the kcat/Km substitutes the KD value in the calculation
(DDGbinding = RT ln(kcat/Km –G41variant/kcat/Km –WT).
(EPS)
Figure S9
Compound 21 does not bind AtPDF1B at S1’. (A) 21
is shown in ball-and-stick format in its FO–FC electron density omit
The Dynamics of Induced Fit at High Resolution
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May 2011 | Volume 9 | Issue 5 | e1001066
map contoured at 2s. (B) Binding site of 21 into AtPDF1B is
detailed. Red and blue residues indicate residues that accommo-
date
the
‘‘phenylalanine’’
and
‘‘trimethyl’’
groups
of
21,
respectively. (C) Overall view of 21 binding site (left). Molecular
surface of AtPDF is represented, as well as 21 in ball-and-stick
format. Residues belonging to the 21 binding pocket are colored in
orange. For comparison, molecular surface of EcPDF (PDB code
1G2A) in the same orientation is also represented, with residues
forming the new ligand binding pocket colored in orange.
Actinonin is represented in ball-and-stick format and is seen
through the molecular surface of each PDF. (D) Ball-and-stick
representation of the interaction network around compound 21.
The metal cation is shown as a grey sphere.
(EPS)
Figure S10
Poorly active versions of AtPDF are in a closed
conformation incompatible with actinonin binding. (A) Free and
close AtPDF were superimposed as in Figure 1C and are figured in
brown and yellow, respectively. Both the G41M (chain B, shown
in orange) and the free C-deleted weakly active AtPDF versions
([63], colored in purple, PDB entry code 3CPM) were superim-
posed, to the two structures, showing that they both fit better to the
ligand-bound full-length close form than to the free open form, but
that the closure is further pronounced, burying the entrance to a
ligand. (B) Close-up showing that the shape of the S1’ pocket of the
poorly active closed versions make it poorly available to P1’
recognition (see circled Ile142 and Ile130 side chains).
(EPS)
Table S1
Catalytic properties of AtPDF. Nm, not measurable;
ND, not determined; WT, is wild-type. aKinetic constants were
determined using the coupled assay as indicated in Materials and
Methods with substrate Fo-Met-Ala-Ser, in the presence of 100
nM enzyme variant and 750 mM NiCl2, at 37uC. The relative
value of kcat/Km for wild-type AtPDF was set at 100%. bData
correspond to the binding constant of actinonin as obtained either
from ITC or from enzymatic analysis when indicated with an
asterisk. cData from Table S3. dGyration radii are from [83].
(DOC)
Table S2
Crystallographic data and refinement statistics. Values
in parentheses are for the outer resolution shell. aRsym (I) =
ShklSi|Ihkl,i 2 ,Ihkl.|/ShklSi|Ihkl,i|, where ,Ihkl. is the mean
intensity of the multiple Ihkl,i observations for symmetry-related
reflections. bRwork = 1006(Shkl|Fobs 2 Fcalc|/Shkl|Fobs|). Rfree
is a test set including ,5% of the data. cPercentage of residues in
most-favored/additionally
allowed/generously
allowed/disal-
lowed regions of the Ramachandran plot. dCompound 21 was
added first, and actinonin afterwards.
(DOC)
Table S3
Kinetic parameters for inhibition of some AtPDF
variants by actinonin. The enzyme concentration used in the assay
was 100 nM. Prior to kinetic analysis for determination of KI*app
values, actinonin was incubated in the presence of each variant set
at the final concentration for 10 min at 37uC; kinetic assay was
started
by
adding
a
small
volume
of
the
substrate.
For
determination of KI, k5, and k6 values, actinonin was not pre-
incubated with enzyme and kinetic assay was started by adding the
enzyme.
(DOCX)
Movie S1
Dynamics of actinonin binding to peptide deformylase
and closure of the active site.
(WMV)
Movie S2
Progressive motions of the main side chains at the
active site and final locking of the hydrogen bond.
(WMV)
Acknowledgments
We are strongly indebted to James Fraser and Tom Alber (University of
California, Berkeley, USA) for introducing us to Ringer before the release
of the freely available downloadable version. We thank Benoıˆt Gigant,
Virginie Gueguen-Chaignon, Solange Morera, and Philippe Peynot
(CNRS, Gif-sur-Yvette, France) for help with data processing and access
to the crystallization facilities. We also thank Magali Nicaise-Aumont
(IBBMC, Orsay, France), who performed the microcalorimetry experi-
ments. We are grateful to the staff of the European Synchrotron Radiation
Facility (ESRF) and SOLEIL beamlines for their help during data
collection.
Author Contributions
The author(s) have made the following declarations about their
contributions: Conceived and designed the experiments: SF CG TM.
Performed the experiments: AB SF. Analyzed the data: FD MD SF CG
TM. Contributed reagents/materials/analysis tools: IA MD CG TM.
Wrote the paper: CG TM.
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|
3M6Q
|
Crystal structure of Arabidopsis thaliana peptide deformylase 1B (AtPDF1B) G41Q mutant in complex with actinonin
|
Trapping Conformational States Along Ligand-Binding
Dynamics of Peptide Deformylase: The Impact of
Induced Fit on Enzyme Catalysis
Sonia Fieulaine1, Adrien Boularot1, Isabelle Artaud2,3, Michel Desmadril4,5, Fre´de´ric Dardel6,7, Thierry
Meinnel1*, Carmela Giglione1*
1 CNRS, ISV, UPR2355, Gif-sur-Yvette, France, 2 Universite´ Paris Descartes, UMR8601, Paris, France, 3 CNRS, UMR8601, Paris, France, 4 Universite´ Paris-Sud, IBBMC,
UMR8619, Orsay, France, 5 CNRS, IBBMC, UMR8619, Orsay, France, 6 Universite´ Paris Descartes, UMR8015, Paris, France, 7 CNRS, UMR8015, Paris, France
Abstract
For several decades, molecular recognition has been considered one of the most fundamental processes in biochemistry.
For enzymes, substrate binding is often coupled to conformational changes that alter the local environment of the active
site to align the reactive groups for efficient catalysis and to reach the transition state. Adaptive substrate recognition is a
well-known concept; however, it has been poorly characterized at a structural level because of its dynamic nature. Here, we
provide a detailed mechanism for an induced-fit process at atomic resolution. We take advantage of a slow, tight binding
inhibitor-enzyme system, actinonin-peptide deformylase. Crystal structures of the initial open state and final closed state
were solved, as well as those of several intermediate mimics captured during the process. Ligand-induced reshaping of a
hydrophobic pocket drives closure of the active site, which is finally ‘‘zipped up’’ by additional binding interactions.
Together with biochemical analyses, these data allow a coherent reconstruction of the sequence of events leading from the
encounter complex to the key-lock binding state of the enzyme. A ‘‘movie’’ that reconstructs this entire process can be
further extrapolated to catalysis.
Citation: Fieulaine S, Boularot A, Artaud I, Desmadril M, Dardel F, et al. (2011) Trapping Conformational States Along Ligand-Binding Dynamics of Peptide
Deformylase: The Impact of Induced Fit on Enzyme Catalysis. PLoS Biol 9(5): e1001066. doi:10.1371/journal.pbio.1001066
Academic Editor: Gregory A. Petsko, Brandeis University, United States of America
Received January 7, 2011; Accepted April 14, 2011; Published May 24, 2011
Copyright: 2011 Fieulaine et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by the Centre National de la Recherche Scientifique (CNRS, France), grant ANR-06-MIME-010-01 (Agence Nationale de la
Recherche, France), and grant #4920 from the Association pour la Recherche sur le Cancer (Villejuif, France). SF was partly supported by a postdoctoral fellowship
from the Fondation pour la Recherche Me´dicale (France). AB was supported by CNRS, France. The funders had no role in study design, data collection and
analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
Abbreviations: DSC, differential scanning calorimetry; Fo, formyl; PDF, peptide deformylase; r.m.s.d., root mean square deviation.
* E-mail: Thierry.Meinnel@isv.cnrs-gif.fr (TM); Carmela.Giglione@isv.cnrs-gif.fr (CG)
Introduction
Flexibility of proteins around their active site is a central feature
of molecular biochemistry [1–5]. Although this has been a central
concept
in
biochemistry
for
half
a
century,
the
detailed
mechanisms describing how the active enzyme conformation is
achieved have remained largely elusive, as a consequence of their
transient
nature.
Direct
structural
evidence
and/or
kinetic
analyses have only recently emerged [6–10]. Three classic
‘‘textbook’’ models are used to describe the formation of the
ligand-enzyme complex: (i) the Fischer’s ‘‘lock-and key’’ model, (ii)
the Koshland’s induced-fit model, and (iii) the selected-shift model
or conformational selection mechanism [6–8,11–13]. In the
Fischer’s ‘‘lock-and key’’ model, the conformations of free and
ligand-bound proteins are essentially the same. In the induced-fit
model, ligand binding induces a conformational change in the
protein, leading to the precise orientation of the catalytic groups
and implying the existence of initial molecular matches that
provide sufficient affinity prior to conformational adaptation [14].
In contrast, the selected-fit model assumes an equilibrium between
multiple conformational states, in which the ligand is able to select
and stabilize a complementary protein conformation. In this case,
the conformational change precedes ligand binding, in contrast to
the induced-fit model in which binding occurs first. The
conformational selection and/or induced-fit processes have been
shown to be involved in a number of enzymes [12,13,15,16]. For
several of these studies, conformational selection is proposed
because the experimental data support that, even in the absence of
the ligand, the enzyme samples multiple conformational states,
including the ligand-bound (active) state [6]. Although direct
structural evidence and/or kinetic analyses have provided clues
[6–8,12,13,16], how we can distinguish whether a protein binds its
ligand in an induced- or selected-fit mechanism remains critical
and often controversial.
The enzyme-inhibitor interaction is a form of molecular
recognition that is more amenable to investigation than the
enzyme-substrate interaction as there is no chemical transforma-
tion of the ligand during this process. In this context, slow, tight-
binding inhibition is an interesting interaction process, as it closely
mimics the substrate recognition process and has been shown to be
commonly involved in adaptive conformational changes [12,
17,18]. In slow, tight-binding inhibition, the degree of inhibition at
a fixed concentration of compound varies over time, leading to a
curvature of the reaction progress curve over time during which
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the uninhibited reaction progress curve is linear [19]. Indeed, the
slow, tight-binding inhibition is a two-step mechanism that
depends on the rate and strength of inhibitor interactions with
the enzyme. Binding of the inhibitor (I) to the enzyme (E) leads to
the rapid formation of a non-covalent enzyme-inhibitor complex
(E:I) followed by monomolecular slower step (k5) in which the E:I is
transformed into a more stable complex (E:I*) that relaxes and
dissociates at a very slow rate, mainly inferred by the k6 value
when k6,,k5,,k4, (Figure 1A; see also footnote f in Table 1).
Although only a few studies have investigated the mechanisms
of slow, tight-binding inhibitors, such molecules are favored for use
as
therapeutics,
as
they
usually
exhibit
unique
inhibitory
properties, including selective potency and long-lasting effects
[20–26]. Here, we explore the precise structural inhibitory
mechanism of actinonin (Figure 1A; [27]), which is a slow, tight-
binding inhibitor of peptide deformylase (PDF), a metal cation-
dependent enzyme [28,29]. The function of the active-site metal is
to activate the reactive water molecule involved in peptide
hydrolysis [30]. PDF is the first enzyme in the N-terminal
methionine excision pathway, an essential and ubiquitous process
that contributes to the diversity of N-terminal amino acids [31,32].
Actinonin is a natural product with antibiotic activity that inhibits
PDF by mimicking the structure of its natural substrates (nascent
peptide chains starting with Fo-Met-Aaa, where Fo is a formyl
group and Aaa is any amino acid) in their transition state
(Figure 1B). The transition state inhibitor actinonin, as well as
other structurally related inhibitors, has been shown to systemat-
ically exhibit a ‘‘slow-binding’’ inhibition behavior (Figure 1A),
regardless of the organism of origin of the PDF [29,33].
Using structural, biocomputing, and enzymatic analyses, we
were able to (i) reveal that the free enzyme is in an open
conformation and that actinonin induces transition of the enzyme
into a closed conformation; (ii) show that there is no evidence for
the occurrence of a closed conformation in the apostructure of the
open enzyme, which, together with detailed kinetic analyses,
makes the closed form fully compatible with an induced-fit model;
and (iii) identify the sequence of molecular events leading to the
final, bound, closed complex (E:I*). Moreover, using several
rationally designed point mutants of the enzyme, ligand-induced
intermediates, which mimic conformational states that normally
would not be expected to accumulate with the wild-type (WT)
enzyme, were trapped. These conformations recapitulate physical
states that the WT enzyme must pass through during its overall
transition from the apo-enzyme to the E:I* complex. ‘‘Freezing’’ of
ligand-induced intermediate states provides direct evidence for an
induced-fit mechanism and allows the reconstruction of a virtual
‘‘movie’’ that recapitulates this mechanism. Since PDF is one
example of an enzyme remaining active in the crystalline state and
because actinonin closely mimics the natural substrates bound to
PDF in the transition state as shown previously with the Escherichia
coli form (EcPDF; see Figure 1B) [34,35], we propose a model
suggesting that induced fit also contributes to efficient catalysis.
Results
Slow, Tight Binding of the Transition-State Analog
Actinonin to Peptide Deformylase
In the present study, at the atomic level we explored the precise
inhibitory mechanism of actinonin on Arabidopsis thaliana PDF1B
(AtPDF), a close eukaryotic homologue of EcPDF (Figure S1)
[36,37]. Measurements of the kinetic parameters of the second step
of the binding mechanism (k5) revealed a timescale in the 10-s range
(Table 1), which is consistent with the collective motion of a large
domain [4,5]. This finding is supported by NMR studies [38,39],
which showed that actinonin binding induces drastic changes in the
heteronuclear single quantum coherence (HSQC) spectrum of
EcPDF, since most resonances undergo significant shifts that affect a
large part of the structure [40,41]. The existence of alternative
conformational states of EcPDF is further supported by recent
biophysical studies [42]. Previously reported snapshots of a series of
different conformations of the enlarged and mobile loop—the so-
called CD loop—of the dimeric PDF from Leptospira interrogans PDF
(LiPDF) in the presence or absence of inhibitor led to the hypothesis
of the existence of an equilibrium between a closed and open form
of the CD-loop of PDF enzymes, suggesting a selected-shift model to
the authors [43]. Taken together, these data suggest that the binding
of actinonin to PDF is accompanied or preceded by conformational
changes within the enzyme. Paradoxically, this proposal has not
been currently supported by the available structural data. Indeed,
free and complexed crystal structures have provided no evidence for
any significant conformational change in PDF structure induced by
the binding of ligand [35,43–47].
Tight inhibition in the closed state is associated with the KI*
apparent equilibrium constant (Figure 1A). A KI* value (see Table 1
and Materials and Methods for the biochemical definition of KI*)
of 0.9 nM for actinonin could be measured for AtPDF; that is, a
value very similar to that obtained for bacterial PDFs, including
EcPDF and Bacillus stearothermophilus PDF2 (BsPDF2, Table 1).
Tightening of the initial encounter complex (E:I) resulted in a final
complex (E:I*) in which the potency of actinonin (KI/KI*) was
enhanced by more than two orders of magnitude and exhibited a
very slow off-rate (k6, Table 1). The dissociation constant value of
AtPDF for actinonin was also assessed using isothermal titration
calorimetry (ITC) experiments (Table S1 and Figure S2A). The
corresponding ITC titration curves (Figure S2A) are consistent
with a very strong affinity of the ligand for the enzyme [48],
enabling us to determine an accurate Kd. Moreover, these studies
generated values similar to those measured by other means for
AtPDF and EcPDF [42,49].
Author Summary
The notion of induced fit when a protein binds its ligand—
like a glove adapting to the shape of a hand—is a central
concept of structural biochemistry introduced over 50
years ago. A detailed molecular demonstration of this
phenomenon has eluded biochemists, however, largely
due to the difficulty of capturing the steps of this very
transient process: the ‘‘conformational change.’’ In this
study, we were able to see this process by using X-ray
diffraction to determine more than 10 distinct structures
adopted by a single enzyme when it binds a ligand. To do
this, we took advantage of the ‘‘slow, tight-binding’’ of a
potent inhibitor to its specific target enzyme to trap
intermediates in the binding process, which allowed us to
monitor the action of an enzyme in real-time at atomic
resolution. We showed the kinetics of the conformational
change from an initial open state, including the encounter
complex, to the final closed state of the enzyme. From
these
data
and
other
biochemical
and
biophysical
analyses, we make a coherent causal reconstruction of
the sequence of events leading to inhibition of the
enzyme’s activity. We also generated a movie that
reconstructs the sequence of events during the encounter.
Our data provide new insights into how enzymes achieve a
catalytically competent conformation in which the reactive
groups are brought into close proximity, resulting in
catalysis.
The Dynamics of Induced Fit at High Resolution
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Ligand-Induced Conformational Closure of AtPDF in the
Crystalline State
Occurrence of a conformational change induced by drug binding
was visualized via the resolution of several crystal structure forms of
AtPDF, the free form and/or in a complex with actinonin (Table
S2). The data reveal a structural switch between the two forms that
can account for both the thermodynamic and kinetic data. The
enzyme was observed in two states, a novel open apo-form and a
closed, induced, actinonin-bound complex (Figure 1C). Binding of
actinonin resulted in a tightening of the active site through the
collective closure of the entire N-terminal portion of the protein
(strands b1, b2, and b3; helix a1; and CD-loop, see Movies S1 and
S2, Figure 1C, and Figure S1). The amplitude of the structural
change was maximal for Pro60 (Figure S1), the Ca of which was
shifted 4 A˚ upon actinonin binding. This collective movement
involved the formation of a ‘‘super b-sheet’’ as the result of the large
rearrangement of b-strands 4 and 5 relative to the rest of the
structure in which actinonin forms an additional strand bridging the
two b-sheets (b1 andb2) on either side of the active site (Figure 1D
and Figure S1B). As actinonin is a peptide-like compound (see
Introduction and Figure 1B), this behavior closely mimics what
occurs in the natural protein substrates of PDF, which also form this
strand-bridging interaction. This phenomenon also accounts for the
strong stabilization of the protein by actinonin, which was also
challenged by differential scanning calorimetry (DSC) experiments:
the Tm of AtPDF increased from 61uC to 81uC upon binding of the
inhibitor (Figure 1D, see also below).
Thus far, this closure of the enzyme induced by actinonin is part
of the rare structural evidence for the slow, tight-binding
mechanism at an atomic scale. The open state, which has never
been observed, was captured not only in the two molecules of the
asymmetric subunit but also in different crystals and under two
distinct crystallization conditions (Table S2 and Figure 2). All
r.m.s.d. values were smaller than 0.25 A˚ . The closure is very
unlikely to result from crystal packing constraints, as soaking the
apo-AtPDF crystals in a solution containing actinonin induced the
Figure 1. Slow, tight-binding inhibition of PDF by actinonin induces conformational change in the protein. (A) Inhibition by a two-step
mechanism, involving a tightening of the initial enzyme-inhibitor complex (E?I) to form a more stable complex (E?I*), with the chemical structure of
actinonin (I), the natural inhibitor of PDF enzymes (E). (B) Structures of EcPDF bound to actinonin (left) and to the transition state resulting from the
cleavage of its substrate, Fo-Met-Ala-Ser (right) [34,35]. (C) Superimposition of free and actinonin-bound AtPDF indicated in green and purple,
respectively. The three conserved motifs of the PDF enzymes family are indicated in orange and numbered I, II, and III. Molecules A of both models
were superimposed, resulting in an r.m.s.d. of 0.9 A˚ for 100% of the Ca. Left inset, close-up comparison of the open and closed forms figured in the
ribbon representation. (D) Baseline-corrected DSC thermograms of free and actinonin-bound WT AtPDF recorded under the same experimental
conditions.
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structural transition from the open to the closed state within the
crystals without cracking them or altering their diffracting power.
Thus, crystal packing is compatible with both states of the enzyme
(Figure S3). Therefore, the open structure most likely corresponds
to a stable state in solution.
The closed final conformation was identical to that previously
reported for PDF complexes obtained either with actinonin or with a
product of the reaction [34,35,44,50], indicating that this structure is
common for the ligands (compare Figures 1B and 2A, and Figure S4).
Hydrogen bonding was also conserved, especially the bond between
the backbone nitrogen of Ile42 (corresponding to Ile44 in EcPDF, see
Figure 1B and Figure S5A) and the alkyl carbonyl chain of actinonin,
which potently contributes to the formation of the super b-sheet
(Movie S2 and Figure S1B, see also below). Between the open and
closed states, the side chains of Ile42, Phe58, and Ile130 underwent
significant structural changes (Figure 3A and D and Figure S6),
corresponding to a hydophobic pocket rearrangement, with Ile42
being the most affected (Figure 3). Interestingly, Ile42 is the second
residue of the conserved active-site motif G41IGLAAXG (motif 1)
that was previously shown to be essential for activity [51].
To assess and visualize the differences between the two states, two
independent structural parameters were measured: the r.m.s.d.
value with respect to the open form and the aperture angle (dap),
which measures the angle made between the N- and C-domains
through three fixed-points, corresponding to the Ca of three
conserved residues, each sitting in one of the three conserved motifs
(Figure 2A). The bi-dimensional graph of these two parameters is a
good representation of the closing motion snapshots (Figure 2B)
shown in Movie S1. With this tool at this stage, two states could be
defined: the closed (C) and open (O) states (Figure 2B).
Evidence for a Pure Induced-Fit Mechanism in the
Binding of Actinonin to AtPDF
Recent quantitative analyses of both conformational selection
and induced fit have led to an integrated continuum—a so-called
‘‘flux-description’’—of
these
two
limiting
mechanisms
[16].
According to this model, conformation selection tends to be
preferred at low ligand concentrations (mM range)—that is, using
detailed kinetic studies—whereas induced fit dominates at high
ligand and enzyme concentrations (mM range) obtained, for
instance, in NMR or crystallographic approaches. Structural
studies are most useful to reveal subpopulations of biological
significance.
We investigated the existence of lowly populated, alternative
conformations of apoPDF. To probe the occurrence of alternate
conformers in the crystalline state of PDF, the new Ringer
program is the most suitable investigation tool [52,53]. Ringer
searches for evidence of alternate rotamers by systematically
sampling electron density maps—free of model bias—around the
dihedral angles of protein side chains. Two independent WT open
datasets of the apoenzyme, including a high-resolution set (1.3 A˚ ),
were used in the analysis. Ringer analysis revealed the existence of
only one rotamer of most side chains of either molecule in the
asymmetric unit, including the three main residues primarily
involved in conformation change—that is, Ile 42, Phe58, and
Ile130
(Figure
4A).
Ringer
analysis
showed
evidence
for
unmodeled alternate conformers for very few residues, including
Ile121 and Phe87, or Phe119 to a much lesser extent (Figure S7).
There is therefore no evidence for the occurrence of a closed
conformation in the apostructure of AtPDF, supporting the
hypothesis that the conformational change was essentially induced
by the binding of actinonin rather than from conformational
selection among multiple states occurring in the crystalline state.
To further investigate the mechanism involved, we followed a
kinetic approach aimed at discriminating between induced fit and
population shift at low ligand concentrations (sub-mM range) [12].
The experimentally observed pseudo-first-order rate constant for
the approach to equilibrium between the free components and the
binary AtPDF-actinonin complex (kobs) was measured and plotted
as a function of actinonin concentration. This plot yielded a
hyperbolic saturation curve with a positive slope, as fully expected
for a pure induced-fit mechanism (Figure 4B and C). In contrast, if
the enzyme sampled two or more conformational states, the curve
would imply that the value of kobs decreases with increasing ligand
concentration (see, for instance, curve C in Figure 1 in [12]). The
same conclusion can be reached for EcPDF and BsPDF2
(Figure 4B and C) and was already reported by others for S.
aureus PDF [29].
Together, these data indicate that a pure induced-fit mechanism
triggered by the binding of actinonin appears to direct the
conformational change both in solution and in the crystalline state.
Single Variants at Gly41 Exhibit Strongly Reduced
Actinonin-Binding Potency and Catalytic Efficiency
When dealing with an induced-fit mechanism, knowledge of the
initial O and final C state is crucial but does not provide direct
information on the position of actinonin in the encounter complex
or on the sequential mechanism of the transition process. We
suspected that the conserved glycine-rich motif 1 (G41IGLAAXQ)
could contribute to the flexibility required for the observed
structural transition. Evidence for such flexibility comes from
NMR analysis of EcPDF in which a few residues show exchange
cross-peaks of an additional, alternative form [38]. The most
strongly affected residues are Cys90, one of the metal ligands, its
neighbor Leu91, and both of the alanines within the above
conserved glycine-rich motif (Figure S1B), suggesting that EcPDF
undergoes conformational dynamics in a similar region.
To unravel the dynamics of the recognition process, we
surmised that it should be possible to freeze the conformational
Table 1. Comparison of the main kinetic and thermodynamic
parameters describing the inhibition of PDF by actinonin.
Parameter
AtPDFa
EcPDFa
BsPDF2a,b
KI (nM)d
140610
112610
185615
KI* (nM)c
0.960.5
1.360.2
2.960.8
KI/KI*
155615
86610
6467
k5 (s21) 6103d
6369
170620
7268
k6 (s21) 6104d
461
1962
1163
k4 (s21)e
140610
112610
185615
t1/2 (min)f
2965
661
1.160.2
aThe enzyme concentrations used in the assay were 100, 50, and 25 nM for
AtPDF, EcPDF, and BsPDF2, respectively.
bData from [49].
cPrior to kinetic analysis for determination of the KI* value, actinonin was
incubated at the final concentration in the presence of the studied enzyme set
for 10 min at 37uC. The kinetic assay was initiated by the addition of a small
volume of the substrate.
dFor determination of KI, k5, and k6 values, actinonin was not preincubated with
the enzyme. The kinetic assay was initiated by the addition of the enzyme.
ek4 corresponds to the kinetic constant of the dissociation of the primary
enzyme-actinonin complex. It is assumed that the rate of complex association
is diffusion-limited (see Table 7.3 in [19]), that is, k3—the kinetic constant of the
association of the primary enzyme-actinonin complex—is 109 M21.s21.
ft1/2 is 0.693(k4+k5+k6)/k4k6 (see case of induced fit and calculation in Table 1 of
[12]). In this case, t1/2,0.693/k6 because k6,,k5,,k4.
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Figure 2. Four distinct conformational states of PDF enzymes. (A) AtPDF and several other representative bacterial PDFs are superimposed. A
zoom is displayed on the right of the panel. Superimpositions were realized using ‘‘module superpose’’ in the CCP4i package and the ‘‘secondary
structure matching’’ tool. The extent of aperture/closure of PDF enzymes was assessed primarily by the measurement of the aperture angle (dap), the
angle made between the Ca of three strictly conserved residues (C, H, and I) of all PDFs, each characterizing a secondary crucial structure module of
the active site crevice, namely b4, a2, and b1 (see Figure S1C). Each single residue belongs to one of the three conserved motifs (motifs 2, 3, and 1,
respectively) and corresponds respectively to Cys91, His137, and Ile42 in AtPDF. The dap was measured in each case (see B). (B) The dap values
combined with those of the r.m.s.d. associated with the superimposition of the open structure of AtPDF allows the identification of four
conformational states: open (O), intermediate (I), closed (C), and super-closed (S). We compared AtPDF1B (this work and PDB CODE 3CPM; brown,
orange, and yellow in A and B; black in C), EcPDF (1BS7, free enzyme; 1BS6, with Met-Ala-Ser; 1G2A, with actinonin; magenta), BsPDF2 (1LQY, with
actinonin; green), LiPDF (1SV2, free; 1SZZ, with actinonin; red), and PfPDF (1JYM, free; blue).
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Figure 3. Effect of actinonin binding on the conformation of key residues in PDF. Conformation of key residues Ile42, Phe58, and Ile130 in
the different complexes: (A) in unbound WT AtPDF, (B and C) in the structure of G41Q and G41M actinonin-bound variants, respectively, and (D) of
actinonin-bound WT protein. In the final complex (D), a hydrogen bond is formed between actinonin and the peptidic bond, which links Gly41 and
Ile42. During the deformylation reaction, which is catalyzed by the PDF enzyme, the N-terminal formyl-methionine fits into the S1’ pocket. The
solvent-accessible surface of this pocket is represented here, and only the aliphatic chain of actinonin is shown, mimicking the N-terminal
methionine. (E) Free WT enzyme with the S1’ pocket shown open in two orientations (top and bottom). (F and G) S1’ pocket in the G41Q and G41M
variant structures, respectively, shown in two orientations (top and bottom). (H) After the complete conformational modifications of actinonin-bound
WT protein induced by actinonin binding, the S1’ pocket is shown closed in two orientations (top and bottom). (I) The four models are superimposed;
the ligand-binding site is magnified: unbound WT AtPDF; G41Q and G41M actinonin-bound enzyme; and WT actinonin-bound enzyme are indicated
in brown, red, orange, and yellow, respectively. Actinonin is indicated by lines. (J) A detailed view of the AtPDF ligand-binding site for all the
complexes, which are superimposed, as indicated in the same colors. Arrows indicate the direction of the closing movement within the enzyme, from
the open, unbound state to the closed, bound state.
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change along the pathway by introducing selected, minor
variations within the above-mentioned crucial residues involved
in the collective motion. In this respect, site-directed mutagenesis
of AtPDF was performed on Gly41, Ile42, and Ile130. Single
substitutions were made at Gly41 (G41A/Q/M), Ile42 (I42A/F/
N/W), and Ile130 (I130A/F), and the variants were purified and
characterized. These mutant proteins showed no change in overall
stability, as evidenced by DSC experiments (unpublished data).
However, two variants of G41, G41Q and G41M, showed
dramatic effects; the kcat/Km values were reduced by three orders
of magnitude due to large decreases in the kcat values compared to
the WT enzyme (Figure 5A and Table S1). The reduced kcat/Km
values suggest an altered ability of these variants to attain the final
enzyme-transition state complex and, as a result, to give rise to
possible states different from the final E:I* complex. Substitutions
at positions 42 and 130 only caused small reductions in the kcat
values (Figure 5A, Figure S2C, and Table S1). The actinonin-
binding potency of both G41 variants was also greatly reduced
(Table S1 and Figure S2B). The time-dependent inhibition by
actinonin of the most active variants was then studied (Table S3).
Figure 4. Evidence for an induced fit in crystalline and solution states of AtPDF. (A) Absence of evidence for alternative conformers in the
crystalline state of AtPDF. Ringer plots of electron density (r) versus x1 angle for representative residues of the 3D apo-structure of AtPDF. Data were
obtained with the 3M6O, 3PN2, and 3PN3 datasets (2.0 and 1.3 A˚ resolution, respectively, see Table S1). The secondary peaks in the Ile residues are
observed because Ile is a branched amino acid. To reveal an alternative conformation with Ile, three peaks should be observed. (B) kobs is a saturable
function of actinonin with various PDFs, including AtPDF. Data obtained for kobs, the experimentally observed pseudo-first-order rate constant for the
approach to equilibrium between the free components and the binary PDF-actinonin complex, were obtained at various concentrations of actinonin
in the presence of EcPDF, AtPDF, and BsPDF2. A direct plot is shown. Inset, time-course measurement of deformylation as a function of varying
actinonin concentrations. (C) Inverted plot of the data in panel B, which is expected to be a straight line if the kobs is ..k6 in the case of induced fit
[19]. The correlation coefficient of each line is 1.00, 0.99, and 1.00 for AtPDF, BsPDF2, and EcPDF, respectively, indicative of the accuracy of the
conclusion.
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The half-lives of the final complexes—as assessed by comparison
of the 1/k6 values—were always significantly smaller (Table S3),
suggesting that the conformational change induced by actinonin
binding still occurred, but the C state is destabilized relative to the
O state in the mutants compared to the WT. Accordingly,
actinonin strongly stabilized almost all of the variants; Tm was
increased by more than 20uC. This differs from the G41M and
G41Q variants, which both showed increases in the Tm of only
12uC, consistent with reduced binding potency (Table S1).
Conformational Changes of Gly41 Variants Are Affected
On-Pathway
The two most interesting variants, G41Q and G41M, could be
crystallized under the same conditions as the WT protein. In the
case of G41Q, the structure of the apo-protein did not show any
modifications compared to the WT structure and remained in an
O conformation (Figure 2B; ‘‘O’’ zone). In contrast, the 3D
structure of the G41M variant showed that the asymmetric unit
was composed of two molecules with distinct structures. One
molecule (chain A) is in the O state and is similar to the structures
of the WT and the G41Q variant (Figure 2B; zone ‘‘O’’). The
second molecule (chain B) is in a C state, closer to that observed
for the WT chain in the presence of actinonin (‘‘C’’), a so-called
‘‘superclosed’’ state (Figure 2B; zone ‘‘S’’), suggesting that the
substitution modified the equilibrium between the two states in
solution either (i) at the step of protein synthesis by providing two
conformers, the inter-conversions of which are blocked due to
steric hindrance brought by the new bulkier side-chain at position
41, or (ii) by dramatically unbalancing the free inter-conversion
between the O and S conformers towards the S state. Ringer
analysis indicates that in the free G41M variant, many residues
show evidence for unmodeled alternate conformers—including
positions 58, 42, and 130—in keeping with the second hypothesis.
For all variants of position G41, addition of actinonin to the
crystal (Figure 3 and Figure S6) induced a closure of the protein
within the crystal. Nevertheless, as expected from in silico graphic
modeling followed by energy minimization, the occurrence of a
bulky side chain at position 41 prevented the completion of the
closure in the presence of the ligand and, hence, the formation of
the hydrogen bond between the backbone nitrogen of Ile42 and
actinonin. This finding is consistent with the strongly reduced Tm
of the complex of the variants with actinonin compared to WT as
measured by DSC. Remarkably, both S and O forms of the G41M
apo-structures in the asymmetric unit of the crystal yielded a
unique intermediary structure (‘‘I’’ state) upon actinonin binding
(r.m.s.d. between the molecules is ,0.25 A˚ ; see also Figure 2B,
zone ‘‘I’’). In this case, it is likely that the induced-fit mechanism
drives the equilibrium by capturing only the O population and
closing it to an intermediary step, thus depleting the pool of O
conformers that is shifted sequentially back from the remaining
pool of S conformers and allows the complete binding of actinonin
to the enzyme.
In line with the rational design of the PDF mutants, the extent
of the structural differences suggests that the underlying motions
are dependent on the length of the side chain (Figure S8).
Together, these data account for the reduced catalytic rate, as the
hydrogen bond is strictly required for the substrate to be efficiently
cleaved by PDFs (Figure S8A) [54]. Therefore, from both
structural and kinetic analyses, each substitution most likely
reproduces intermediates along the pathway that lead to the
closure of PDF around its substrate (Figure S2B).
Conformational Changes of Gly41 Variants Recapitulate
Closing Intermediates
Analysis of the structures allows us to propose the following
sequence of atomic events (Figures 3 and 2B and Figure S6). To
name the various sites of the ligand and subsites of PDF, we will
use the usual nomenclature found in [55], which defines the
various binding pockets of a protease, where P1’ is the first side
chain at the C-terminal side of the cleavage site and its binding
pocket is S1’, also referred to as the hydrophobic pocket in the case
of PDF. First, actinonin aligns along the S1’ pocket to form the
encounter complex, which shifts the Ile130 side chain to avoid
steric hindrance in the S1’ pocket, promotes rotation of the Ile42
side chain, and finally rearranges the phenyl group of Phe58.
These
events
achieve an
optimal
hydrophobic
S1’
pocket
conformation (Figure 3), and the concomitant closure leads to
the formation of a hydrogen bond between the first carbonyl
group of actinonin and the backbone nitrogen of Ile42. The initial
N-O distance is reduced from 5 A˚ to 2.8 A˚ , which is an optimal
value for hydrogen bonding (Movie S2 and Figure S8B). Thus, the
primary driving force for the active site closure appears to be the
P1’:S1’ hydrophobic interaction. The C state is ultimately locked
by the super-b-sheet hydrogen bonds extending across the ligand,
including those involving Ile42. The DDGbinding value (2.2–
2.4 kcal/mol, Figure S8B), as calculated from the Kd values for
actinonin binding to wild-type (WT) and G41M and G41Q, is
consistent with the loss of a hydrogen bond that also contributes to
the conformational stability of the protein [56,57]. Thus, this bond
contributes to the major binding free energy difference between
the two complexes (3.1 kcal/mol; Figure S8B, Tables S1 and S3,
and [29]). Interestingly, the above DDGbinding values also correlate
with the DDGES values derived from the kcat/Km and kcat
measurements [19]. This dataset strongly correlates with the
Figure 5. Inhibition and enzymatic reactions progress through an induced fit pathway. (A) The catalytic parameters Km and kcat, for all
AtPDF variants are provided as a percentage of the wild-type values (WT). Detailed values are presented in Table S2. (B) Schematic model for
actinonin binding to AtPDF in favor of an induced-fit pathway. PDF might exist in at least two conformational states, open (O) or closed (C). The
relative abundance of each conformation would vary, depending on the enzyme type. With AtPDF, it is likely that the most abundant form is the O
one, which is the only form leading to a productive complex. The superclosed form (S) is likely to show reduced affinity for the ligand because of
steric occlusion of the active site. At the initial stage, the inhibitor (shown in red) binds to AtPDF (indicated in brown) in the O conformation. To reach
the final key-lock state (productive closed conformation, C), two major and extreme pathways can be used. According to the conformational selection
pathway, the inhibitor selects the C conformation. This pathway, which is represented by the dashed arrow, does not occur within the crystal. In
contrast, the G41Q and G41M mutants, by providing the structure of the enzyme in intermediate conformations (I), prove the existence of the so-
called encounter complex and confirm that the inhibitor binds to the enzyme when it is in the O conformation. The ligand-binding site is then
reorganized to yield the C enzyme conformation, that is, the key-lock state. Indeed, the inhibitor binds to the enzyme through the induced-fit
pathway. Each timescale was calculated using the data available in the text and corresponds to t1/2 values deduced from the calculation of 0.693/
(kinetic constant of interest). The kcat value (k2) was used to assess the timescale of catalysis in panel C, whereas, in (B), k4 assesses the first step of
inhibition, and k6 is used in the case of the slow step. For the SO conversion (left, B), the lifetime of the minor form of EcPDF was used to assess the
order of magnitude (see text and [38]). (C) Schematic model for the deformylation reaction catalyzed by PDF. Since actinonin is a pseudo-peptidic
inhibitor, it is likely that a peptidic substrate can bind to the PDF enzyme through an induced-fit pathway, as described in (B). The key-lock state
represents a transition state in which the N-formylated substrate is deformylated to yield the final reaction product.
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gyration and van der Waals radii of the side chain at position 41 as
well as the N-O distance between the first carbonyl group of
actinonin and the backbone nitrogen of Ile42 (Figure S8). These
results suggest that the capacity of both G41M and G41Q variants
to form the transition state is a consequence of their inability to
reach the fully closed state.
Thus, our study of the designed Gly41 mutant enzymes reveals
that, in addition to the initial and final states observed for the WT
enzyme, the conformations of the Gly41 variants correspond
indeed to on-pathway intermediates, thus providing snapshots
along the trajectory from the O to the C state of the enzyme
(Figures 2B and 3). The 3D structure of the variants in the absence
of ligand is similar to that of WT, and a strict correlation exists
between the completeness of the conformational change and both
binding potency and catalytic efficiency. This suggests that both
events require complete protein closure to generate a productive
complex.
The
strong
stabilization
of
AtPDF
by
actinonin
(Figure 1D) closely mimics what occurs with its natural substrates
when it reaches the transition state [34,58]. Indeed, as expected,
the enzyme facilitates the final C conformation by lowering its
final energy [6]. Optimal arrangement of the S1’ pocket (Figure 3)
proceeds
along
the
reaction
process
towards
the
final
C
conformation, triggering the alignment of reactive groups in an
optimal arrangement for ligand recognition. Upon binding,
actinonin alters the thermodynamic landscape for the structural
transition between the O and C states. This ligand is a potent
inhibitor because it can trigger the above sequence of events
similar to the substrate, but unlike the substrate, it is non-
hydrolyzable. Thus, by mimicking the transition state and being
non-hydrolyzable (Figure 1B), the final C complex is long lasting.
Ligand-Induced Conformational Closure Is Initially
Triggered by the Binding of the P1’ Group in the S1’
Pocket
Given the similarity between actinonin and natural substrate
binding, the very slow kinetics of inhibitor binding (10-s time-scale)
remains puzzling compared to the 10 ms required for catalysis
(deduced from the kcat). This finding could be explained as a
conformational effect during the formation of the hydrogen bond,
aligning the substrate as an additional beta-sheet and eventually
stabilizing the entire enzyme-ligand complex. The significantly
longer time needed to reach the most stable state compared to the
substrate would most likely be due to the presence of the flexible
and one carbon longer metal-binding group in actinonin (i.e.,
hydroxamate versus formyl, Figure 1B). This suggestion is in line
with the overall data obtained when we investigated more deeply
the role of the first carbonyl group of the ligand. This group is well
known
to
exert
a
crucial
effect
in
both
productive
and
unproductive ligand binding (i.e., substrate and inhibitor) [54].
In this respect, we studied the binding of compound 6b (Figure
S5B), a PDF ligand that does not exhibit a reactive group at this
position [49]. We observed that this compound binds strongly to
both EcPDF (KI* = 6366 nM) and AtPDF (KI* = 400635 nM)
but, unlike actinonin, does not display slow, tight binding as KI* =
KI. This impact on binding is consistent with the absence of the
hydrogen bond involving the first carbonyl group of the ligand.
The 3D structure of AtPDF was determined after soaking the
compound in crystals of the free, open AtPDF form. Upon binding,
6b induced a complete conformational change, identical to that
observed with actinonin (Figures 2B and 6A; ‘‘O’’ state). This
result further suggests that the conformational change is not
induced initially by the formation of this hydrogen bond and that
the encounter complex is primarily driven by the fit within the S1’
pocket. This also reveals that the timescale of the large
conformational change is several orders of magnitude faster than
the kinetics of slow binding and fully compatible with both the first
step of actinonin binding (k4 = 140 s21; see Table 1) and the
catalytic rate of the substrate (kcat = 37 s21; see Table 1 and Table
S3). The 3D structure also revealed that both the P1’ and the
hydroxamate groups are bound similarly to the corresponding
groups of actinonin (Figure 6B). As expected, no additional
bonding occurs, especially around the backbone nitrogen of Ile42
(Figure 6C).
Taken together, these data allow us to conclude that the
conformational change observed upon ligand binding is triggered
primarily by binding in the S1’ pocket. As revealed by the binding of
6b, the one carbon longer metal-binding group fits, immediately
upon recognition of the P1’ group, in the S1’ pocket and forms a
bidentate complex with the metal cation, mimicking the transition
state as a result. Thus, the active site is very confined and rigid due
to the presence and length of the hydroxamate group (compare
right and left panels in Figure 1B). As a result, compared to the
complex made with the substrate, it is likely that the formation of the
hydrogen bond involving the carbonyl of actinonin and the
backbone nitrogen of Ile42 becomes strongly rate-limiting (k5
= 0.044 s21; Table 1). Once this hydrogen link is locked, the
uncleavable bond, mimicking the labile formyl group at the
transition state, stabilizes the enzyme-inhibitor complex, making it
long-lasting (k6 = 0.0006 s21; Table 1) and providing a mechanistic
explanation for the slow-binding effect that involves both large and
fine conformational changes. The large conformational change is
similar to the one occurring with the substrate, whereas the second is
more subtle and locks the hydrogen bond involving the backbone
nitrogen of Ile42. The second step is rate-limiting with some
transition state analogs such as actinonin (Figure 5B and C).
Proper Positioning of the Carbonyl Group Is Required to
Stabilize the Complex at S1’
Compound 21 corresponds to another interesting derivative
designed to probe the impact of the peptide bond in PDF binding
[49]. In addition to the hydroxamate group, this compound
features both a hydrophobic benzyl group at P1’ and a reverse
peptide bond. Compound 21 shows modest but significant
inhibitory potency to AtPDF1B (KI* = 400637 nM), confirming
the crucial role of the peptide bond in PDF binding. After soaking
with crystals of apo-AtPDF, compound 21 could be detected in
high-resolution electron density maps (Figure S9A). Unlike 6b, 21
did not bind the active site of the enzyme but an alternative pocket
at the surface of the protein (Figure S9B). A docking study
performed with EcPDF had previously revealed this alternative
binding pocket (Figure S9C; [59]).
The aforementioned data indicate that the occurrence of a S1’-
binding group placed in the unfavorable context of a reverse
peptide bond does not stably promote binding at the active site of
AtPDF. Upon binding of 21, the 3D structure of both molecules of
the asymmetric unit remain in an O conformation (r.m.s.d.
,0.2 A˚ with respect to the apo-structures in the ‘‘O’’ state). This
finding suggests that only the binding of compounds entering the
S1’ pocket, such as actinonin or 6b, induces conformational
change, in keeping with the crucial role of the P1’ group if located
in the frame of a classic peptide bond. Moreover, we noticed that
the binding pocket of 21 was located on the rear side of the true
S1’ pocket and induced a weak modification of the P1’ hosting
platform (Figure S9D). Indeed, when crystals of the 21:AtPDF
complex were soaked in actinonin, the final 3D structure no longer
showed evidence of compound 21 occupancy greater than 5%.
Instead, this structure revealed both actinonin and closing of the
protein (Table S2). The r.m.s.d. between this structure and that
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obtained directly with actinonin was less than 0.2 A˚ ; the actinonin
position was virtually identical, indicating that the protein had
retained full capacity for binding actinonin and closing despite the
presence of compound 21. We conclude that actinonin does
compete with 21 because of the overlap at P1’ of AtPDF1B (Figure
S9C). As the actinonin S1’ subsite strongly mimics that of a true
substrate, this result also explains the inhibitory behavior of 21
towards AtPDF.
Discussion
Although PDF catalysis has been extensively studied and the
mechanism has been elucidated [34], how the enzyme achieves the
catalytically competent state remains unknown. Here, we provide
insight on how the enzyme might reach a catalytically competent
conformation, demonstrating that the reactive groups move into
proximity to promote catalysis (Figures 2B and 5C). We suggest
Figure 6. Effect of 6b binding on the conformation of key residues of PDF. Superimposition of free, 6b-, and actinonin-bound AtPDF
indicated in brown, red, and yellow, respectively. (A) Molecule A in the three models was superimposed, resulting in an r.m.s.d. of 0.9 A˚ for 100% of
the Ca. Actinonin is shown in yellow and 6b in red. (B) Conformation of key residues Ile42, Phe58, and Ile130 in the different complexes and in
unbound WT AtPDF. Actinonin is shown in yellow and 6b in red. (C) A detailed view of the AtPDF ligand-binding site for both actinonin and 6b
complexes, which are indicated by sticks and are superimposed. The two ligands are colored in pale and dark grey, respectively. The hydrogen bond
made by actinonin only is shown.
doi:10.1371/journal.pbio.1001066.g006
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that the motions of the catalytic centre starting with free ligand-
PDF favor a final configuration that is optimal for binding and/or
catalysis (Figures 1B, 2B, and 5B and C). In our model, we propose
that free PDF might exist in at least two conformational states, that
is, open (O) or super-closed (S). The relative abundance of each
conformation varies by enzyme type and incubation conditions,
explaining why both conformations have not been trapped thus
far. In the case of AtPDF, it is likely that the most abundant form
corresponds to an O state, which is the form that leads to a
productive complex. Indeed, in the NMR spectra for EcPDF, a
few residues show exchange cross-peaks from an additional,
alternative form [38]. The most strongly affected residues are
Cys90, one of the metal ligands, its neighbor Leu91, as well as
Ala47 and Ala48 on the facing strand. This suggests that EcPDF
exists in at least two conformations (‘‘S’’ and ‘‘C’’; see Figure 2B),
which undergo slow interconversion on the NMR timescale. The
3D structure of the major conformation (75%, lifetime 300 ms)
could be solved at high resolution, but the structure of the minor
form (25%, lifetime 100 ms), which exhibits very weak signals,
could
not
be
solved
[38].
This
conformation
appears
to
correspond to that of the complex obtained with the product of
the reaction (Met-Ala-Ser). A very similar situation—although
more balanced between the two states—appears to occur in the
case of variant G41M, suggesting that a mechanism involving
conformational selection followed by induced fit is a general model
for PDF and that AtPDF is a specific case where population shift
virtually does not occur as the free enzyme is completely in the O
conformation. This is also in line with data obtained with L.
interrogans PDF (LiPDF), which reveal conformers in both the S and
C states (see Figure 2B) and suggest a population-shift mechanism
[43]. It is interesting to note that LiPDF is a poorly active PDF
[60]. According to the representation shown in Figure 2B,
Plasmodium falciparum PDF (PfPDF), a poorly active PDF [61],
was retrieved only in the S state. Finally, weak decompaction of
the structure of Bacillus cereus and Staphylococcus aureus PDFs in the
presence of actinonin have been described [45,46]. These
examples suggest that the enzyme is trapped in the S conformer
in the free state and converts to the C conformer when bound to
actinonin, suggesting that the S conformer is overrepresented in
solution compared to the O state, unlike AtPDF.
This study of AtPDF—including 10 different crystal structures of
apo- and complexed enzyme variants—reveals the 3D structure of
a PDF in at least four distinct states. This includes the O form, the
occurrence of which is crucial for catalysis, as it is the active form.
Here, we propose that the transition from the O to the C state is
directly induced by the ligand. Indeed, the O form, which is
captured in the crystal, undergoes closure directly upon ligand
binding in our soaking experiments. Progression to this closure
involves intermediary states (‘‘I’’) similar to those observed with
variants G41Q and G41M in the presence of actinonin (see
Figure 2B). Extrapolating the situation to catalysis, which occurs in
the crystalline states of PDF, it is likely that hydrolysis of the
substrate frees the enzyme in its S state, which in turn needs to
open to accommodate a new substrate (Figures 2B and 5C). This is
well illustrated in the 3D structure of EcPDF complexed with a
product of the reaction, obtained after co-crystallization of the
enzyme with the substrate in a closed conformation [34]. The S
free form is likely to exhibit a slower on-rate for the ligand (k3)
compared to the O form because of steric occlusion of the active
site (Figure S10). In support of this hypothesis, recent data show
that the 3D structure of a C-terminally truncated, poorly active
version of AtPDF is in the C conformation in the unbound state,
although crystallized under conditions identical to ours [62,63].
This structure is similar to that of chain B, one of the two
molecules of the asymmetric subunit of variant G41M (Figure 2B).
This suggests that alterations remote from the active site
significantly unbalance the equilibrium between the two conform-
ers, thus altering the efficiency of the reaction (Figure 5C). As the S
version corresponds to a significantly less active version of AtPDF
compared to that reported in our present work, this further
confirms that, compared to the O state, the S state has a
significantly weaker propensity to bind substrate or a close mimic
ligand, such as actinonin. Comparison of the 3D structures of the
free-closed and the ligand-bound-closed forms reveals some
differences responsible for the slight steric reduction of the active
site of free-closed AtPDF1B with respect to that of the actinonin-
AtPDF1B complex (Figure S10A), including the side chain of Ile42
burying the S1’ binding pocket (Figure S10B). Overall, these data
suggest that an S form might exist under the free state but that it
would feature a k3 value with respect to the ligand that is
significantly weaker than that of the O form, which would strongly
slow down the reaction or the binding as a result.
With the interaction scheme proposed in our model (Figure 5B
and C), the ligand/substrate binds more easily to the O form and
induces the optimal conformation of the enzyme to reach the
transition state, thus allowing the reaction to be efficiently
catalyzed.
In
the
final
model
(Figure
5C),
there
is
both
conformational selection and induced fit subsequently involved
in line with the recently proposed existence of such mixed
mechanisms for other enzymes [15,16]. Nevertheless, in our model
(Figure 5C), we suggest that induced fit is the primary mechanism,
as it provides energy input from the ligand, which eventually drives
the enzyme towards the productive key-lock complex. Unambig-
uous distinction between the relative contributions of the two
mechanisms is deduced from the observation that kobs is a saturable
function of actinonin with various PDF, including EcPDF, BsPDF,
AtPDF (Figure 4B and C), and S. aureus PDF [12,16,29,49].
Using crystallographic reconstruction analysis involving enzyme
variants, motions of small mobile loops and movie reconstructions
of snapshots of catalytic events have been previously documented
[1–3,64–66], often by visualizing the binding of unnatural
inhibitors and not necessarily mimicking closely the substrate
and transition state as actinonin does [67,68]. However, only a few
examples make use of soaking conditions of a crystal to promote
the motion and show the importance of induced fit [1,69]. None of
these data show a motion of the amplitude revealed here with PDF
and a large stabilization of the complex involving the formation of
the four-stranded b-sheet superstructure and the entire N-domain
of the enzyme. Compared to previous crystallographic analyses,
our work integrates biophysical, computational, and kinetic
analyses to reconstruct the whole picture, allowing a better
understanding of the slow-binding mechanism.
While our work primarily focused on an induced-fit mechanism
of enzyme inhibition and catalysis, it should be emphasized that
this phenomenon is also applicable to the broader area of
receptor-ligand interactions. For example, in all cases where
conformational change mechanisms have been proposed for
kinase inhibitors without supporting experimental data [12,26],
further experimental work must be provided to clarify the precise
mechanism. We expect this will have important implications on
how one conducts future drug-discovery efforts against such
enzymes [70].
Materials and Methods
Protein Expression and Purification
Expression and purification of mature Arabidopsis thaliana PDF1B
and all variants (i.e., AtPDF) were derived from the previously
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described protocol [37]: the lysis supernatant after sonication was
applied on a Q-Sepharose column (GE Healthcare; buffers A and
B as described containing 5 mM NiCl2) followed by Superdex-75
chromatography (GE Healthcare) using buffer C consisting of
buffer A supplemented with 0.1 M NaCl. For crystallization
experiments, the protein was purified further. The sample was
concentrated on an Amicon Ultra-15 centrifugal filter unit
(Millipore Corp.) with a 5-kDa cut-off and applied to a MonoQ
HR5/5 column (GE Healthcare) previously equilibrated in buffer
A (50 mM Hepes, pH 7.5, and 5 mM NiCl2). Elution was
performed with a 50-mL gradient from 0% to 100% buffer B.
The buffer of the pooled purified AtPDF1B was exchanged using a
PD-10 desalting column (GE Healthcare) to yield a protein
solution in 50 mM Hepes, pH 7.5, 0.1 M NaCl, and 5 mM NiCl2
(buffer C). The protein was concentrated on an Amicon Ultra-15
centrifugal filter unit. The resulting AtPDF1B preparation was
frozen in aliquots and stored at 280uC (for crystallization
purposes) or diluted 2-fold in 100% glycerol and stored at
220uC (for enzymatic purposes). The typical yield was 5–10 mg
AtPDF per liter of culture. All purification procedures were
performed at 4uC. Samples of the collected fractions were
analyzed by SDS-PAGE on 12% acrylamide gels, and protein
concentrations were estimated from the calculated extinction
coefficients for each variant.
Site-directed mutagenesis of AtPDF sequence in plasmid
pQdef1bDN [36] was carried out using the QuickChange Site-
Directed Mutagenesis Kit (Stratagene).
Enzymology
Assay of PDF activity was coupled to formate dehydrogenase,
where the absorbance of NADH at 340 nm was measured at 37uC
as previously described [71]. For measurements of classical kinetic
parameters (i.e., Km and kcat), the reaction was initiated by addition
of the substrate Fo-Met-Ala-Ser to the mixture containing purified
enzyme in the presence of 1 mM NiCl2. The kinetics parameters
were derived from iterative non-linear least square calculations
using the Michaelis-Menten equation based on the experimental
data (Sigma-Plot; Kinetics module). For determination of kinetic
parameters related to actinonin, the reaction mixture contained
750 mM NiCl2. In some cases, the mixture containing PDF and
actinonin was incubated for 15 min at 37uC before kinetic
analysis, which was initiated by the addition of substrate. The
same protocol was used to determine the dissociation constant of
actinonin [KI* = k4/(k3+k3k5/k6)], but the initial reaction velocities
were measured with varying concentrations of Fo-Met-Ala-Ser and
actinonin. The data were then calculated according to the method
of Henderson, which can be used to determine the dissociation
constant
of
the tight-binding
competitive enzyme
inhibitor
[28,49,72] by varying both the inhibitor and substrate concentra-
tions. To determine KI, k5, and k6, the reaction was initiated by the
addition of enzyme as previously described [29,49]. KI*app
measurements were used for comparative studies of AtPDF
variants (Table S3) at a concentration of 2 mM substrate by
varying the concentration of actinonin. KI*app is the slope of the
v[Actinonin]/v0 line curve. kobs was fitted from the kinetic data without
preincubation with vI = vs + (v0 2 vs)e2kobst where vI is the observed
velocity at a given concentration of inhibitor I, v0 is the velocity,
and vs is the steady-state velocity [18]. From the set of values
obtained at various concentrations of I, k5 and k6 could be derived
using kobs = k6 + k5[I]/(KI + [I]). By choosing a set of values with
kobs..k6, 1/kobs
= 1/k5(KI/[I] +1) and 1/kobs
=
f(1/[I]) is
expected to be a straight line in case of induced fit whose positive
slope corresponds to 1/k5. k6 was derived from equation k6 = k5/
(KI/KI*21) [18,19].
Microcalorimetry
ITC experiments were performed using a VP-ITC isothermal
titration calorimeter (Microcal Corp.). Experiments were per-
formed at 37uC. For each experiment, injections of 10 mL
actinonin (180 mM) were added using a computer-controlled
300 mL microsyringe at intervals of 240 s into the Ni-AtPDF
variant solution (5 to 10 mM, cell volume = 2.1 mL) dissolved in
buffer C with stirring at 310 rpm. A theoretical titration curve was
fitted to the experimental data using the ORIGIN software
(Microcal). This software uses the relationship between the heat
generated after each injection and DHu (enthalpy change in kcal/
mol), KA (the association binding constant in M21), n (number of
binding sites per monomer), total protein concentration, and free
and total ligand concentrations. The thermal stability of the WT
and variants of Ni-AtPDF1B was studied by DSC using VP-DSC
calorimetry (Microcal Corp.). DSC measurements were made with
10 mM protein solutions in buffer C. The actinonin concentration
was 20 mM. The same buffer was used as a reference. All solutions
were degassed just before loading into the calorimeter. Scanning
was performed at 1uC/min. The temperature dependence of the
partial molar capacity (Cp) was expressed in kcal/K after
subtracting the buffer signal using Origin(R) software.
Crystallization and Soaking Experiments
Crystallization conditions were screened by a robot using the
sitting drop vapor diffusion method. Crystals were obtained and
optimized at 20uC with 15%–20% PEG-3350 and either 0.1 or
0.2 M zinc acetate. The drops were formed by mixing 2 mL of a
solution containing 2 to 4 mg/mL protein and 2 mL of the
crystallization solution. Crystals were soaked for 24 h by adding
actinonin to the crystallization drops at a final concentration of
5 mM. Cryoprotection was achieved by placing crystals for 30 s in
a solution that was composed of 20% PEG-3350 and 0.2 M zinc
acetate, supplemented with 5%, 10%, and 15% glycerol. Crystals
were then directly flash frozen in liquid nitrogen using cryoloops
(Hampton Research). Crystals were also grown under conditions
described for the C-terminally deleted, weakly active version of
AtPDF [63].
X-Ray Diffraction Data Collection
Data collections were performed at 100 K at the European
Synchrotron Radiation Facility (Grenoble, France) on station
ID29, FIP-BM30A, ID14-1, and ID23-2, and at SOLEIL (Gif-sur-
Yvette, France) on station PROXIMA1. In each case, a single
crystal was used to collect a complete dataset. Data were processed
and scaled using XDS software [73]. Two crystal forms were
encountered with different cell parameters. In each case, b
parameter was nearly equal to a, and data could be indexed into
two space groups, P212121 or P43212. The data are shown in Table
S2.
Structure Determination and Refinement
The
structure
of
free
AtPDF
was
solved
by
molecular
replacement with Phaser [74] followed by a rigid-body refinement
by CNS [75] using coordinates from the Plasmodium falciparum PDF
(PDB code 1RL4) [76] as a search model. The structures of
actinonin-bound proteins—that is, WT and mutants—were solved
using rigid-body refinement by CNS of the free AtPDF structure.
The ten final models were obtained by manual rebuilding using
TURBO-FRODO [77] and combined with refinement of only
calculated phases using CNS and Refmac [78] software. No non-
crystallographic symmetries were used. Quality control of the
three models was performed using the PROCHECK program
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[79]. To probe for alternative conformers, Ringer was used [53].
Ringer is a program to detect molecular motions by automatic X-
ray electron density sampling, and can be accessed at http://
ucxray.berkeley.edu/ringer.htm.
Accession Numbers
PDB codes for the PDF structures presented within this
manuscript are as follows: 3M6O, 3PN2, 3M6P, 3O3J, 3PN3,
3PN4, 3PN5, 3M6Q, 3PN6, and 3M6R. UniProtKB accession
numbers for other PDF studied are P0A6K3 (EcPDF) and O31410
(BsPDF).
Supporting Information
Figure S1
Alignment of PDF sequences and secondary structures.
(A) PDF1B from Arabidopsis thaliana (AtPDF1B) is compared with
bacterial type 1B (EcPDF and LiPDF), pathogenic protozoa
(PfPDF1B), eukaryotic mitochondrial PDF1A from A. thaliana
(AtPDF1A), and bacterial type 2 (BsPDF2). This figure was created
with ENDscript [80]. The sequence alignment was realized with the
algorithm muscle included in ENDscript, and modified according to
the superimposition of structures. The blue frames indicate
conserved residues, white characters in red boxes indicate strict
identity, and red characters in yellow boxes indicate homology. The
secondary structures at the top (a-helices, 310 helices, b-strands, and
b-turns are shown by medium squiggles, small squiggles, arrows,
and TT letters, respectively) were predicted by DSSP [81]. Relative
accessibility (acc) of subunit A is shown by a blue-colored bar below
sequence. White is buried, cyan is intermediate, and blue with red
borders is highly exposed. A red box means that relative accessibility
is not calculated for the residue, because it is truncated. Hydropathy
(hyd) is calculated from the sequence according to [82]. It is shown
by a second bar below accessibility: pink is hydrophobic, grey is
intermediate, and cyan is hydrophilic. Motifs 1 (41GwGwAAXQ48),
2 (89EGCLS93), and 3 (133HEwDH137), where w is a hydrophobic
amino acid, are labeled by red stars below the sequence alignment.
To simplify the nomenclature, AtPDF1B is referred to as AtPDF
throughout the text. (B) Topology cartoon of AtPDF, free (left) or
actinonin bound (right), in the same color code as (A). Actinonin
(represented by the yellow arrow) binding to the ligand binding site
allows the linkage of the two distinct b-sheets into one single b-sheet,
by mimicking an additional b-strand. PDB sum (http://www.ebi.ac.
uk/thornton-srv/databases/pdbsum/) was used. (C) 3-D structure
of AtPDF is represented showing the position of the residues
discussed in the text, indicated in red.
(EPS)
Figure S2
Microcalorimetric titration of AtPDF with actinonin.
Data were obtained at 37uC by an automated sequence of 28
injections of 180 mM actinonin from a 300 ml syringe into the
reaction cell, which contain 9.85 mM AtPDF. The volume of each
reaction was 10 ml, and injections were made at 240 s intervals.
Top, raw data from the titration. Each peak corresponds to the
injection. Bottom, the peaks in the upper panel were integrated
with ORIGIN software and the values were plotted versus
injection number. Each point corresponds to the heat in mcal
generated by the reaction upon each injection. The solid line is the
curve fit to the data by the Origin program. This fit yields values
for Kd. Experiments were done with wild type protein and others
variants, and gave similar raw data and curve fit. (A) WT; (B)
variant G41M; (C) variant I42W.
(EPS)
Figure S3
Binding of actinonin to AtPDF does barely modify the
crystal packing. (A) Crystal pack of the two complexes: open, free
complex (left) and bound to actinonin (right) (B). Non-crystallo-
graphic contacts into asymmetric unit are not modified by closing
movement of the protein due to actinonin binding, except for zinc
atom number 6. This metal ion is coordinated by side chains of
Asp40 and Glu63, and water molecules, Asp40 and Glu63 being
hydrogen bonded by side chain of Lys38 of the other subunit of
the asymmetric unit. With the closing movement of the protein
into the crystal, Ca of Asp40 shifted by 3.1 A˚ and its side chain
flipped by 90u. Therefore, it does no longer participate to the
coordination shell of this Zn2+ ion. However, it is still hydrogen
bonded by Lys38 from chain B.
(EPS)
Figure S4
Binding of actinonin to AtPDF closely mimics both
actinonin and product binding to EcPDF. Superimposition of
EcPDF and AtPDF bound to either actinonin (1LRU PDB code,
panel A) or Met-Ala-Ser (1BS6 PDB code, panel B), the product of
the
reaction.
The
r.m.s.d.
value
is
1.11
A˚
for
151
Ca
superimposed.
(EPS)
Figure S5
The ligand binding site of AtPDF. This picture shows
the residues of AtPDF that are in contact with actinonin (left) and
6b (right) according to the 3-D structure; this should be compared
to the similar scheme shown in Figure 1B for EcPDF.
(EPS)
Figure S6
Electronic densities of the moving side-chains and of
actinonin at the binding site in some variants of AtPDF. Actinonin
and selected residues (G/Q/M41, I42, F58, and I130) are drawn
in stick and are shown in their FO–FC electron density omit maps
contoured at 2s, in free wild-type AtPDF (two crystallization
conditions, WT1 and WT2), and ligand-bound WT (actinonin, 6b
and 21), G41Q, and G41M variants.
(EPS)
Figure S7
Only few residues show alternative conformation in
AtPDF. Alternative conformers in the crystalline state of AtPDF.
Ringer
plots
of
electron
density
(r)
versus
x1
angle
for
representative residues of the 3-D apostructure of AtPDF. Data
were obtained with the 3M6O dataset (see Table S1). The
secondary peaks in the Ile residues are observed because Ile is a
branched amino acid. To evidence an alternative conformation
with Ile, three peaks should be observed.
(EPS)
Figure S8
Impact of induced fit on the binding free energy of
actinonin depends on the capacity to stabilize a hydrogen bond
with PDF. (A) The gyration radii [83] of the side chain occurring
at position 41 is displayed with black squares and compared to the
kcat/Km values (grey bars). (B) The distance between the NH of I42
and the CO of actinonin was measured in each case. The
percentage of the distance required to make a hydrogen bond (2.8
A˚ ) is reported (dark squares). The difference of binding free energy
(DDGbinding) between the open, free state and the variants closed
complexes of the G41 variants are displayed as grey bars. The
values were calculated as follows. For the WT, it corresponds to
the RT ln(KI*/KI) value [29], where R is the ideal gas constant and
T is the temperature in Kelvin. RT is 0.616 kcal.mol21 at 37uC.
For the G41M and G41Q variants, the DDGbinding corresponds to
RT ln(KI-G41variant/KD-WT). The obtained values are similar to that
obtained if the kcat/Km substitutes the KD value in the calculation
(DDGbinding = RT ln(kcat/Km –G41variant/kcat/Km –WT).
(EPS)
Figure S9
Compound 21 does not bind AtPDF1B at S1’. (A) 21
is shown in ball-and-stick format in its FO–FC electron density omit
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May 2011 | Volume 9 | Issue 5 | e1001066
map contoured at 2s. (B) Binding site of 21 into AtPDF1B is
detailed. Red and blue residues indicate residues that accommo-
date
the
‘‘phenylalanine’’
and
‘‘trimethyl’’
groups
of
21,
respectively. (C) Overall view of 21 binding site (left). Molecular
surface of AtPDF is represented, as well as 21 in ball-and-stick
format. Residues belonging to the 21 binding pocket are colored in
orange. For comparison, molecular surface of EcPDF (PDB code
1G2A) in the same orientation is also represented, with residues
forming the new ligand binding pocket colored in orange.
Actinonin is represented in ball-and-stick format and is seen
through the molecular surface of each PDF. (D) Ball-and-stick
representation of the interaction network around compound 21.
The metal cation is shown as a grey sphere.
(EPS)
Figure S10
Poorly active versions of AtPDF are in a closed
conformation incompatible with actinonin binding. (A) Free and
close AtPDF were superimposed as in Figure 1C and are figured in
brown and yellow, respectively. Both the G41M (chain B, shown
in orange) and the free C-deleted weakly active AtPDF versions
([63], colored in purple, PDB entry code 3CPM) were superim-
posed, to the two structures, showing that they both fit better to the
ligand-bound full-length close form than to the free open form, but
that the closure is further pronounced, burying the entrance to a
ligand. (B) Close-up showing that the shape of the S1’ pocket of the
poorly active closed versions make it poorly available to P1’
recognition (see circled Ile142 and Ile130 side chains).
(EPS)
Table S1
Catalytic properties of AtPDF. Nm, not measurable;
ND, not determined; WT, is wild-type. aKinetic constants were
determined using the coupled assay as indicated in Materials and
Methods with substrate Fo-Met-Ala-Ser, in the presence of 100
nM enzyme variant and 750 mM NiCl2, at 37uC. The relative
value of kcat/Km for wild-type AtPDF was set at 100%. bData
correspond to the binding constant of actinonin as obtained either
from ITC or from enzymatic analysis when indicated with an
asterisk. cData from Table S3. dGyration radii are from [83].
(DOC)
Table S2
Crystallographic data and refinement statistics. Values
in parentheses are for the outer resolution shell. aRsym (I) =
ShklSi|Ihkl,i 2 ,Ihkl.|/ShklSi|Ihkl,i|, where ,Ihkl. is the mean
intensity of the multiple Ihkl,i observations for symmetry-related
reflections. bRwork = 1006(Shkl|Fobs 2 Fcalc|/Shkl|Fobs|). Rfree
is a test set including ,5% of the data. cPercentage of residues in
most-favored/additionally
allowed/generously
allowed/disal-
lowed regions of the Ramachandran plot. dCompound 21 was
added first, and actinonin afterwards.
(DOC)
Table S3
Kinetic parameters for inhibition of some AtPDF
variants by actinonin. The enzyme concentration used in the assay
was 100 nM. Prior to kinetic analysis for determination of KI*app
values, actinonin was incubated in the presence of each variant set
at the final concentration for 10 min at 37uC; kinetic assay was
started
by
adding
a
small
volume
of
the
substrate.
For
determination of KI, k5, and k6 values, actinonin was not pre-
incubated with enzyme and kinetic assay was started by adding the
enzyme.
(DOCX)
Movie S1
Dynamics of actinonin binding to peptide deformylase
and closure of the active site.
(WMV)
Movie S2
Progressive motions of the main side chains at the
active site and final locking of the hydrogen bond.
(WMV)
Acknowledgments
We are strongly indebted to James Fraser and Tom Alber (University of
California, Berkeley, USA) for introducing us to Ringer before the release
of the freely available downloadable version. We thank Benoıˆt Gigant,
Virginie Gueguen-Chaignon, Solange Morera, and Philippe Peynot
(CNRS, Gif-sur-Yvette, France) for help with data processing and access
to the crystallization facilities. We also thank Magali Nicaise-Aumont
(IBBMC, Orsay, France), who performed the microcalorimetry experi-
ments. We are grateful to the staff of the European Synchrotron Radiation
Facility (ESRF) and SOLEIL beamlines for their help during data
collection.
Author Contributions
The author(s) have made the following declarations about their
contributions: Conceived and designed the experiments: SF CG TM.
Performed the experiments: AB SF. Analyzed the data: FD MD SF CG
TM. Contributed reagents/materials/analysis tools: IA MD CG TM.
Wrote the paper: CG TM.
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|
3M6R
|
Crystal structure of Arabidopsis thaliana peptide deformylase 1B (AtPDF1B) G41M mutant in complex with actinonin
|
Trapping Conformational States Along Ligand-Binding
Dynamics of Peptide Deformylase: The Impact of
Induced Fit on Enzyme Catalysis
Sonia Fieulaine1, Adrien Boularot1, Isabelle Artaud2,3, Michel Desmadril4,5, Fre´de´ric Dardel6,7, Thierry
Meinnel1*, Carmela Giglione1*
1 CNRS, ISV, UPR2355, Gif-sur-Yvette, France, 2 Universite´ Paris Descartes, UMR8601, Paris, France, 3 CNRS, UMR8601, Paris, France, 4 Universite´ Paris-Sud, IBBMC,
UMR8619, Orsay, France, 5 CNRS, IBBMC, UMR8619, Orsay, France, 6 Universite´ Paris Descartes, UMR8015, Paris, France, 7 CNRS, UMR8015, Paris, France
Abstract
For several decades, molecular recognition has been considered one of the most fundamental processes in biochemistry.
For enzymes, substrate binding is often coupled to conformational changes that alter the local environment of the active
site to align the reactive groups for efficient catalysis and to reach the transition state. Adaptive substrate recognition is a
well-known concept; however, it has been poorly characterized at a structural level because of its dynamic nature. Here, we
provide a detailed mechanism for an induced-fit process at atomic resolution. We take advantage of a slow, tight binding
inhibitor-enzyme system, actinonin-peptide deformylase. Crystal structures of the initial open state and final closed state
were solved, as well as those of several intermediate mimics captured during the process. Ligand-induced reshaping of a
hydrophobic pocket drives closure of the active site, which is finally ‘‘zipped up’’ by additional binding interactions.
Together with biochemical analyses, these data allow a coherent reconstruction of the sequence of events leading from the
encounter complex to the key-lock binding state of the enzyme. A ‘‘movie’’ that reconstructs this entire process can be
further extrapolated to catalysis.
Citation: Fieulaine S, Boularot A, Artaud I, Desmadril M, Dardel F, et al. (2011) Trapping Conformational States Along Ligand-Binding Dynamics of Peptide
Deformylase: The Impact of Induced Fit on Enzyme Catalysis. PLoS Biol 9(5): e1001066. doi:10.1371/journal.pbio.1001066
Academic Editor: Gregory A. Petsko, Brandeis University, United States of America
Received January 7, 2011; Accepted April 14, 2011; Published May 24, 2011
Copyright: 2011 Fieulaine et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by the Centre National de la Recherche Scientifique (CNRS, France), grant ANR-06-MIME-010-01 (Agence Nationale de la
Recherche, France), and grant #4920 from the Association pour la Recherche sur le Cancer (Villejuif, France). SF was partly supported by a postdoctoral fellowship
from the Fondation pour la Recherche Me´dicale (France). AB was supported by CNRS, France. The funders had no role in study design, data collection and
analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
Abbreviations: DSC, differential scanning calorimetry; Fo, formyl; PDF, peptide deformylase; r.m.s.d., root mean square deviation.
* E-mail: Thierry.Meinnel@isv.cnrs-gif.fr (TM); Carmela.Giglione@isv.cnrs-gif.fr (CG)
Introduction
Flexibility of proteins around their active site is a central feature
of molecular biochemistry [1–5]. Although this has been a central
concept
in
biochemistry
for
half
a
century,
the
detailed
mechanisms describing how the active enzyme conformation is
achieved have remained largely elusive, as a consequence of their
transient
nature.
Direct
structural
evidence
and/or
kinetic
analyses have only recently emerged [6–10]. Three classic
‘‘textbook’’ models are used to describe the formation of the
ligand-enzyme complex: (i) the Fischer’s ‘‘lock-and key’’ model, (ii)
the Koshland’s induced-fit model, and (iii) the selected-shift model
or conformational selection mechanism [6–8,11–13]. In the
Fischer’s ‘‘lock-and key’’ model, the conformations of free and
ligand-bound proteins are essentially the same. In the induced-fit
model, ligand binding induces a conformational change in the
protein, leading to the precise orientation of the catalytic groups
and implying the existence of initial molecular matches that
provide sufficient affinity prior to conformational adaptation [14].
In contrast, the selected-fit model assumes an equilibrium between
multiple conformational states, in which the ligand is able to select
and stabilize a complementary protein conformation. In this case,
the conformational change precedes ligand binding, in contrast to
the induced-fit model in which binding occurs first. The
conformational selection and/or induced-fit processes have been
shown to be involved in a number of enzymes [12,13,15,16]. For
several of these studies, conformational selection is proposed
because the experimental data support that, even in the absence of
the ligand, the enzyme samples multiple conformational states,
including the ligand-bound (active) state [6]. Although direct
structural evidence and/or kinetic analyses have provided clues
[6–8,12,13,16], how we can distinguish whether a protein binds its
ligand in an induced- or selected-fit mechanism remains critical
and often controversial.
The enzyme-inhibitor interaction is a form of molecular
recognition that is more amenable to investigation than the
enzyme-substrate interaction as there is no chemical transforma-
tion of the ligand during this process. In this context, slow, tight-
binding inhibition is an interesting interaction process, as it closely
mimics the substrate recognition process and has been shown to be
commonly involved in adaptive conformational changes [12,
17,18]. In slow, tight-binding inhibition, the degree of inhibition at
a fixed concentration of compound varies over time, leading to a
curvature of the reaction progress curve over time during which
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the uninhibited reaction progress curve is linear [19]. Indeed, the
slow, tight-binding inhibition is a two-step mechanism that
depends on the rate and strength of inhibitor interactions with
the enzyme. Binding of the inhibitor (I) to the enzyme (E) leads to
the rapid formation of a non-covalent enzyme-inhibitor complex
(E:I) followed by monomolecular slower step (k5) in which the E:I is
transformed into a more stable complex (E:I*) that relaxes and
dissociates at a very slow rate, mainly inferred by the k6 value
when k6,,k5,,k4, (Figure 1A; see also footnote f in Table 1).
Although only a few studies have investigated the mechanisms
of slow, tight-binding inhibitors, such molecules are favored for use
as
therapeutics,
as
they
usually
exhibit
unique
inhibitory
properties, including selective potency and long-lasting effects
[20–26]. Here, we explore the precise structural inhibitory
mechanism of actinonin (Figure 1A; [27]), which is a slow, tight-
binding inhibitor of peptide deformylase (PDF), a metal cation-
dependent enzyme [28,29]. The function of the active-site metal is
to activate the reactive water molecule involved in peptide
hydrolysis [30]. PDF is the first enzyme in the N-terminal
methionine excision pathway, an essential and ubiquitous process
that contributes to the diversity of N-terminal amino acids [31,32].
Actinonin is a natural product with antibiotic activity that inhibits
PDF by mimicking the structure of its natural substrates (nascent
peptide chains starting with Fo-Met-Aaa, where Fo is a formyl
group and Aaa is any amino acid) in their transition state
(Figure 1B). The transition state inhibitor actinonin, as well as
other structurally related inhibitors, has been shown to systemat-
ically exhibit a ‘‘slow-binding’’ inhibition behavior (Figure 1A),
regardless of the organism of origin of the PDF [29,33].
Using structural, biocomputing, and enzymatic analyses, we
were able to (i) reveal that the free enzyme is in an open
conformation and that actinonin induces transition of the enzyme
into a closed conformation; (ii) show that there is no evidence for
the occurrence of a closed conformation in the apostructure of the
open enzyme, which, together with detailed kinetic analyses,
makes the closed form fully compatible with an induced-fit model;
and (iii) identify the sequence of molecular events leading to the
final, bound, closed complex (E:I*). Moreover, using several
rationally designed point mutants of the enzyme, ligand-induced
intermediates, which mimic conformational states that normally
would not be expected to accumulate with the wild-type (WT)
enzyme, were trapped. These conformations recapitulate physical
states that the WT enzyme must pass through during its overall
transition from the apo-enzyme to the E:I* complex. ‘‘Freezing’’ of
ligand-induced intermediate states provides direct evidence for an
induced-fit mechanism and allows the reconstruction of a virtual
‘‘movie’’ that recapitulates this mechanism. Since PDF is one
example of an enzyme remaining active in the crystalline state and
because actinonin closely mimics the natural substrates bound to
PDF in the transition state as shown previously with the Escherichia
coli form (EcPDF; see Figure 1B) [34,35], we propose a model
suggesting that induced fit also contributes to efficient catalysis.
Results
Slow, Tight Binding of the Transition-State Analog
Actinonin to Peptide Deformylase
In the present study, at the atomic level we explored the precise
inhibitory mechanism of actinonin on Arabidopsis thaliana PDF1B
(AtPDF), a close eukaryotic homologue of EcPDF (Figure S1)
[36,37]. Measurements of the kinetic parameters of the second step
of the binding mechanism (k5) revealed a timescale in the 10-s range
(Table 1), which is consistent with the collective motion of a large
domain [4,5]. This finding is supported by NMR studies [38,39],
which showed that actinonin binding induces drastic changes in the
heteronuclear single quantum coherence (HSQC) spectrum of
EcPDF, since most resonances undergo significant shifts that affect a
large part of the structure [40,41]. The existence of alternative
conformational states of EcPDF is further supported by recent
biophysical studies [42]. Previously reported snapshots of a series of
different conformations of the enlarged and mobile loop—the so-
called CD loop—of the dimeric PDF from Leptospira interrogans PDF
(LiPDF) in the presence or absence of inhibitor led to the hypothesis
of the existence of an equilibrium between a closed and open form
of the CD-loop of PDF enzymes, suggesting a selected-shift model to
the authors [43]. Taken together, these data suggest that the binding
of actinonin to PDF is accompanied or preceded by conformational
changes within the enzyme. Paradoxically, this proposal has not
been currently supported by the available structural data. Indeed,
free and complexed crystal structures have provided no evidence for
any significant conformational change in PDF structure induced by
the binding of ligand [35,43–47].
Tight inhibition in the closed state is associated with the KI*
apparent equilibrium constant (Figure 1A). A KI* value (see Table 1
and Materials and Methods for the biochemical definition of KI*)
of 0.9 nM for actinonin could be measured for AtPDF; that is, a
value very similar to that obtained for bacterial PDFs, including
EcPDF and Bacillus stearothermophilus PDF2 (BsPDF2, Table 1).
Tightening of the initial encounter complex (E:I) resulted in a final
complex (E:I*) in which the potency of actinonin (KI/KI*) was
enhanced by more than two orders of magnitude and exhibited a
very slow off-rate (k6, Table 1). The dissociation constant value of
AtPDF for actinonin was also assessed using isothermal titration
calorimetry (ITC) experiments (Table S1 and Figure S2A). The
corresponding ITC titration curves (Figure S2A) are consistent
with a very strong affinity of the ligand for the enzyme [48],
enabling us to determine an accurate Kd. Moreover, these studies
generated values similar to those measured by other means for
AtPDF and EcPDF [42,49].
Author Summary
The notion of induced fit when a protein binds its ligand—
like a glove adapting to the shape of a hand—is a central
concept of structural biochemistry introduced over 50
years ago. A detailed molecular demonstration of this
phenomenon has eluded biochemists, however, largely
due to the difficulty of capturing the steps of this very
transient process: the ‘‘conformational change.’’ In this
study, we were able to see this process by using X-ray
diffraction to determine more than 10 distinct structures
adopted by a single enzyme when it binds a ligand. To do
this, we took advantage of the ‘‘slow, tight-binding’’ of a
potent inhibitor to its specific target enzyme to trap
intermediates in the binding process, which allowed us to
monitor the action of an enzyme in real-time at atomic
resolution. We showed the kinetics of the conformational
change from an initial open state, including the encounter
complex, to the final closed state of the enzyme. From
these
data
and
other
biochemical
and
biophysical
analyses, we make a coherent causal reconstruction of
the sequence of events leading to inhibition of the
enzyme’s activity. We also generated a movie that
reconstructs the sequence of events during the encounter.
Our data provide new insights into how enzymes achieve a
catalytically competent conformation in which the reactive
groups are brought into close proximity, resulting in
catalysis.
The Dynamics of Induced Fit at High Resolution
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Ligand-Induced Conformational Closure of AtPDF in the
Crystalline State
Occurrence of a conformational change induced by drug binding
was visualized via the resolution of several crystal structure forms of
AtPDF, the free form and/or in a complex with actinonin (Table
S2). The data reveal a structural switch between the two forms that
can account for both the thermodynamic and kinetic data. The
enzyme was observed in two states, a novel open apo-form and a
closed, induced, actinonin-bound complex (Figure 1C). Binding of
actinonin resulted in a tightening of the active site through the
collective closure of the entire N-terminal portion of the protein
(strands b1, b2, and b3; helix a1; and CD-loop, see Movies S1 and
S2, Figure 1C, and Figure S1). The amplitude of the structural
change was maximal for Pro60 (Figure S1), the Ca of which was
shifted 4 A˚ upon actinonin binding. This collective movement
involved the formation of a ‘‘super b-sheet’’ as the result of the large
rearrangement of b-strands 4 and 5 relative to the rest of the
structure in which actinonin forms an additional strand bridging the
two b-sheets (b1 andb2) on either side of the active site (Figure 1D
and Figure S1B). As actinonin is a peptide-like compound (see
Introduction and Figure 1B), this behavior closely mimics what
occurs in the natural protein substrates of PDF, which also form this
strand-bridging interaction. This phenomenon also accounts for the
strong stabilization of the protein by actinonin, which was also
challenged by differential scanning calorimetry (DSC) experiments:
the Tm of AtPDF increased from 61uC to 81uC upon binding of the
inhibitor (Figure 1D, see also below).
Thus far, this closure of the enzyme induced by actinonin is part
of the rare structural evidence for the slow, tight-binding
mechanism at an atomic scale. The open state, which has never
been observed, was captured not only in the two molecules of the
asymmetric subunit but also in different crystals and under two
distinct crystallization conditions (Table S2 and Figure 2). All
r.m.s.d. values were smaller than 0.25 A˚ . The closure is very
unlikely to result from crystal packing constraints, as soaking the
apo-AtPDF crystals in a solution containing actinonin induced the
Figure 1. Slow, tight-binding inhibition of PDF by actinonin induces conformational change in the protein. (A) Inhibition by a two-step
mechanism, involving a tightening of the initial enzyme-inhibitor complex (E?I) to form a more stable complex (E?I*), with the chemical structure of
actinonin (I), the natural inhibitor of PDF enzymes (E). (B) Structures of EcPDF bound to actinonin (left) and to the transition state resulting from the
cleavage of its substrate, Fo-Met-Ala-Ser (right) [34,35]. (C) Superimposition of free and actinonin-bound AtPDF indicated in green and purple,
respectively. The three conserved motifs of the PDF enzymes family are indicated in orange and numbered I, II, and III. Molecules A of both models
were superimposed, resulting in an r.m.s.d. of 0.9 A˚ for 100% of the Ca. Left inset, close-up comparison of the open and closed forms figured in the
ribbon representation. (D) Baseline-corrected DSC thermograms of free and actinonin-bound WT AtPDF recorded under the same experimental
conditions.
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structural transition from the open to the closed state within the
crystals without cracking them or altering their diffracting power.
Thus, crystal packing is compatible with both states of the enzyme
(Figure S3). Therefore, the open structure most likely corresponds
to a stable state in solution.
The closed final conformation was identical to that previously
reported for PDF complexes obtained either with actinonin or with a
product of the reaction [34,35,44,50], indicating that this structure is
common for the ligands (compare Figures 1B and 2A, and Figure S4).
Hydrogen bonding was also conserved, especially the bond between
the backbone nitrogen of Ile42 (corresponding to Ile44 in EcPDF, see
Figure 1B and Figure S5A) and the alkyl carbonyl chain of actinonin,
which potently contributes to the formation of the super b-sheet
(Movie S2 and Figure S1B, see also below). Between the open and
closed states, the side chains of Ile42, Phe58, and Ile130 underwent
significant structural changes (Figure 3A and D and Figure S6),
corresponding to a hydophobic pocket rearrangement, with Ile42
being the most affected (Figure 3). Interestingly, Ile42 is the second
residue of the conserved active-site motif G41IGLAAXG (motif 1)
that was previously shown to be essential for activity [51].
To assess and visualize the differences between the two states, two
independent structural parameters were measured: the r.m.s.d.
value with respect to the open form and the aperture angle (dap),
which measures the angle made between the N- and C-domains
through three fixed-points, corresponding to the Ca of three
conserved residues, each sitting in one of the three conserved motifs
(Figure 2A). The bi-dimensional graph of these two parameters is a
good representation of the closing motion snapshots (Figure 2B)
shown in Movie S1. With this tool at this stage, two states could be
defined: the closed (C) and open (O) states (Figure 2B).
Evidence for a Pure Induced-Fit Mechanism in the
Binding of Actinonin to AtPDF
Recent quantitative analyses of both conformational selection
and induced fit have led to an integrated continuum—a so-called
‘‘flux-description’’—of
these
two
limiting
mechanisms
[16].
According to this model, conformation selection tends to be
preferred at low ligand concentrations (mM range)—that is, using
detailed kinetic studies—whereas induced fit dominates at high
ligand and enzyme concentrations (mM range) obtained, for
instance, in NMR or crystallographic approaches. Structural
studies are most useful to reveal subpopulations of biological
significance.
We investigated the existence of lowly populated, alternative
conformations of apoPDF. To probe the occurrence of alternate
conformers in the crystalline state of PDF, the new Ringer
program is the most suitable investigation tool [52,53]. Ringer
searches for evidence of alternate rotamers by systematically
sampling electron density maps—free of model bias—around the
dihedral angles of protein side chains. Two independent WT open
datasets of the apoenzyme, including a high-resolution set (1.3 A˚ ),
were used in the analysis. Ringer analysis revealed the existence of
only one rotamer of most side chains of either molecule in the
asymmetric unit, including the three main residues primarily
involved in conformation change—that is, Ile 42, Phe58, and
Ile130
(Figure
4A).
Ringer
analysis
showed
evidence
for
unmodeled alternate conformers for very few residues, including
Ile121 and Phe87, or Phe119 to a much lesser extent (Figure S7).
There is therefore no evidence for the occurrence of a closed
conformation in the apostructure of AtPDF, supporting the
hypothesis that the conformational change was essentially induced
by the binding of actinonin rather than from conformational
selection among multiple states occurring in the crystalline state.
To further investigate the mechanism involved, we followed a
kinetic approach aimed at discriminating between induced fit and
population shift at low ligand concentrations (sub-mM range) [12].
The experimentally observed pseudo-first-order rate constant for
the approach to equilibrium between the free components and the
binary AtPDF-actinonin complex (kobs) was measured and plotted
as a function of actinonin concentration. This plot yielded a
hyperbolic saturation curve with a positive slope, as fully expected
for a pure induced-fit mechanism (Figure 4B and C). In contrast, if
the enzyme sampled two or more conformational states, the curve
would imply that the value of kobs decreases with increasing ligand
concentration (see, for instance, curve C in Figure 1 in [12]). The
same conclusion can be reached for EcPDF and BsPDF2
(Figure 4B and C) and was already reported by others for S.
aureus PDF [29].
Together, these data indicate that a pure induced-fit mechanism
triggered by the binding of actinonin appears to direct the
conformational change both in solution and in the crystalline state.
Single Variants at Gly41 Exhibit Strongly Reduced
Actinonin-Binding Potency and Catalytic Efficiency
When dealing with an induced-fit mechanism, knowledge of the
initial O and final C state is crucial but does not provide direct
information on the position of actinonin in the encounter complex
or on the sequential mechanism of the transition process. We
suspected that the conserved glycine-rich motif 1 (G41IGLAAXQ)
could contribute to the flexibility required for the observed
structural transition. Evidence for such flexibility comes from
NMR analysis of EcPDF in which a few residues show exchange
cross-peaks of an additional, alternative form [38]. The most
strongly affected residues are Cys90, one of the metal ligands, its
neighbor Leu91, and both of the alanines within the above
conserved glycine-rich motif (Figure S1B), suggesting that EcPDF
undergoes conformational dynamics in a similar region.
To unravel the dynamics of the recognition process, we
surmised that it should be possible to freeze the conformational
Table 1. Comparison of the main kinetic and thermodynamic
parameters describing the inhibition of PDF by actinonin.
Parameter
AtPDFa
EcPDFa
BsPDF2a,b
KI (nM)d
140610
112610
185615
KI* (nM)c
0.960.5
1.360.2
2.960.8
KI/KI*
155615
86610
6467
k5 (s21) 6103d
6369
170620
7268
k6 (s21) 6104d
461
1962
1163
k4 (s21)e
140610
112610
185615
t1/2 (min)f
2965
661
1.160.2
aThe enzyme concentrations used in the assay were 100, 50, and 25 nM for
AtPDF, EcPDF, and BsPDF2, respectively.
bData from [49].
cPrior to kinetic analysis for determination of the KI* value, actinonin was
incubated at the final concentration in the presence of the studied enzyme set
for 10 min at 37uC. The kinetic assay was initiated by the addition of a small
volume of the substrate.
dFor determination of KI, k5, and k6 values, actinonin was not preincubated with
the enzyme. The kinetic assay was initiated by the addition of the enzyme.
ek4 corresponds to the kinetic constant of the dissociation of the primary
enzyme-actinonin complex. It is assumed that the rate of complex association
is diffusion-limited (see Table 7.3 in [19]), that is, k3—the kinetic constant of the
association of the primary enzyme-actinonin complex—is 109 M21.s21.
ft1/2 is 0.693(k4+k5+k6)/k4k6 (see case of induced fit and calculation in Table 1 of
[12]). In this case, t1/2,0.693/k6 because k6,,k5,,k4.
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Figure 2. Four distinct conformational states of PDF enzymes. (A) AtPDF and several other representative bacterial PDFs are superimposed. A
zoom is displayed on the right of the panel. Superimpositions were realized using ‘‘module superpose’’ in the CCP4i package and the ‘‘secondary
structure matching’’ tool. The extent of aperture/closure of PDF enzymes was assessed primarily by the measurement of the aperture angle (dap), the
angle made between the Ca of three strictly conserved residues (C, H, and I) of all PDFs, each characterizing a secondary crucial structure module of
the active site crevice, namely b4, a2, and b1 (see Figure S1C). Each single residue belongs to one of the three conserved motifs (motifs 2, 3, and 1,
respectively) and corresponds respectively to Cys91, His137, and Ile42 in AtPDF. The dap was measured in each case (see B). (B) The dap values
combined with those of the r.m.s.d. associated with the superimposition of the open structure of AtPDF allows the identification of four
conformational states: open (O), intermediate (I), closed (C), and super-closed (S). We compared AtPDF1B (this work and PDB CODE 3CPM; brown,
orange, and yellow in A and B; black in C), EcPDF (1BS7, free enzyme; 1BS6, with Met-Ala-Ser; 1G2A, with actinonin; magenta), BsPDF2 (1LQY, with
actinonin; green), LiPDF (1SV2, free; 1SZZ, with actinonin; red), and PfPDF (1JYM, free; blue).
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Figure 3. Effect of actinonin binding on the conformation of key residues in PDF. Conformation of key residues Ile42, Phe58, and Ile130 in
the different complexes: (A) in unbound WT AtPDF, (B and C) in the structure of G41Q and G41M actinonin-bound variants, respectively, and (D) of
actinonin-bound WT protein. In the final complex (D), a hydrogen bond is formed between actinonin and the peptidic bond, which links Gly41 and
Ile42. During the deformylation reaction, which is catalyzed by the PDF enzyme, the N-terminal formyl-methionine fits into the S1’ pocket. The
solvent-accessible surface of this pocket is represented here, and only the aliphatic chain of actinonin is shown, mimicking the N-terminal
methionine. (E) Free WT enzyme with the S1’ pocket shown open in two orientations (top and bottom). (F and G) S1’ pocket in the G41Q and G41M
variant structures, respectively, shown in two orientations (top and bottom). (H) After the complete conformational modifications of actinonin-bound
WT protein induced by actinonin binding, the S1’ pocket is shown closed in two orientations (top and bottom). (I) The four models are superimposed;
the ligand-binding site is magnified: unbound WT AtPDF; G41Q and G41M actinonin-bound enzyme; and WT actinonin-bound enzyme are indicated
in brown, red, orange, and yellow, respectively. Actinonin is indicated by lines. (J) A detailed view of the AtPDF ligand-binding site for all the
complexes, which are superimposed, as indicated in the same colors. Arrows indicate the direction of the closing movement within the enzyme, from
the open, unbound state to the closed, bound state.
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change along the pathway by introducing selected, minor
variations within the above-mentioned crucial residues involved
in the collective motion. In this respect, site-directed mutagenesis
of AtPDF was performed on Gly41, Ile42, and Ile130. Single
substitutions were made at Gly41 (G41A/Q/M), Ile42 (I42A/F/
N/W), and Ile130 (I130A/F), and the variants were purified and
characterized. These mutant proteins showed no change in overall
stability, as evidenced by DSC experiments (unpublished data).
However, two variants of G41, G41Q and G41M, showed
dramatic effects; the kcat/Km values were reduced by three orders
of magnitude due to large decreases in the kcat values compared to
the WT enzyme (Figure 5A and Table S1). The reduced kcat/Km
values suggest an altered ability of these variants to attain the final
enzyme-transition state complex and, as a result, to give rise to
possible states different from the final E:I* complex. Substitutions
at positions 42 and 130 only caused small reductions in the kcat
values (Figure 5A, Figure S2C, and Table S1). The actinonin-
binding potency of both G41 variants was also greatly reduced
(Table S1 and Figure S2B). The time-dependent inhibition by
actinonin of the most active variants was then studied (Table S3).
Figure 4. Evidence for an induced fit in crystalline and solution states of AtPDF. (A) Absence of evidence for alternative conformers in the
crystalline state of AtPDF. Ringer plots of electron density (r) versus x1 angle for representative residues of the 3D apo-structure of AtPDF. Data were
obtained with the 3M6O, 3PN2, and 3PN3 datasets (2.0 and 1.3 A˚ resolution, respectively, see Table S1). The secondary peaks in the Ile residues are
observed because Ile is a branched amino acid. To reveal an alternative conformation with Ile, three peaks should be observed. (B) kobs is a saturable
function of actinonin with various PDFs, including AtPDF. Data obtained for kobs, the experimentally observed pseudo-first-order rate constant for the
approach to equilibrium between the free components and the binary PDF-actinonin complex, were obtained at various concentrations of actinonin
in the presence of EcPDF, AtPDF, and BsPDF2. A direct plot is shown. Inset, time-course measurement of deformylation as a function of varying
actinonin concentrations. (C) Inverted plot of the data in panel B, which is expected to be a straight line if the kobs is ..k6 in the case of induced fit
[19]. The correlation coefficient of each line is 1.00, 0.99, and 1.00 for AtPDF, BsPDF2, and EcPDF, respectively, indicative of the accuracy of the
conclusion.
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The half-lives of the final complexes—as assessed by comparison
of the 1/k6 values—were always significantly smaller (Table S3),
suggesting that the conformational change induced by actinonin
binding still occurred, but the C state is destabilized relative to the
O state in the mutants compared to the WT. Accordingly,
actinonin strongly stabilized almost all of the variants; Tm was
increased by more than 20uC. This differs from the G41M and
G41Q variants, which both showed increases in the Tm of only
12uC, consistent with reduced binding potency (Table S1).
Conformational Changes of Gly41 Variants Are Affected
On-Pathway
The two most interesting variants, G41Q and G41M, could be
crystallized under the same conditions as the WT protein. In the
case of G41Q, the structure of the apo-protein did not show any
modifications compared to the WT structure and remained in an
O conformation (Figure 2B; ‘‘O’’ zone). In contrast, the 3D
structure of the G41M variant showed that the asymmetric unit
was composed of two molecules with distinct structures. One
molecule (chain A) is in the O state and is similar to the structures
of the WT and the G41Q variant (Figure 2B; zone ‘‘O’’). The
second molecule (chain B) is in a C state, closer to that observed
for the WT chain in the presence of actinonin (‘‘C’’), a so-called
‘‘superclosed’’ state (Figure 2B; zone ‘‘S’’), suggesting that the
substitution modified the equilibrium between the two states in
solution either (i) at the step of protein synthesis by providing two
conformers, the inter-conversions of which are blocked due to
steric hindrance brought by the new bulkier side-chain at position
41, or (ii) by dramatically unbalancing the free inter-conversion
between the O and S conformers towards the S state. Ringer
analysis indicates that in the free G41M variant, many residues
show evidence for unmodeled alternate conformers—including
positions 58, 42, and 130—in keeping with the second hypothesis.
For all variants of position G41, addition of actinonin to the
crystal (Figure 3 and Figure S6) induced a closure of the protein
within the crystal. Nevertheless, as expected from in silico graphic
modeling followed by energy minimization, the occurrence of a
bulky side chain at position 41 prevented the completion of the
closure in the presence of the ligand and, hence, the formation of
the hydrogen bond between the backbone nitrogen of Ile42 and
actinonin. This finding is consistent with the strongly reduced Tm
of the complex of the variants with actinonin compared to WT as
measured by DSC. Remarkably, both S and O forms of the G41M
apo-structures in the asymmetric unit of the crystal yielded a
unique intermediary structure (‘‘I’’ state) upon actinonin binding
(r.m.s.d. between the molecules is ,0.25 A˚ ; see also Figure 2B,
zone ‘‘I’’). In this case, it is likely that the induced-fit mechanism
drives the equilibrium by capturing only the O population and
closing it to an intermediary step, thus depleting the pool of O
conformers that is shifted sequentially back from the remaining
pool of S conformers and allows the complete binding of actinonin
to the enzyme.
In line with the rational design of the PDF mutants, the extent
of the structural differences suggests that the underlying motions
are dependent on the length of the side chain (Figure S8).
Together, these data account for the reduced catalytic rate, as the
hydrogen bond is strictly required for the substrate to be efficiently
cleaved by PDFs (Figure S8A) [54]. Therefore, from both
structural and kinetic analyses, each substitution most likely
reproduces intermediates along the pathway that lead to the
closure of PDF around its substrate (Figure S2B).
Conformational Changes of Gly41 Variants Recapitulate
Closing Intermediates
Analysis of the structures allows us to propose the following
sequence of atomic events (Figures 3 and 2B and Figure S6). To
name the various sites of the ligand and subsites of PDF, we will
use the usual nomenclature found in [55], which defines the
various binding pockets of a protease, where P1’ is the first side
chain at the C-terminal side of the cleavage site and its binding
pocket is S1’, also referred to as the hydrophobic pocket in the case
of PDF. First, actinonin aligns along the S1’ pocket to form the
encounter complex, which shifts the Ile130 side chain to avoid
steric hindrance in the S1’ pocket, promotes rotation of the Ile42
side chain, and finally rearranges the phenyl group of Phe58.
These
events
achieve an
optimal
hydrophobic
S1’
pocket
conformation (Figure 3), and the concomitant closure leads to
the formation of a hydrogen bond between the first carbonyl
group of actinonin and the backbone nitrogen of Ile42. The initial
N-O distance is reduced from 5 A˚ to 2.8 A˚ , which is an optimal
value for hydrogen bonding (Movie S2 and Figure S8B). Thus, the
primary driving force for the active site closure appears to be the
P1’:S1’ hydrophobic interaction. The C state is ultimately locked
by the super-b-sheet hydrogen bonds extending across the ligand,
including those involving Ile42. The DDGbinding value (2.2–
2.4 kcal/mol, Figure S8B), as calculated from the Kd values for
actinonin binding to wild-type (WT) and G41M and G41Q, is
consistent with the loss of a hydrogen bond that also contributes to
the conformational stability of the protein [56,57]. Thus, this bond
contributes to the major binding free energy difference between
the two complexes (3.1 kcal/mol; Figure S8B, Tables S1 and S3,
and [29]). Interestingly, the above DDGbinding values also correlate
with the DDGES values derived from the kcat/Km and kcat
measurements [19]. This dataset strongly correlates with the
Figure 5. Inhibition and enzymatic reactions progress through an induced fit pathway. (A) The catalytic parameters Km and kcat, for all
AtPDF variants are provided as a percentage of the wild-type values (WT). Detailed values are presented in Table S2. (B) Schematic model for
actinonin binding to AtPDF in favor of an induced-fit pathway. PDF might exist in at least two conformational states, open (O) or closed (C). The
relative abundance of each conformation would vary, depending on the enzyme type. With AtPDF, it is likely that the most abundant form is the O
one, which is the only form leading to a productive complex. The superclosed form (S) is likely to show reduced affinity for the ligand because of
steric occlusion of the active site. At the initial stage, the inhibitor (shown in red) binds to AtPDF (indicated in brown) in the O conformation. To reach
the final key-lock state (productive closed conformation, C), two major and extreme pathways can be used. According to the conformational selection
pathway, the inhibitor selects the C conformation. This pathway, which is represented by the dashed arrow, does not occur within the crystal. In
contrast, the G41Q and G41M mutants, by providing the structure of the enzyme in intermediate conformations (I), prove the existence of the so-
called encounter complex and confirm that the inhibitor binds to the enzyme when it is in the O conformation. The ligand-binding site is then
reorganized to yield the C enzyme conformation, that is, the key-lock state. Indeed, the inhibitor binds to the enzyme through the induced-fit
pathway. Each timescale was calculated using the data available in the text and corresponds to t1/2 values deduced from the calculation of 0.693/
(kinetic constant of interest). The kcat value (k2) was used to assess the timescale of catalysis in panel C, whereas, in (B), k4 assesses the first step of
inhibition, and k6 is used in the case of the slow step. For the SO conversion (left, B), the lifetime of the minor form of EcPDF was used to assess the
order of magnitude (see text and [38]). (C) Schematic model for the deformylation reaction catalyzed by PDF. Since actinonin is a pseudo-peptidic
inhibitor, it is likely that a peptidic substrate can bind to the PDF enzyme through an induced-fit pathway, as described in (B). The key-lock state
represents a transition state in which the N-formylated substrate is deformylated to yield the final reaction product.
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gyration and van der Waals radii of the side chain at position 41 as
well as the N-O distance between the first carbonyl group of
actinonin and the backbone nitrogen of Ile42 (Figure S8). These
results suggest that the capacity of both G41M and G41Q variants
to form the transition state is a consequence of their inability to
reach the fully closed state.
Thus, our study of the designed Gly41 mutant enzymes reveals
that, in addition to the initial and final states observed for the WT
enzyme, the conformations of the Gly41 variants correspond
indeed to on-pathway intermediates, thus providing snapshots
along the trajectory from the O to the C state of the enzyme
(Figures 2B and 3). The 3D structure of the variants in the absence
of ligand is similar to that of WT, and a strict correlation exists
between the completeness of the conformational change and both
binding potency and catalytic efficiency. This suggests that both
events require complete protein closure to generate a productive
complex.
The
strong
stabilization
of
AtPDF
by
actinonin
(Figure 1D) closely mimics what occurs with its natural substrates
when it reaches the transition state [34,58]. Indeed, as expected,
the enzyme facilitates the final C conformation by lowering its
final energy [6]. Optimal arrangement of the S1’ pocket (Figure 3)
proceeds
along
the
reaction
process
towards
the
final
C
conformation, triggering the alignment of reactive groups in an
optimal arrangement for ligand recognition. Upon binding,
actinonin alters the thermodynamic landscape for the structural
transition between the O and C states. This ligand is a potent
inhibitor because it can trigger the above sequence of events
similar to the substrate, but unlike the substrate, it is non-
hydrolyzable. Thus, by mimicking the transition state and being
non-hydrolyzable (Figure 1B), the final C complex is long lasting.
Ligand-Induced Conformational Closure Is Initially
Triggered by the Binding of the P1’ Group in the S1’
Pocket
Given the similarity between actinonin and natural substrate
binding, the very slow kinetics of inhibitor binding (10-s time-scale)
remains puzzling compared to the 10 ms required for catalysis
(deduced from the kcat). This finding could be explained as a
conformational effect during the formation of the hydrogen bond,
aligning the substrate as an additional beta-sheet and eventually
stabilizing the entire enzyme-ligand complex. The significantly
longer time needed to reach the most stable state compared to the
substrate would most likely be due to the presence of the flexible
and one carbon longer metal-binding group in actinonin (i.e.,
hydroxamate versus formyl, Figure 1B). This suggestion is in line
with the overall data obtained when we investigated more deeply
the role of the first carbonyl group of the ligand. This group is well
known
to
exert
a
crucial
effect
in
both
productive
and
unproductive ligand binding (i.e., substrate and inhibitor) [54].
In this respect, we studied the binding of compound 6b (Figure
S5B), a PDF ligand that does not exhibit a reactive group at this
position [49]. We observed that this compound binds strongly to
both EcPDF (KI* = 6366 nM) and AtPDF (KI* = 400635 nM)
but, unlike actinonin, does not display slow, tight binding as KI* =
KI. This impact on binding is consistent with the absence of the
hydrogen bond involving the first carbonyl group of the ligand.
The 3D structure of AtPDF was determined after soaking the
compound in crystals of the free, open AtPDF form. Upon binding,
6b induced a complete conformational change, identical to that
observed with actinonin (Figures 2B and 6A; ‘‘O’’ state). This
result further suggests that the conformational change is not
induced initially by the formation of this hydrogen bond and that
the encounter complex is primarily driven by the fit within the S1’
pocket. This also reveals that the timescale of the large
conformational change is several orders of magnitude faster than
the kinetics of slow binding and fully compatible with both the first
step of actinonin binding (k4 = 140 s21; see Table 1) and the
catalytic rate of the substrate (kcat = 37 s21; see Table 1 and Table
S3). The 3D structure also revealed that both the P1’ and the
hydroxamate groups are bound similarly to the corresponding
groups of actinonin (Figure 6B). As expected, no additional
bonding occurs, especially around the backbone nitrogen of Ile42
(Figure 6C).
Taken together, these data allow us to conclude that the
conformational change observed upon ligand binding is triggered
primarily by binding in the S1’ pocket. As revealed by the binding of
6b, the one carbon longer metal-binding group fits, immediately
upon recognition of the P1’ group, in the S1’ pocket and forms a
bidentate complex with the metal cation, mimicking the transition
state as a result. Thus, the active site is very confined and rigid due
to the presence and length of the hydroxamate group (compare
right and left panels in Figure 1B). As a result, compared to the
complex made with the substrate, it is likely that the formation of the
hydrogen bond involving the carbonyl of actinonin and the
backbone nitrogen of Ile42 becomes strongly rate-limiting (k5
= 0.044 s21; Table 1). Once this hydrogen link is locked, the
uncleavable bond, mimicking the labile formyl group at the
transition state, stabilizes the enzyme-inhibitor complex, making it
long-lasting (k6 = 0.0006 s21; Table 1) and providing a mechanistic
explanation for the slow-binding effect that involves both large and
fine conformational changes. The large conformational change is
similar to the one occurring with the substrate, whereas the second is
more subtle and locks the hydrogen bond involving the backbone
nitrogen of Ile42. The second step is rate-limiting with some
transition state analogs such as actinonin (Figure 5B and C).
Proper Positioning of the Carbonyl Group Is Required to
Stabilize the Complex at S1’
Compound 21 corresponds to another interesting derivative
designed to probe the impact of the peptide bond in PDF binding
[49]. In addition to the hydroxamate group, this compound
features both a hydrophobic benzyl group at P1’ and a reverse
peptide bond. Compound 21 shows modest but significant
inhibitory potency to AtPDF1B (KI* = 400637 nM), confirming
the crucial role of the peptide bond in PDF binding. After soaking
with crystals of apo-AtPDF, compound 21 could be detected in
high-resolution electron density maps (Figure S9A). Unlike 6b, 21
did not bind the active site of the enzyme but an alternative pocket
at the surface of the protein (Figure S9B). A docking study
performed with EcPDF had previously revealed this alternative
binding pocket (Figure S9C; [59]).
The aforementioned data indicate that the occurrence of a S1’-
binding group placed in the unfavorable context of a reverse
peptide bond does not stably promote binding at the active site of
AtPDF. Upon binding of 21, the 3D structure of both molecules of
the asymmetric unit remain in an O conformation (r.m.s.d.
,0.2 A˚ with respect to the apo-structures in the ‘‘O’’ state). This
finding suggests that only the binding of compounds entering the
S1’ pocket, such as actinonin or 6b, induces conformational
change, in keeping with the crucial role of the P1’ group if located
in the frame of a classic peptide bond. Moreover, we noticed that
the binding pocket of 21 was located on the rear side of the true
S1’ pocket and induced a weak modification of the P1’ hosting
platform (Figure S9D). Indeed, when crystals of the 21:AtPDF
complex were soaked in actinonin, the final 3D structure no longer
showed evidence of compound 21 occupancy greater than 5%.
Instead, this structure revealed both actinonin and closing of the
protein (Table S2). The r.m.s.d. between this structure and that
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obtained directly with actinonin was less than 0.2 A˚ ; the actinonin
position was virtually identical, indicating that the protein had
retained full capacity for binding actinonin and closing despite the
presence of compound 21. We conclude that actinonin does
compete with 21 because of the overlap at P1’ of AtPDF1B (Figure
S9C). As the actinonin S1’ subsite strongly mimics that of a true
substrate, this result also explains the inhibitory behavior of 21
towards AtPDF.
Discussion
Although PDF catalysis has been extensively studied and the
mechanism has been elucidated [34], how the enzyme achieves the
catalytically competent state remains unknown. Here, we provide
insight on how the enzyme might reach a catalytically competent
conformation, demonstrating that the reactive groups move into
proximity to promote catalysis (Figures 2B and 5C). We suggest
Figure 6. Effect of 6b binding on the conformation of key residues of PDF. Superimposition of free, 6b-, and actinonin-bound AtPDF
indicated in brown, red, and yellow, respectively. (A) Molecule A in the three models was superimposed, resulting in an r.m.s.d. of 0.9 A˚ for 100% of
the Ca. Actinonin is shown in yellow and 6b in red. (B) Conformation of key residues Ile42, Phe58, and Ile130 in the different complexes and in
unbound WT AtPDF. Actinonin is shown in yellow and 6b in red. (C) A detailed view of the AtPDF ligand-binding site for both actinonin and 6b
complexes, which are indicated by sticks and are superimposed. The two ligands are colored in pale and dark grey, respectively. The hydrogen bond
made by actinonin only is shown.
doi:10.1371/journal.pbio.1001066.g006
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that the motions of the catalytic centre starting with free ligand-
PDF favor a final configuration that is optimal for binding and/or
catalysis (Figures 1B, 2B, and 5B and C). In our model, we propose
that free PDF might exist in at least two conformational states, that
is, open (O) or super-closed (S). The relative abundance of each
conformation varies by enzyme type and incubation conditions,
explaining why both conformations have not been trapped thus
far. In the case of AtPDF, it is likely that the most abundant form
corresponds to an O state, which is the form that leads to a
productive complex. Indeed, in the NMR spectra for EcPDF, a
few residues show exchange cross-peaks from an additional,
alternative form [38]. The most strongly affected residues are
Cys90, one of the metal ligands, its neighbor Leu91, as well as
Ala47 and Ala48 on the facing strand. This suggests that EcPDF
exists in at least two conformations (‘‘S’’ and ‘‘C’’; see Figure 2B),
which undergo slow interconversion on the NMR timescale. The
3D structure of the major conformation (75%, lifetime 300 ms)
could be solved at high resolution, but the structure of the minor
form (25%, lifetime 100 ms), which exhibits very weak signals,
could
not
be
solved
[38].
This
conformation
appears
to
correspond to that of the complex obtained with the product of
the reaction (Met-Ala-Ser). A very similar situation—although
more balanced between the two states—appears to occur in the
case of variant G41M, suggesting that a mechanism involving
conformational selection followed by induced fit is a general model
for PDF and that AtPDF is a specific case where population shift
virtually does not occur as the free enzyme is completely in the O
conformation. This is also in line with data obtained with L.
interrogans PDF (LiPDF), which reveal conformers in both the S and
C states (see Figure 2B) and suggest a population-shift mechanism
[43]. It is interesting to note that LiPDF is a poorly active PDF
[60]. According to the representation shown in Figure 2B,
Plasmodium falciparum PDF (PfPDF), a poorly active PDF [61],
was retrieved only in the S state. Finally, weak decompaction of
the structure of Bacillus cereus and Staphylococcus aureus PDFs in the
presence of actinonin have been described [45,46]. These
examples suggest that the enzyme is trapped in the S conformer
in the free state and converts to the C conformer when bound to
actinonin, suggesting that the S conformer is overrepresented in
solution compared to the O state, unlike AtPDF.
This study of AtPDF—including 10 different crystal structures of
apo- and complexed enzyme variants—reveals the 3D structure of
a PDF in at least four distinct states. This includes the O form, the
occurrence of which is crucial for catalysis, as it is the active form.
Here, we propose that the transition from the O to the C state is
directly induced by the ligand. Indeed, the O form, which is
captured in the crystal, undergoes closure directly upon ligand
binding in our soaking experiments. Progression to this closure
involves intermediary states (‘‘I’’) similar to those observed with
variants G41Q and G41M in the presence of actinonin (see
Figure 2B). Extrapolating the situation to catalysis, which occurs in
the crystalline states of PDF, it is likely that hydrolysis of the
substrate frees the enzyme in its S state, which in turn needs to
open to accommodate a new substrate (Figures 2B and 5C). This is
well illustrated in the 3D structure of EcPDF complexed with a
product of the reaction, obtained after co-crystallization of the
enzyme with the substrate in a closed conformation [34]. The S
free form is likely to exhibit a slower on-rate for the ligand (k3)
compared to the O form because of steric occlusion of the active
site (Figure S10). In support of this hypothesis, recent data show
that the 3D structure of a C-terminally truncated, poorly active
version of AtPDF is in the C conformation in the unbound state,
although crystallized under conditions identical to ours [62,63].
This structure is similar to that of chain B, one of the two
molecules of the asymmetric subunit of variant G41M (Figure 2B).
This suggests that alterations remote from the active site
significantly unbalance the equilibrium between the two conform-
ers, thus altering the efficiency of the reaction (Figure 5C). As the S
version corresponds to a significantly less active version of AtPDF
compared to that reported in our present work, this further
confirms that, compared to the O state, the S state has a
significantly weaker propensity to bind substrate or a close mimic
ligand, such as actinonin. Comparison of the 3D structures of the
free-closed and the ligand-bound-closed forms reveals some
differences responsible for the slight steric reduction of the active
site of free-closed AtPDF1B with respect to that of the actinonin-
AtPDF1B complex (Figure S10A), including the side chain of Ile42
burying the S1’ binding pocket (Figure S10B). Overall, these data
suggest that an S form might exist under the free state but that it
would feature a k3 value with respect to the ligand that is
significantly weaker than that of the O form, which would strongly
slow down the reaction or the binding as a result.
With the interaction scheme proposed in our model (Figure 5B
and C), the ligand/substrate binds more easily to the O form and
induces the optimal conformation of the enzyme to reach the
transition state, thus allowing the reaction to be efficiently
catalyzed.
In
the
final
model
(Figure
5C),
there
is
both
conformational selection and induced fit subsequently involved
in line with the recently proposed existence of such mixed
mechanisms for other enzymes [15,16]. Nevertheless, in our model
(Figure 5C), we suggest that induced fit is the primary mechanism,
as it provides energy input from the ligand, which eventually drives
the enzyme towards the productive key-lock complex. Unambig-
uous distinction between the relative contributions of the two
mechanisms is deduced from the observation that kobs is a saturable
function of actinonin with various PDF, including EcPDF, BsPDF,
AtPDF (Figure 4B and C), and S. aureus PDF [12,16,29,49].
Using crystallographic reconstruction analysis involving enzyme
variants, motions of small mobile loops and movie reconstructions
of snapshots of catalytic events have been previously documented
[1–3,64–66], often by visualizing the binding of unnatural
inhibitors and not necessarily mimicking closely the substrate
and transition state as actinonin does [67,68]. However, only a few
examples make use of soaking conditions of a crystal to promote
the motion and show the importance of induced fit [1,69]. None of
these data show a motion of the amplitude revealed here with PDF
and a large stabilization of the complex involving the formation of
the four-stranded b-sheet superstructure and the entire N-domain
of the enzyme. Compared to previous crystallographic analyses,
our work integrates biophysical, computational, and kinetic
analyses to reconstruct the whole picture, allowing a better
understanding of the slow-binding mechanism.
While our work primarily focused on an induced-fit mechanism
of enzyme inhibition and catalysis, it should be emphasized that
this phenomenon is also applicable to the broader area of
receptor-ligand interactions. For example, in all cases where
conformational change mechanisms have been proposed for
kinase inhibitors without supporting experimental data [12,26],
further experimental work must be provided to clarify the precise
mechanism. We expect this will have important implications on
how one conducts future drug-discovery efforts against such
enzymes [70].
Materials and Methods
Protein Expression and Purification
Expression and purification of mature Arabidopsis thaliana PDF1B
and all variants (i.e., AtPDF) were derived from the previously
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described protocol [37]: the lysis supernatant after sonication was
applied on a Q-Sepharose column (GE Healthcare; buffers A and
B as described containing 5 mM NiCl2) followed by Superdex-75
chromatography (GE Healthcare) using buffer C consisting of
buffer A supplemented with 0.1 M NaCl. For crystallization
experiments, the protein was purified further. The sample was
concentrated on an Amicon Ultra-15 centrifugal filter unit
(Millipore Corp.) with a 5-kDa cut-off and applied to a MonoQ
HR5/5 column (GE Healthcare) previously equilibrated in buffer
A (50 mM Hepes, pH 7.5, and 5 mM NiCl2). Elution was
performed with a 50-mL gradient from 0% to 100% buffer B.
The buffer of the pooled purified AtPDF1B was exchanged using a
PD-10 desalting column (GE Healthcare) to yield a protein
solution in 50 mM Hepes, pH 7.5, 0.1 M NaCl, and 5 mM NiCl2
(buffer C). The protein was concentrated on an Amicon Ultra-15
centrifugal filter unit. The resulting AtPDF1B preparation was
frozen in aliquots and stored at 280uC (for crystallization
purposes) or diluted 2-fold in 100% glycerol and stored at
220uC (for enzymatic purposes). The typical yield was 5–10 mg
AtPDF per liter of culture. All purification procedures were
performed at 4uC. Samples of the collected fractions were
analyzed by SDS-PAGE on 12% acrylamide gels, and protein
concentrations were estimated from the calculated extinction
coefficients for each variant.
Site-directed mutagenesis of AtPDF sequence in plasmid
pQdef1bDN [36] was carried out using the QuickChange Site-
Directed Mutagenesis Kit (Stratagene).
Enzymology
Assay of PDF activity was coupled to formate dehydrogenase,
where the absorbance of NADH at 340 nm was measured at 37uC
as previously described [71]. For measurements of classical kinetic
parameters (i.e., Km and kcat), the reaction was initiated by addition
of the substrate Fo-Met-Ala-Ser to the mixture containing purified
enzyme in the presence of 1 mM NiCl2. The kinetics parameters
were derived from iterative non-linear least square calculations
using the Michaelis-Menten equation based on the experimental
data (Sigma-Plot; Kinetics module). For determination of kinetic
parameters related to actinonin, the reaction mixture contained
750 mM NiCl2. In some cases, the mixture containing PDF and
actinonin was incubated for 15 min at 37uC before kinetic
analysis, which was initiated by the addition of substrate. The
same protocol was used to determine the dissociation constant of
actinonin [KI* = k4/(k3+k3k5/k6)], but the initial reaction velocities
were measured with varying concentrations of Fo-Met-Ala-Ser and
actinonin. The data were then calculated according to the method
of Henderson, which can be used to determine the dissociation
constant
of
the tight-binding
competitive enzyme
inhibitor
[28,49,72] by varying both the inhibitor and substrate concentra-
tions. To determine KI, k5, and k6, the reaction was initiated by the
addition of enzyme as previously described [29,49]. KI*app
measurements were used for comparative studies of AtPDF
variants (Table S3) at a concentration of 2 mM substrate by
varying the concentration of actinonin. KI*app is the slope of the
v[Actinonin]/v0 line curve. kobs was fitted from the kinetic data without
preincubation with vI = vs + (v0 2 vs)e2kobst where vI is the observed
velocity at a given concentration of inhibitor I, v0 is the velocity,
and vs is the steady-state velocity [18]. From the set of values
obtained at various concentrations of I, k5 and k6 could be derived
using kobs = k6 + k5[I]/(KI + [I]). By choosing a set of values with
kobs..k6, 1/kobs
= 1/k5(KI/[I] +1) and 1/kobs
=
f(1/[I]) is
expected to be a straight line in case of induced fit whose positive
slope corresponds to 1/k5. k6 was derived from equation k6 = k5/
(KI/KI*21) [18,19].
Microcalorimetry
ITC experiments were performed using a VP-ITC isothermal
titration calorimeter (Microcal Corp.). Experiments were per-
formed at 37uC. For each experiment, injections of 10 mL
actinonin (180 mM) were added using a computer-controlled
300 mL microsyringe at intervals of 240 s into the Ni-AtPDF
variant solution (5 to 10 mM, cell volume = 2.1 mL) dissolved in
buffer C with stirring at 310 rpm. A theoretical titration curve was
fitted to the experimental data using the ORIGIN software
(Microcal). This software uses the relationship between the heat
generated after each injection and DHu (enthalpy change in kcal/
mol), KA (the association binding constant in M21), n (number of
binding sites per monomer), total protein concentration, and free
and total ligand concentrations. The thermal stability of the WT
and variants of Ni-AtPDF1B was studied by DSC using VP-DSC
calorimetry (Microcal Corp.). DSC measurements were made with
10 mM protein solutions in buffer C. The actinonin concentration
was 20 mM. The same buffer was used as a reference. All solutions
were degassed just before loading into the calorimeter. Scanning
was performed at 1uC/min. The temperature dependence of the
partial molar capacity (Cp) was expressed in kcal/K after
subtracting the buffer signal using Origin(R) software.
Crystallization and Soaking Experiments
Crystallization conditions were screened by a robot using the
sitting drop vapor diffusion method. Crystals were obtained and
optimized at 20uC with 15%–20% PEG-3350 and either 0.1 or
0.2 M zinc acetate. The drops were formed by mixing 2 mL of a
solution containing 2 to 4 mg/mL protein and 2 mL of the
crystallization solution. Crystals were soaked for 24 h by adding
actinonin to the crystallization drops at a final concentration of
5 mM. Cryoprotection was achieved by placing crystals for 30 s in
a solution that was composed of 20% PEG-3350 and 0.2 M zinc
acetate, supplemented with 5%, 10%, and 15% glycerol. Crystals
were then directly flash frozen in liquid nitrogen using cryoloops
(Hampton Research). Crystals were also grown under conditions
described for the C-terminally deleted, weakly active version of
AtPDF [63].
X-Ray Diffraction Data Collection
Data collections were performed at 100 K at the European
Synchrotron Radiation Facility (Grenoble, France) on station
ID29, FIP-BM30A, ID14-1, and ID23-2, and at SOLEIL (Gif-sur-
Yvette, France) on station PROXIMA1. In each case, a single
crystal was used to collect a complete dataset. Data were processed
and scaled using XDS software [73]. Two crystal forms were
encountered with different cell parameters. In each case, b
parameter was nearly equal to a, and data could be indexed into
two space groups, P212121 or P43212. The data are shown in Table
S2.
Structure Determination and Refinement
The
structure
of
free
AtPDF
was
solved
by
molecular
replacement with Phaser [74] followed by a rigid-body refinement
by CNS [75] using coordinates from the Plasmodium falciparum PDF
(PDB code 1RL4) [76] as a search model. The structures of
actinonin-bound proteins—that is, WT and mutants—were solved
using rigid-body refinement by CNS of the free AtPDF structure.
The ten final models were obtained by manual rebuilding using
TURBO-FRODO [77] and combined with refinement of only
calculated phases using CNS and Refmac [78] software. No non-
crystallographic symmetries were used. Quality control of the
three models was performed using the PROCHECK program
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[79]. To probe for alternative conformers, Ringer was used [53].
Ringer is a program to detect molecular motions by automatic X-
ray electron density sampling, and can be accessed at http://
ucxray.berkeley.edu/ringer.htm.
Accession Numbers
PDB codes for the PDF structures presented within this
manuscript are as follows: 3M6O, 3PN2, 3M6P, 3O3J, 3PN3,
3PN4, 3PN5, 3M6Q, 3PN6, and 3M6R. UniProtKB accession
numbers for other PDF studied are P0A6K3 (EcPDF) and O31410
(BsPDF).
Supporting Information
Figure S1
Alignment of PDF sequences and secondary structures.
(A) PDF1B from Arabidopsis thaliana (AtPDF1B) is compared with
bacterial type 1B (EcPDF and LiPDF), pathogenic protozoa
(PfPDF1B), eukaryotic mitochondrial PDF1A from A. thaliana
(AtPDF1A), and bacterial type 2 (BsPDF2). This figure was created
with ENDscript [80]. The sequence alignment was realized with the
algorithm muscle included in ENDscript, and modified according to
the superimposition of structures. The blue frames indicate
conserved residues, white characters in red boxes indicate strict
identity, and red characters in yellow boxes indicate homology. The
secondary structures at the top (a-helices, 310 helices, b-strands, and
b-turns are shown by medium squiggles, small squiggles, arrows,
and TT letters, respectively) were predicted by DSSP [81]. Relative
accessibility (acc) of subunit A is shown by a blue-colored bar below
sequence. White is buried, cyan is intermediate, and blue with red
borders is highly exposed. A red box means that relative accessibility
is not calculated for the residue, because it is truncated. Hydropathy
(hyd) is calculated from the sequence according to [82]. It is shown
by a second bar below accessibility: pink is hydrophobic, grey is
intermediate, and cyan is hydrophilic. Motifs 1 (41GwGwAAXQ48),
2 (89EGCLS93), and 3 (133HEwDH137), where w is a hydrophobic
amino acid, are labeled by red stars below the sequence alignment.
To simplify the nomenclature, AtPDF1B is referred to as AtPDF
throughout the text. (B) Topology cartoon of AtPDF, free (left) or
actinonin bound (right), in the same color code as (A). Actinonin
(represented by the yellow arrow) binding to the ligand binding site
allows the linkage of the two distinct b-sheets into one single b-sheet,
by mimicking an additional b-strand. PDB sum (http://www.ebi.ac.
uk/thornton-srv/databases/pdbsum/) was used. (C) 3-D structure
of AtPDF is represented showing the position of the residues
discussed in the text, indicated in red.
(EPS)
Figure S2
Microcalorimetric titration of AtPDF with actinonin.
Data were obtained at 37uC by an automated sequence of 28
injections of 180 mM actinonin from a 300 ml syringe into the
reaction cell, which contain 9.85 mM AtPDF. The volume of each
reaction was 10 ml, and injections were made at 240 s intervals.
Top, raw data from the titration. Each peak corresponds to the
injection. Bottom, the peaks in the upper panel were integrated
with ORIGIN software and the values were plotted versus
injection number. Each point corresponds to the heat in mcal
generated by the reaction upon each injection. The solid line is the
curve fit to the data by the Origin program. This fit yields values
for Kd. Experiments were done with wild type protein and others
variants, and gave similar raw data and curve fit. (A) WT; (B)
variant G41M; (C) variant I42W.
(EPS)
Figure S3
Binding of actinonin to AtPDF does barely modify the
crystal packing. (A) Crystal pack of the two complexes: open, free
complex (left) and bound to actinonin (right) (B). Non-crystallo-
graphic contacts into asymmetric unit are not modified by closing
movement of the protein due to actinonin binding, except for zinc
atom number 6. This metal ion is coordinated by side chains of
Asp40 and Glu63, and water molecules, Asp40 and Glu63 being
hydrogen bonded by side chain of Lys38 of the other subunit of
the asymmetric unit. With the closing movement of the protein
into the crystal, Ca of Asp40 shifted by 3.1 A˚ and its side chain
flipped by 90u. Therefore, it does no longer participate to the
coordination shell of this Zn2+ ion. However, it is still hydrogen
bonded by Lys38 from chain B.
(EPS)
Figure S4
Binding of actinonin to AtPDF closely mimics both
actinonin and product binding to EcPDF. Superimposition of
EcPDF and AtPDF bound to either actinonin (1LRU PDB code,
panel A) or Met-Ala-Ser (1BS6 PDB code, panel B), the product of
the
reaction.
The
r.m.s.d.
value
is
1.11
A˚
for
151
Ca
superimposed.
(EPS)
Figure S5
The ligand binding site of AtPDF. This picture shows
the residues of AtPDF that are in contact with actinonin (left) and
6b (right) according to the 3-D structure; this should be compared
to the similar scheme shown in Figure 1B for EcPDF.
(EPS)
Figure S6
Electronic densities of the moving side-chains and of
actinonin at the binding site in some variants of AtPDF. Actinonin
and selected residues (G/Q/M41, I42, F58, and I130) are drawn
in stick and are shown in their FO–FC electron density omit maps
contoured at 2s, in free wild-type AtPDF (two crystallization
conditions, WT1 and WT2), and ligand-bound WT (actinonin, 6b
and 21), G41Q, and G41M variants.
(EPS)
Figure S7
Only few residues show alternative conformation in
AtPDF. Alternative conformers in the crystalline state of AtPDF.
Ringer
plots
of
electron
density
(r)
versus
x1
angle
for
representative residues of the 3-D apostructure of AtPDF. Data
were obtained with the 3M6O dataset (see Table S1). The
secondary peaks in the Ile residues are observed because Ile is a
branched amino acid. To evidence an alternative conformation
with Ile, three peaks should be observed.
(EPS)
Figure S8
Impact of induced fit on the binding free energy of
actinonin depends on the capacity to stabilize a hydrogen bond
with PDF. (A) The gyration radii [83] of the side chain occurring
at position 41 is displayed with black squares and compared to the
kcat/Km values (grey bars). (B) The distance between the NH of I42
and the CO of actinonin was measured in each case. The
percentage of the distance required to make a hydrogen bond (2.8
A˚ ) is reported (dark squares). The difference of binding free energy
(DDGbinding) between the open, free state and the variants closed
complexes of the G41 variants are displayed as grey bars. The
values were calculated as follows. For the WT, it corresponds to
the RT ln(KI*/KI) value [29], where R is the ideal gas constant and
T is the temperature in Kelvin. RT is 0.616 kcal.mol21 at 37uC.
For the G41M and G41Q variants, the DDGbinding corresponds to
RT ln(KI-G41variant/KD-WT). The obtained values are similar to that
obtained if the kcat/Km substitutes the KD value in the calculation
(DDGbinding = RT ln(kcat/Km –G41variant/kcat/Km –WT).
(EPS)
Figure S9
Compound 21 does not bind AtPDF1B at S1’. (A) 21
is shown in ball-and-stick format in its FO–FC electron density omit
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map contoured at 2s. (B) Binding site of 21 into AtPDF1B is
detailed. Red and blue residues indicate residues that accommo-
date
the
‘‘phenylalanine’’
and
‘‘trimethyl’’
groups
of
21,
respectively. (C) Overall view of 21 binding site (left). Molecular
surface of AtPDF is represented, as well as 21 in ball-and-stick
format. Residues belonging to the 21 binding pocket are colored in
orange. For comparison, molecular surface of EcPDF (PDB code
1G2A) in the same orientation is also represented, with residues
forming the new ligand binding pocket colored in orange.
Actinonin is represented in ball-and-stick format and is seen
through the molecular surface of each PDF. (D) Ball-and-stick
representation of the interaction network around compound 21.
The metal cation is shown as a grey sphere.
(EPS)
Figure S10
Poorly active versions of AtPDF are in a closed
conformation incompatible with actinonin binding. (A) Free and
close AtPDF were superimposed as in Figure 1C and are figured in
brown and yellow, respectively. Both the G41M (chain B, shown
in orange) and the free C-deleted weakly active AtPDF versions
([63], colored in purple, PDB entry code 3CPM) were superim-
posed, to the two structures, showing that they both fit better to the
ligand-bound full-length close form than to the free open form, but
that the closure is further pronounced, burying the entrance to a
ligand. (B) Close-up showing that the shape of the S1’ pocket of the
poorly active closed versions make it poorly available to P1’
recognition (see circled Ile142 and Ile130 side chains).
(EPS)
Table S1
Catalytic properties of AtPDF. Nm, not measurable;
ND, not determined; WT, is wild-type. aKinetic constants were
determined using the coupled assay as indicated in Materials and
Methods with substrate Fo-Met-Ala-Ser, in the presence of 100
nM enzyme variant and 750 mM NiCl2, at 37uC. The relative
value of kcat/Km for wild-type AtPDF was set at 100%. bData
correspond to the binding constant of actinonin as obtained either
from ITC or from enzymatic analysis when indicated with an
asterisk. cData from Table S3. dGyration radii are from [83].
(DOC)
Table S2
Crystallographic data and refinement statistics. Values
in parentheses are for the outer resolution shell. aRsym (I) =
ShklSi|Ihkl,i 2 ,Ihkl.|/ShklSi|Ihkl,i|, where ,Ihkl. is the mean
intensity of the multiple Ihkl,i observations for symmetry-related
reflections. bRwork = 1006(Shkl|Fobs 2 Fcalc|/Shkl|Fobs|). Rfree
is a test set including ,5% of the data. cPercentage of residues in
most-favored/additionally
allowed/generously
allowed/disal-
lowed regions of the Ramachandran plot. dCompound 21 was
added first, and actinonin afterwards.
(DOC)
Table S3
Kinetic parameters for inhibition of some AtPDF
variants by actinonin. The enzyme concentration used in the assay
was 100 nM. Prior to kinetic analysis for determination of KI*app
values, actinonin was incubated in the presence of each variant set
at the final concentration for 10 min at 37uC; kinetic assay was
started
by
adding
a
small
volume
of
the
substrate.
For
determination of KI, k5, and k6 values, actinonin was not pre-
incubated with enzyme and kinetic assay was started by adding the
enzyme.
(DOCX)
Movie S1
Dynamics of actinonin binding to peptide deformylase
and closure of the active site.
(WMV)
Movie S2
Progressive motions of the main side chains at the
active site and final locking of the hydrogen bond.
(WMV)
Acknowledgments
We are strongly indebted to James Fraser and Tom Alber (University of
California, Berkeley, USA) for introducing us to Ringer before the release
of the freely available downloadable version. We thank Benoıˆt Gigant,
Virginie Gueguen-Chaignon, Solange Morera, and Philippe Peynot
(CNRS, Gif-sur-Yvette, France) for help with data processing and access
to the crystallization facilities. We also thank Magali Nicaise-Aumont
(IBBMC, Orsay, France), who performed the microcalorimetry experi-
ments. We are grateful to the staff of the European Synchrotron Radiation
Facility (ESRF) and SOLEIL beamlines for their help during data
collection.
Author Contributions
The author(s) have made the following declarations about their
contributions: Conceived and designed the experiments: SF CG TM.
Performed the experiments: AB SF. Analyzed the data: FD MD SF CG
TM. Contributed reagents/materials/analysis tools: IA MD CG TM.
Wrote the paper: CG TM.
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The Dynamics of Induced Fit at High Resolution
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|
3M6U
|
Multi-site-specific 16S rRNA methyltransferase RsmF from Thermus thermophilus in space group 43
|
Multi-site-specific 16S rRNA methyltransferase RsmF
from Thermus thermophilus
HASAN DEMIRCI,1,3 LINE H.G. LARSEN,2,3 TRINE HANSEN,2 ANETTE RASMUSSEN,2 ASHWIN CADAMBI,1
STEVEN T. GREGORY,1 FINN KIRPEKAR,2 and GERWALD JOGL1
1Department of Molecular Biology, Cell Biology and Biochemistry, Brown University, Providence, Rhode Island 02912, USA
2Department of Biochemistry and Molecular Biology, University of Southern Denmark, 5230 Odense M, Denmark
ABSTRACT
Cells devote a significant effort toward the production of multiple modified nucleotides in rRNAs, which fine tune the ribosome
function. Here, we report that two methyltransferases, RsmB and RsmF, are responsible for all four 5-methylcytidine (m5C)
modifications in 16S rRNA of Thermus thermophilus. Like Escherichia coli RsmB, T. thermophilus RsmB produces m5C967. In
contrast to E. coli RsmF, which introduces a single m5C1407 modification, T. thermophilus RsmF modifies three positions,
generating m5C1400 and m5C1404 in addition to m5C1407. These three residues are clustered near the decoding site of the
ribosome, but are situated in distinct structural contexts, suggesting a requirement for flexibility in the RsmF active site that is
absent from the E. coli enzyme. Two of these residues, C1400 and C1404, are sufficiently buried in the mature ribosome
structure so as to require extensive unfolding of the rRNA to be accessible to RsmF. In vitro, T. thermophilus RsmF methylates
C1400, C1404, and C1407 in a 30S subunit substrate, but only C1400 and C1404 when naked 16S rRNA is the substrate. The
multispecificity of T. thermophilus RsmF is potentially explained by three crystal structures of the enzyme in a complex with
cofactor S-adenosyl-methionine at up to 1.3 A˚ resolution. In addition to confirming the overall structural similarity to E. coli
RsmF, these structures also reveal that key segments in the active site are likely to be dynamic in solution, thereby expanding
substrate recognition by T. thermophilus RsmF.
Keywords: rRNA methyltransferase; 5-methylcytidine; RsmB; RsmF; RNA mass spectrometry
INTRODUCTION
Ribosomal RNAs (rRNAs) are post-transcriptionally mod-
ified in all three domains of life, and many modifications
are phylogenetically conserved. Most modifications are
located in functionally important regions of the ribosome,
where they probably act to fine tune protein synthesis (Agris
2004; Gustilo et al. 2008). Complete modification maps of
bacterial 16S rRNAs have been determined for only a hand-
ful of species, and among these are the enteric bacterium
Escherichia coli and the extremely thermophilic bacterium
Thermus thermophilus (Guymon et al. 2006). Despite the
large phylogenetic divergence of these two organisms, their
ribosome modification patterns are quite similar. Of the 11
E. coli and 14 T. thermophilus 16S rRNA modifications,
eight are identical. This suggests a set of common functional
requirements conserved since divergence from their last
common ancestor, and also suggests common recognition
mechanisms among their modifying enzymes.
For most ribosome modifications, a single enzyme recog-
nizes and modifies a single site. However, there exist nota-
ble exceptions. Among these are dimethylation of two adja-
cent adenosines in 16S rRNA by KsgA (Helser et al. 1972);
pseudouridylation of three adjacent residues in tRNAs by
TruA (Hur and Stroud 2007); pseudouridylation of several
tRNA residues by Pus1 (Motorin et al. 1998), Pus2 (Behm-
Ansmant et al. 2007), or Pus7 (Behm-Ansmant et al. 2003);
or methylation of four tRNA positions by Saccharomyces
cerevisiae Trm4 (Motorin and Grosjean 1999). Even with
these multi-site-specific enzymes, however, homologs from
various species generally modify the same residues.
E. coli 16S rRNA contains two 5-methylcytidine (m5C)
residues, located in or near the highly conserved decoding
3These authors contributed equally to this work.
Abbreviations: rRNA, ribosomal RNA; AdoMet, S-adenosyl-L-methionine;
m5C, 5-methyl-cytidine; MALDI mass spectrometry, matrix-assisted laser
desorption ionization mass spectrometry.
Reprint requests to: Gerwald Jogl, Department of Molecular Biology,
Cell Biology and Biochemistry, Brown University, Box G-E129, Provi-
dence, RI 02912, USA; e-mail: Gerwald_Jogl@brown.edu; fax: (401)
863-6114; or Finn Kirpekar, Department of Biochemistry and Molecular
Biology, University of Southern Denmark, Campusvej 55, 5230 Odense M,
Denmark; e-mail: f.kir@bmb.sdu.dk; fax: (+45) 65502467.
Article published online ahead of print. Article and publication date
are at http://www.rnajournal.org/cgi/doi/10.1261/rna.2088310.
1584
RNA (2010), 16:1584–1596. Published by Cold Spring Harbor Laboratory Press. Copyright 2010 RNA Society.
center of the 30S subunit (Fig. 1). An m5C967 modification
is produced by RsmB (also called Fmu), while an m5C1407
modification is produced by RsmF, formerly known as
YebU (Andersen and Douthwaite 2006). T. thermophilus
16S rRNA contains m5C967 and m5C1407, as well as two
additional m5C nucleotides, m5C1400 and m5C1404 (E. coli
rRNA numbering used throughout) (Guymon et al. 2006).
While the m5C967 and m5C1407 modifications are pre-
sumably produced by RsmB and RsmF homologs, respec-
tively, the source of the two additional m5C residues has
been unknown. Here we demonstrate that T. thermophilus
RsmF is a multi-site-specific methyltransferase and, in
contrast to the single-site-specific E. coli RsmF, is respon-
sible for the synthesis of three modifications: m5C1407,
m5C1400, and m5C1404. We also demonstrate that RsmB is
responsible for the synthesis of m5C967 in T. thermophilus
as well as is in E. coli, thereby accounting for all four m5C
modifications of 16S rRNA. We present crystal structures
of T. thermophilus RsmF up to 1.3 A˚ resolution that reveal
a dynamic region in the active site that is absent from the E.
coli RsmF structure, providing a possible explanation for
the expanded recognition capacity of the T. thermophilus
methyltransferase.
RESULTS
Identification of T. thermophilus 16S rRNA
m5C-methyltransferases
With the E. coli RsmB and RsmF protein sequences as
queries, we used conventional BLAST searches (Altschul
et al. 1990) to identify potential homologs encoded by the
T. thermophilus HB8 genome (data not shown). Both RsmB
and RsmF have the highest similarity to the T. thermophilus
protein encoded by TTHA1387 (BLAST scores of 106 and
190, respectively) and second-highest similarity to the pro-
tein encoded by TTHA0851 (BLAST scores of 93 and 81,
respectively). The simplest interpretation of these results is
that TTHA1387 encodes RsmF, responsible for methylation
of C1407, leaving TTHA0851 as the most likely candidate
for the gene encoding RsmB, responsible for methylation of
C967. The similarities of the two E. coli enzymes with other
T. thermophilus proteins were far too low to reveal potential
candidates responsible for methylation of C1400 and C1404.
We next constructed T. thermophilus strains in which
either TTHA0851 or TTHA1387 was inactivated by the
homologous recombination and insertion of a heat stable
kanamycin-resistance gene. 16S rRNA was isolated from
these null mutants and subfragments of z50 nucleotides
(nt) around the regions of interest were further purified,
digested with RNase T1, and analyzed by MALDI mass
spectrometry (Fig. 2). Comparison of the TTHA1387 null
mutant to wild-type T. thermophilus HB8 indicates three
clear differences, each corresponding to the disappearance
of a methyl group (z14.0 Da). The RNase T1 digestion
fragment harboring m5C1407, the nucleotide methylated
by RsmF in E. coli, is absent in the null mutant, indicating
that TTHA1387 is indeed rsmF. The predicted RNase T1
fragment reduced by 14.0 Da is obscured by another RNase
T1 fragment that is present in both the wild-type and
TTHA1387 null mutants (Fig. 2B). Unexpectedly, two addi-
tional RNase T1 fragments are also reduced by 14.0 Da.
One of these contains C1400 while the other contains
C1404. This latter RNase T1 fragment from wild-type
T. thermophilus contains three methyl groups, two on
m4Cm1402 and one on m5C1404 (Guymon et al. 2006),
preventing an unambiguous identification of the missing
methyl group. We therefore performed tandem mass spec-
trometry on the 1402CCCG1405 RNase T1 fragment with two
methyl groups from the TTHA1387 null mutant and com-
pared it with the triply methylated wild-type RNase T1
fragment (Fig. 2C). The clear w2 ions, as well as the less
intense z3 ions, display a 14.0 Da mass difference between
the two samples, showing that the methylations on
m4Cm1402 were not affected by inactivation of TTHA1387.
Tandem mass spectrometry was also performed on the
RNase T1 fragments appearing as a consequence of the lack
of methylations on C1400 and C1407 (data not shown).
As expected, the C1400-containing fragment revealed no
FIGURE 1. Secondary structure diagram of the 39 minor domain of
16S rRNA indicating the position of the three RsmF substrate
nucleotides.
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1585
indications of a methyl group, whereas
the RNase T1 fragment with C1407 ex-
hibited a fragmentation pattern corre-
sponding to the expected mass overlap
with an RNase T1 fragment of a different
sequence. In summary, our data lead us
to conclude that TTHA1387 encodes an
RsmF m5C methyltransferase responsible
for synthesizing m5C1400, m5C1404, and
m5C1407 in 16S rRNA of T. thermophilus.
The
above
results
left
us
with
TTHA0851 as the sole candidate for
the gene encoding the m5C967 methyl-
transferase. An approach conceptually
identical to that described above re-
vealed that disruption of TTHA0851
reduced the relevant RNase T1 fragment
by 14.0 Da (Supplemental Fig. 1A).
Since this fragment is methylated at
G966 and C967 in the wild-type strain
(Guymon et al. 2006), tandem mass
spectrometry was again performed (Sup-
plemental Fig. 1B), showing that only
the methyl group on C967 was absent. In
agreement with the suggested nomen-
clature for rRNA modifying enzymes
(Andersen and Douthwaite 2006), we
hereafter refer to TTHA0851 as rsmB
due to substrate specificity identical to
the originally identified enzyme from
E. coli (Gu et al. 1999).
Effect of temperature on growth
of the rsmF null mutant
One possible explanation for methyla-
tion of multiple sites by RsmF is that
such additional methyl groups improve
ribosomal function at elevated temper-
ature. To address this possibility, we ex-
amined the effect of temperature on
growth of the rsmF null mutant. Wild-
type T. thermophilus and the rsmF null
mutant were cocultured at three tem-
peratures and these cocultures were se-
rially subcultured for seven cycles of
24 h each. The proportion of wild-type
and rsmF null mutant cells in each mixed
culture was determined by spreading di-
lutions onto TEM plates with or without
kanamycin. After seven cycles, no differ-
ence in the relative proportions of the
wild-type and rsmF mutant was exhib-
ited at 70°C. However, at 60°C, the rsmF
null mutant constituted only around 5%
FIGURE 2. (Legend on next page)
Demirci et al.
1586
RNA, Vol. 16, No. 8
of the population, and at 80°C the rsmF null mutant was
unable to grow at all. Thus, methylation by RsmF appears to
facilitate growth at temperatures outside the optimal growth
temperature.
RsmF substrate preference
The T. thermophilus rsmB and rsmF genes were each cloned
into an E. coli expression plasmid in order to produce
proteins for X-ray crystallography and in vitro methylation
studies. The expression constructs were equipped with
C-terminal histidine6 tags to facilitate protein purification.
While we achieved a high expression level of RsmF, we were
unable to do so with RsmB despite a series of optimization
attempts. Consequently, in vitro substrate and structure
analyses were performed exclusively with RsmF.
E. coli RsmF requires the 30S ribosomal subunit as a
substrate when the activity is assayed in vitro (Andersen and
Douthwaite 2006). We assayed 70S ribosomes, 30S ribosomal
subunits, and 16S rRNA for the ability to serve as substrates
for methylation by RsmF in vitro. 16S rRNA subfragments of
z50 nt around the target sites were purified after the in vitro
assay and analyzed by mass spectrometry as described above.
In vitro methylation at 70°C showed an interesting but rather
complex substrate pattern. RsmF completely methylates
C1400 when either 16S rRNA or 30S subunits are used as
a substrate. It methylates C1404 to z35% with 16S rRNA
and completely with 30S subunits, and it produces only trace
amounts of methylation of C1407 with 16S rRNA and z75%
with 30S subunits (Fig. 3). There were no indications of
the 70S ribosome being a substrate in vitro. Curiously,
T. thermophilus RsmF expressed in an E. coli rsmF null
mutant almost completely methylated, in vivo, positions
C1400 and C1404, but not C1407 (data not shown).
X-ray crystal structures of RsmF
We determined the structure of T. thermophilus RsmF (456
amino acids) in three different crystal forms and in a com-
plex with cofactor AdoMet to up to 1.3 A˚ resolution (Figs.
4, 5). The structure was solved in space group P43 (data set
RsmF1, 1.4 A˚ resolution) by molecular replacement using
a search model generated with the program Modeller
(Eswar et al. 2008) from the catalytic domain of the RsmF
homolog YebU from E. coli (Pdb 2FRX) (Hallberg et al.
2006). The structures of the AdoMet-bound form in space
group P2 (RsmF2, 1.82 A˚ resolution), of the AdoMet-
bound form (RsmF3, 1.3 A˚ resolution), and of the apo-
form (RsmF4, 1.68 A˚ resolution) in space group P21212
were subsequently solved by molecular replacement with
the refined RsmF1 model. There are two molecules in the
asymmetric unit in space groups P43 and P2 and one mol-
ecule in space group P21212. Electron density is generally
well defined in all crystal forms. The majority of residues
(92.0%, 92.3%, 93.1%, and 92.6%) are in the most favored
region of the Ramachandran plot for RsmF1, RsmF2,
RsmF3, and RsmF4, respectively, and there are no residues
in the disallowed region. The final models consist of
residues 5–178, 194–198, and 201–456 and five additional
residues from the histidine6 affinity tag in both chains of
data set RsmF1; residues 2–456 and five affinity-tag
residues in both chains of data set RsmF2; residues 1–456
and six affinity-tag residues in data set RsmF3; and resi-
dues 1–456 and seven affinity-tag residues in RsmF4. The
N-terminal a-amino group was ordered in data sets RsmF3
and RsmF4 and contained additional electron density,
which we interpreted as N-(dihydroxymethyl)-L-methio-
nine, the hydrated form of N-formyl-methionine. Data
collection and refinement statistics are given in Table 1.
The overall structure of RsmF consists of a central
canonical class I methyltransferase catalytic domain with
additional N-terminal and C-terminal
domains (Figs. 4, 5). The catalytic do-
main is formed by a central seven-
stranded b-sheet that is flanked on both
sides by three helices of varying lengths.
An inserted region between strand b7
and helix a11 contains additional heli-
ces a9 and a10, which interact with the
two N-terminal helices a1 and a2. A
second inserted region following strand
b9 includes the short helices a13 and
FIGURE 2. (A) MALDI mass spectra of an RNase T1-digested 16S rRNA subfragment (pos.
1378–1432) from wild-type cells (upper panel) or from the TTHA1387 (putative rsmF) null
mutant (lower panel). Expected digestion products are labeled; fragments affected by the null
mutation are set in italics. (B) Expansion of the signals affected by the TTHA1387 null mu-
tation. The sequence and methylation status of the RNase T1 products are indicated. (C)
MALDI tandem mass spectrometry of the methylated RNase T1 fragment of 16S rRNA (pos.
1404–1407); wild-type cells (upper panel), TTHA1387 null mutant (lower panel). Mass
spectrometric fragments used to deduce the methylation status are labeled. The position of
the backbone fragments (nomenclature according to McLuckey et al. [1992]) in the sequence is
shown. MH+, precursor ion selected for fragmentation; C, cytosine; mC, methylated cytosine;
C>p, cytidine-2´-39-monophosphate; me, methyl group.
FIGURE 3. In vitro methylation with 30S ribosomal subunits or 16S
rRNA from the T. thermophilus rsmF null mutant as a substrate. Effect
on C1400-, C1404-, and C1407-harboring RNase T1 products. In
vitro methylated products are set in italics. *, artifact signal arising
from the enzyme preparation.
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1587
a14, which interact with helix a12. Furthermore, RsmF
contains three additional smaller domains, an N-terminal
domain consisting of a three-stranded b-sheet and two flank-
ing helices (Fig. 5B, colored in blue), and two C-terminal
domains consisting of four-stranded b-sheets and two or one
helix (Fig. 5B, colored in magenta and red).
Cofactor binding, substrate docking,
and conformational flexibility in the active site
The coordination of AdoMet in the T. thermophilus RsmF
active site is similar to that seen in other class I methyl-
transferases. However, the previously published structure of
E. coli RsmF did not contain the cofactor AdoMet in the
active site, precluding a direct comparison. Both the T.
thermophilus and E. coli RsmF cofactor-binding sites reveal
a new variation for the methyltransferase signature motif I
(Malone et al. 1995), with the highly conserved GxGxG
sequence replaced by 109AAAPG113. The combination of
three alanines and a proline results in a loop conformation
that is very similar to that observed in other methyltrans-
ferases with a GxGxG motif (e.g., RsmC) (Demirci et al.
2008a). In RsmF, the amide hydrogen atom of the last
glycine residue forms a hydrogen bond with the cofactor
carboxy group (Fig. 6). Other key interactions with AdoMet
are well conserved in RsmF. The cofactor adenine ring is
located in a mainly hydrophobic pocket lined by residues
Val134, Pro160, and Leu211. This pocket is open toward the
solvent. The adenine amino group is not specifically recog-
nized and interacts with solvent water molecules. The ribose
hydroxyl groups form hydrogen bonds with Glu133 and
Arg138, and the methionine amino group interacts with
Asp177. The AdoMet cofactor is bound in a cleft in the
RsmF active site, which suggests that substrate cytidine bases
are inserted into the active site in an unstacked conforma-
tion. Inspection of the electrostatic charge distribution re-
veals a large positively charged surface region, which would
be consistent with binding to an RNA surface and modifi-
cation of the substrate base in an unstacked orientation (Fig.
6C). To evaluate the possible orientation of a substrate base
in the active site, we performed computational docking
calculations with the program Dock6 (Lang et al. 2009). The
resulting positions of cytosine and m5C in the presence of
AdoMet in data set RsmF3 are highly similar to each other,
with m5C placed into the active site with its phosphate group
toward a positively charged pocket at the entrance of the
active site cleft (Fig. 6E). The position of the phosphate
group is close to a sulfate molecule that we observed in data
set RsmF1, providing further support for the results of the
docking calculation (Fig. 6F).
Interestingly, we observed that three active site segments
were disordered in data set RsmF1. These segments include
residues 179–193 (including helices a9 and a10 and the
catalytic Cys180), residues 199–200, and the N-terminal
residues 1–5, which interact with the first two segments (Fig.
6E,F, colored in green). We observed electron density for the
intervening residues 194–198, which formed a lattice con-
tact with a neighboring molecule. However, the position of
FIGURE 4. Structure-based sequence alignment of RsmF from T. thermophilus and E. coli. Secondary structure elements of T. thermophilus RsmF
are indicated on top. The color scheme for the secondary structure elements is as in Figure 5A. The position of the variant methyltransferase motif
I is marked with a red box. A flexible region observed in the active site is marked with a green box; residues interacting with the cofactor are
marked with orange boxes.
Demirci et al.
1588
RNA, Vol. 16, No. 8
these five residues was not related to their position in the
other three data sets, suggesting that the extended active
site region between residues 179 and 201 can reorient in the
RsmF structure. This observation suggests that this active
site region is dynamic, which may be important for sub-
strate binding at 72°C, the optimum growth temperature of
T. thermophilus.
DISCUSSION
Substrate recognition mechanisms
We have identified the two enzymes responsible for the
synthesis of the four m5C modifications of T. thermophilus
16S rRNA, and characterized the RsmF methyltransferase
responsible for synthesizing three of these. rRNA modifying
enzymes in bacteria are generally highly specific, with a
one-to-one association between the modifying enzyme and
the modification. A few cases of multitarget ribosome mod-
ifying enzymes have been reported (Helser et al. 1972;
Demirci et al. 2008b), but to our knowledge T. thermophi-
lus RsmF is the first rRNA methyltransferase found to mod-
ify three different nucleotides. Most ribosome modifying
enzymes probably recognize assembly intermediates, and
the data presented here are consistent with that notion.
T. thermophilus RsmF methylates C1400 and C1404 in
vitro using either 16S rRNA or 30S subunits as substrates,
whereas both E. coli (Andersen and Douthwaite 2006) and
T. thermophilus RsmF exclusively utilize 30S subunits as
substrates for methylation of C1407. This may reflect that
C1400 and C1404 methylations do not rely on the asso-
ciation of ribosomal proteins in order to be recognized by
T. thermophilus RsmF. C1407 methylation, in contrast,
depends on both rRNA and the ribosomal protein for the
recognition by RsmF in both T. thermophilus and E. coli.
More puzzling is the observation that T. thermophilus RsmF
does not methylate E. coli ribosomes in vivo on C1407. It
is perhaps worth noting that methylation of C1407 in the
T. thermophilus 30S ribosomal subunit in vitro was less
efficient than methylation of the other two positions,
indicating the need for a particular intermediate assembly
structure or for accessory factors. The only clear in vitro
FIGURE 5. Overall structure of RsmF. (A) Schematic representation of the position of the substrate bases in the 30S ribosomal subunit (Pdb
entry 2WRI) (Gao et al. 2009). C1400, C1404, and C1407 in helix 44 (green) are shown in stick representation with 5-methyl groups as pink
spheres. (B) Schematic stereo representation of the overall structure of RsmF. Secondary structure elements are in orange and yellow for the
catalytic domain, in salmon and blue for additional N-terminal domains, and in magenta and red for C-terminal domains. The flexible region in
the active site including helices a9 and a10 is indicated with an arrow and colored in green. (C) Topology diagram with secondary structure
elements colored as in B.
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1589
methylation observed at 37°C with T. thermophilus RsmF
was on C1400 with 16S rRNA as the substrate (data not
shown), which is not evidently related to the methylation
pattern in vivo in the heterologous system. It seems unlikely
that the aberrant methylation in the heterologous system
reflects species-specific differences in the mature 30S ribo-
somal subunit, given the extreme sequence and structural
conservation of the decoding site. Instead, it may reflect
differences in 30S subunit assembly in the two organisms,
necessary due to the large difference in growth temperature.
The four m5C residues in T. thermophilus 16S rRNA are
clustered in and around the functionally critical decoding
center at or close to sites of contact with tRNA, mRNA, and
EF-G (Ogle et al. 2002; Selmer et al. 2006; Gao et al. 2009).
m5C1400, m5C1404, and m5C1407 are located in the
subunit body, while m5C967 is located in the subunit head,
about 10 A˚ from m5C1400 (Wimberly et al. 2000). Exam-
ination of the 30S subunit crystal structure (Wimberly et al.
2000) indicates that the three bases methylated by RsmF are
situated in three distinct structural contexts, but provides
few clues to a common mode of substrate recognition.
C1400 is an unpaired base protruding from a sharply bent
segment of rRNA at the junction of helices 43 and 44, while
C1404 and C1407 are engaged in Watson–Crick pairs within
helix 44 (Fig. 4A). The C5 positions of the latter two bases
are not obviously accessible, such that RsmF would need to
approach them from the major groove side. While a flipping
of C1407 out of the helix via the minor groove could allow
access to this base, such a mechanism would be problematic
for C1404, whose minor groove side is packed against the
rest of the 30S subunit.
While m5C1404 and m5C1407 are about 11 A˚ apart
(Selmer et al. 2006), m5C1400 is quite distant from both of
these bases (about 21 and 30 A˚ , respectively). The pro-
truded conformation of m5C1400 in the mature 30S sub-
unit is due in part to base-pairing interactions between
adjacent bases and the 1500 region of 16S rRNA (C1399–
G1504 and G1401–C1501). As the 1500 region is one of the
last segments of 16S rRNA to be synthesized, C1400 could
potentially be positioned much closer to C1404 and C1407
in an assembly intermediate, prior to the formation of the
C1399–G1504 and G1401–C1501 base pairs. RsmF could
utilize a single binding mode to then access all three bases,
further facilitated by its flexible active site domain. Meth-
ylation of C1400 in mature 30S subunits would therefore
involve disruption of the adjacent base pairs. A complete
understanding of the recognition mechanism of these
enzymes will require high-resolution structural data on
assembly intermediate-enzyme complexes. Given the large
number of potential subunit assembly intermediates, pre-
cisely defining the physiological substrate for RsmB and
RsmF will be a formidable task.
TABLE 1. Data collection and refinement statistics
RsmF1
RsmF2
RsmF3
RsmF4
Data collectiona
AdoMet
AdoMet
Space group
P43
P2
P21212
P21212
Cell dimensions
a, b, c (A˚ )
71.0, 71.0, 186.7
66.0, 78.3, 108.1
89.7, 109.0, 51.0
89.8, 109.1, 50.8
a, b, g (°)
90, 90, 90
90, 107.1, 90
90, 90, 90
90, 90, 90
Resolution (A˚ )b
30–1.4 (1.55–1.40)
30–1.82 (1.89–1.82)
30–1.30 (1.34–1.30)
30–1.68 (1.74–1.68)
Rmerge
0.065 (0.59)
0.08(0.38)
0.058(0.36)
0.15 (0.49)
I/sI
29.3(2.15)
12.6 (2.04)
24.3 (2.05)
14.2 (1.73)
Completeness (%)
90.1 (72.6)
97.0(86.5)
95.6(66.3)
99.6 (97.8)
Redundancy
8.9 (5.4)
2.8(2.0)
5.2(2.1)
6.4 (3.9)
Refinement
Resolution (A˚ )
30–1.4 (1.42–1.40)
30–1.82 (1.84–1.82)
30–1.30 (1.32–1.30)
30–1.68 (1.69–1.68)
Number of reflections
161,955 (4356)
91,762 (2583)
119,490/2640
109,375 (3244)
Rwork/Rfree
0.169/0.189 (0.227/0.263)
0.162/0.194 (0.222/0.259)
0.177/0.191 (0.233/0.234)
0.173/0.192 (0.217/0.266)
Number of atoms
Protein
6766
7117
3598
3574
Ligand/ion
20
54
27
1
Water
1486
1285
856
665
B-factors
Protein
24.1
20.4
15.9
17.2
Ligand/ion
34.5
21.6
17.5
19.1
Water
38.7
36.3
33.6
33.6
RMSDs
Bond lengths (A˚ )
0.009
0.006
0.005
0.004
Bond angles (°)
1.22
1.04
1.14
0.94
aOne crystal used for each data set.
bThe highest resolution shell is shown in parentheses.
Demirci et al.
1590
RNA, Vol. 16, No. 8
Structural comparison of RsmF with related
methyltransferases
A database search with Dali (Holm et al. 2008) confirmed
the structural similarity of the T. thermophilus and E. coli
(PDB 2FRX) (Hallberg et al. 2006) RsmF homologs, which
superimpose with a root-mean-square deviation (RMSD)
of 1.6 A˚ for 342 Ca atoms (Fig. 7A) and
are the only two structures in the Pro-
tein Database with this domain organi-
zation. Even so, substantial structural
differences are observed in most of the
loop regions and for the long connect-
ing loop between the methyltransferase
domain and the first C-terminal domain.
While in the active site, the positions
of residues in the cofactor-binding site
and of the two cysteine residues are
conserved (Fig. 7B), there are a number
of positively charged residues (Arg30,
Arg190, Arg194, His195, and Arg203) in
the T. thermophilus structure that are
absent from the E. coli enzyme (Fig. 7B).
Three of these are located in the flexible
region, and the combination of a posi-
tive charge and flexibility close to the
active site is suggestive of a functional
contribution of this region to the mul-
tisite specificity of T. thermophilus RsmF
(Figs. 6C,D, 7B). Methylation of three
rRNA positions may require an increase
in the enzyme’s structural dynamics in
order to accommodate the 30S subunit
in slightly different orientations. Similar
observations have been made for other
multi-site-specific methyltransferases in-
cluding KsgA, which modifies two adja-
cent adenosines in the 30S ribosomal
subunit (O’Farrell et al. 2004; Demirci
et al. 2009), and the PrmA ribosomal
protein methyltransferase, which under-
goes dramatic interdomain movements
to modify multiple lysine residues and the
N-terminal a-amino group on the same
substrate protein (Demirci et al. 2007,
2008b).
The second C-terminal domain in
RsmF is related to the RNA-binding
PUA
(pseudouridine
synthase
and
archaeosine transglycosylase) domains
(Perez-Arellano et al. 2007). The RlmI
methyltransferase,
which
produces
m5C1962 in 23S rRNA (Purta et al.
2008) also contains a PUA domain
(Sunita et al. 2008), although it is
N-terminal to the catalytic methyltransferase domain and
in a different orientation. PUA domains contain six
b-strands, which form a central pseudobarrel closed by
a short 310-helix. A comparison of the C-terminal domain
in RsmF with a typical PUA domain in archaeosine trans-
glycosylase (ArcTGT, Pdb entry 1J2B) reveals that the
central fold is similar (53 Ca atoms align with an RMSD
FIGURE 6. Substrate docking and conformational flexibility. (A) Cofactor-binding site in
RsmF. Bound AdoMet is shown in blue sticks. Hydrogen bonds to coordinating residues are
indicated. (B) Final sA-weighted 2mFO-DFC electron density map of the cofactor-binding site
(data set RsmF2) contoured at the 1s level. (C,D) Comparison of the electrostatic surface
charge distribution between RsmF from T. thermophilus and E. coli. The location of the
C-terminal domains and of the flexible region (labeled ‘‘FLEX’’) is indicated with circles.
AdoMet and docked m5C are shown as sticks. AdoMet from the T. thermophilus structure is
shown with the E. coli structure for comparison. (E) The modeled position of m5C docked into
the active site of RsmF. m5C and AdoMet are shown in tan and blue sticks, respectively.
Residues in the flexible region are shown in green. (F) The active site region in data set RsmF1.
A sulfate ion is observed close to the position of the m5C phosphate group. Residues 194–198 in
the flexible region interacting with a neighboring RsmF molecule are indicated with an arrow.
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1591
of 1.8 A˚ ), but that several connecting loop regions are
substantially shorter and two a-helices and one b-strand of
the pseudobarrel are absent. Thus, this PUA-like domain
differs considerably from typical PUA domains. However,
the similarity to RNA-binding PUA domains and the
positive surface charge distribution observed in both RsmF
structures are suggestive for a conserved function of the
PUA-like domain in RNA recognition.
The next most closely related structure is E. coli RsmB
(474 residues, PDB 1SQF) (Foster et al. 2003). A total of
264 Ca atoms can be aligned with an RMSD of 1.5 A˚
between RsmB and RsmF. Both enzymes retain the same
organization for the N-terminal domains and the core
methyltransferase domain. However, an additional 140-
residue N-terminal RNA-binding domain provides sub-
strate specificity to RsmB, whereas the two C-terminal
domains following the core methyltransferase domain (160
residues) are likely to determine substrate recognition by
RsmF. Thus, these two enzymes have evolved substrate
specificity via acquisition of additional,
unrelated RNA recognition domains.
While there are as yet no enzyme–
substrate complexes for rRNA m5C-
methyltransferases, insights into the RsmF
catalytic mechanism can be gleaned from
a comparison with the RlmD and TrmA
m5U methyltransferases in covalent in-
termediate complexes with RNA oligonu-
cleotides (Lee et al. 2005; Alian et al.2008).
DNA and RNA m5C methyltransferases
use a thiol from a catalytic cysteine residue
to attack the six-position of the pyrimi-
dine base to activate the five-position for
methyl group transfer (Liu and Santi
2000). Cys180 and Cys230 in RsmF are
positioned equivalently to the catalytic
Cys324 and the catalytic base Glu358 of
TrmA. The substrate nucleotides insert
into TrmA and RsmF in unrelated di-
rections, consistent with a lack of struc-
tural homology outside the methyltrans-
ferase domains. Nevertheless, the C5
positions of the pyrimidine rings and of
the 5-methyl carbons are quite similar
with respect to the catalytic cysteine resi-
dues and the AdoMet cofactor. The same
structural homology of the active site
geometry can be observed in comparison
with the RlmD methyltransferase (Sup-
plemental Fig. 2; Lee et al. 2005).
A possible origin of T. thermophilus
RsmF
T. thermophilus RsmF shows the highest
similarities with proteins from close relatives, namely,
Thermus aquaticus and two Meiothermus species of the
Thermaceae family. Remarkably, the next highest similarities
(BLAST scores between 267 and 359) are with the NOL1/
NOP2/Sun proteins from the Gram-positive Firmicutes
phylum (Supplemental Fig. 3). This similarity, together
with the fact that the Thermaceae family and most members
of the Firmicutes identified in Supplemental Figure 3 are
thermophilic, suggest that rsmF has undergone horizontal
transfer between the Thermaceae family and members of the
Firmicutes phylum. Thus, we speculate that this version of
the RsmF protein, which catalyzes methylation of three
cytidines, may be adaptive for existence in thermally
challenging environments. The effect of the loss of methyl-
ation by RsmF on growth at different temperatures is
consistent with this notion. Our hypothesis of horizontal
transfer of rsmF predicts that RsmF of other members of the
Thermaceae family and members of the Firmicutes phylum
will also be found to introduce multiple m5C modifications.
FIGURE 7. Comparison with other methyltransferases. (A) Differences between the overall
structures of RsmF from T. thermophilus (orange) and E. coli (green). AdoMet bound in the T.
thermophilus structure is shown as blue sticks. (B) Comparison of the active site region in both
enzymes. Residues in the cofactor-binding site and in the flexible region in the T. thermophilus
enzyme are shown as sticks. (C) Comparison between RsmF (orange) and a substrate complex
structure of the TrmA methyltransferase (cyan/blue). (D) Comparison of the active site region
illustrating the differences in the insertion direction of the substrate base in RsmF (m5C in
light orange) and in TrmA (m5U in cyan).
Demirci et al.
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RNA, Vol. 16, No. 8
MATERIALS AND METHODS
Cloning of the T. thermophilus rsmB and rsmF genes
The T. thermophilus HB8 loci TTHA0851 (GenBank accession
number BAD70674) and TTHA1387 (GenBank accession number
BAD71210) were PCR amplified from genomic DNA and purified
via the High Pure PCR Template Preparation Kit (Roche). The
100 mL PCRs contained 150 ng DNA, 10 mM of each primer,
10 mM dNTP, 1 unit Phusion DNA polymerase (Finnzymes), and
1x Phusion HF buffer. Primers for rsmB amplification were 59-CC
CTGGACATATGAGGGCCGG-39 and 59-GGCCAAGATCTTGCC
TGAGAG-39, and the temperature cycling was as follows: 98°C/30
sec; 30X (98°C/10 sec, 59°C/20 sec, and 72°C/36 sec); and 72°C/420s.
Primers for rsmF amplification were 59-GCTAGGGTACACATA
TGCTGCCC-39 and 59-GCACGGGGGTGAGATCTAAGCCC-39,
and the temperature cycling was as follows: 98°C/30 sec; 30X
(98°C/10 sec, 62°C /20 sec, 72°C/42 sec); and 72°C/420 sec. The
desired PCR products were purified from agarose gels using the
GFX PCR purification kit (GE Healthcare). The PCR fragments
were digested with NdeI and BglII and inserted into the expression
vector pLJ102 (Andersen and Douthwaite 2006), generating
isopropyl-1-thio-b-D-galactopyranoside (IPTG)-inducible genes
for the recombinant proteins with a C-terminal histidine6 tag.
The constructs (designated pLJ102-RsmB and pLJ102-RsmF) were
used to transform an rsmF-deletion derivative of E. coli CP79
(Andersen and Douthwaite 2006).
Deletion of the T. thermophilus rsmB and rsmF genes
Constructs for inactivation of the T. thermophilus rsmB and rsmF
genes were made by inserting the gene for a heat tolerant
kanamycin (HTK) nucleotidyltransferase (Hoseki et al. 1999) into
the methyltransferase parts of either pLJ102-RsmB or pLJ102-
RsmF. The htk part of pUC18-htk (Hashimoto et al. 2001) was
amplified by PCR with primers that introduced an upstream AvrII
site and a downstream SacI site into the product for later disrup-
tion of rsmB. For rsmF disruption, the PCR primers introduced
SacI restriction enzyme sites both upstream of and downstream
from the htk gene. These sites were used to insert the PCR prod-
ucts into pLJ102-RsmB and pLJ102-RsmF to form the plasmids
pLJ102-RsmBThtk and pLJ102-RsmFThtk, which were propa-
gated in the E. coli strain Top10 (Invitrogen). T. thermophilus
HB8 was transformed with pLJ102-RsmBThtk or pLJ102-
RsmFThtk selecting for kanamycin resistance as described by
others (Hashimoto et al. 2001; Cameron et al. 2004). Kanamycin-
resistant transformants were restreaked twice. Gene disruptions
were verified by PCR with primers distal to the interrupted rsmB
or rsmF genes on genomic DNA; resulting PCR products were
characterized by sequencing.
Growth competition assays
Wild-type and rsmF null mutant liquid cultures were grown at
70°C to saturation, then equal numbers of cells from each were
mixed and incubated in 5 mL TEM medium at 60°C, 70°C, or
80°C. After growth for 24 h, 100 mL of the 60°C culture, 10 mL of
the 70°C culture, and 1000 mL of the 80°C culture were trans-
ferred to a fresh 5-mL medium and incubated at the respective
temperatures for another 24 h. This was repeated in independent
triplicates for seven cycles. Samples of 1 mL were collected at each
dilution and half was plated on TEM plates without antibiotic and
the other half was plated on TEM plates with 30 mg/mL
kanamycin. The plates were incubated at 70°C.
Purification of T. thermophilus ribosomal subunits
and ribosomes
T. thermophilus culture (1 L) was grown in TEM media (contain-
ing 30 mg/mL of kanamycin when appropriate) with shaking at
70°C to an OD600 = 0.6. Cells were harvested and washed once
with 100 mL of buffer A (10 mM NH4Cl, 20 mM MgCl2, 100 mM
KCl, and 10 mM Tris-HCl [pH 7.5]), then suspended in 10 mL of
buffer A, and disrupted by sonication. The lysate was cleared by
centrifugation twice in a Beckman JA20 rotor at 16,000 rpm for
10 min at 4°C. Crude ribosomes were collected by centrifugation
in a Beckman Ti50 rotor at 19,000 rpm for 19 h at 4°C, and
dissolved in buffer A. 70S ribosomes were obtained by centrifu-
gation of 100 A260 units of crude ribosomes through a 10%–40%
sucrose gradient (200 mM NH4Cl, 20 mM MgCl2, 20 mM Tris-
HCl [pH 7.5]) in a Beckman SW28 rotor at 20,000 rpm for 18 h at
4°C. Fractions containing intact 70S ribosomes were pooled and
concentrated by centrifugation in a Beckman Ti50 rotor at 40,000
rpm for 22 h at 4°C, washed, and dissolved in buffer A, and stored
at 80°C.
50S and 30S ribosomal subunits were obtained by adjusting
100 A260 units of crude ribosomes (10 mM NH4Cl, 2 mM MgCl2,
100 mM KCl, and 10 mM Tris-HCl [pH 7.5]) and passing
through a 5%–20% sucrose gradient (200 mM NH4Cl, 1 mM
MgCl2 and 20 mM Tris-HCl [pH 7.5]) in a Sorvall AH-629 rotor
at 20,000 rpm for 18 h at 4°C. After pooling of the relevant
fractions, the subunits were adjusted to 10 mM MgCl2 and
pelleted by centrifugation in a Beckman Ti50 rotor at 40,000
rpm for 22 h at 4°C, washed with and dissolved in 10 mM NH4Cl,
10 mM MgCl2, 100 mM KCl, and 10 mM Tris-HCl (pH 7.5), and
stored at 80°C.
Isolation of 16S rRNA and subfragments
from T. thermophilus and E. coli
Water (400 mL) was added to 100 mL of 30S ribosomal subunits
and the rRNA was extracted with 500 mL phenol, phenol/
chloroform, and chloroform. rRNA was ethanol precipitated
and dissolved in water. Purification of 16S rRNA subfragments
was performed as previously described (Andersen et al. 2004).
Briefly, 16S rRNA was hybridized to an excess of oligodeoxynu-
cleotide complementary to either the region 944–990 or the region
1378–1432. Single-stranded nucleic acids were digested with
Mung Bean Nuclease and RNase A. The resulting mixture was
separated on a polyacrylamide gel. Bands were visualized by ethid-
ium bromide staining, excised, and eluted.
E. coli CP79 with the endogenous rsmF inactivated, but com-
plemented with the T. thermophilus homolog on the plasmid
pLJ102-RsmF, were grown at 37°C to an OD450 = 0.45 in 200 mL
of LB medium containing 100 mg/L of ampicillin. RsmF expres-
sion was induced by addition of IPTG to 1 mM, and incubation
for another 3 h. Cells were harvested by centrifugation at 4°C,
washed in 100 mL TMN buffer (50 mM Tris-HCl [pH 7.8],
10 mM magnesium acetate, 100 mM NH4Cl), and resuspended in
2 mL TMN buffer prior to lysis by sonication (7 3 30 sec on ice)
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1593
and removal of debris by centrifugation (10 min/14,000 rpm/4°C/
microcentrifuge). Total RNA was recovered from the supernatant
by phenol extraction and ethanol precipitation. A 16S rRNA
subfragment was isolated as described above using an oligodeoxy-
nucleotide complementary to the region 1378–1432.
In vitro methylation
Reactions contained 50 pmol of 16S rRNA, 30S subunits, or 70S
ribosomes from the T. thermophilus TTHA1387 null mutant as the
substrate in a total volume of 100 mL (containing 100 mM NH4Cl,
10 mM MgCl2, 40 mM Hepes [pH 7.5]), 6 mM b-mercaptoetha-
nol, and 10% glycerol (prepared as a two times concentrated stock
solution), 1.5 mM S-adenosyl methionine, and 2 mg of recombi-
nantly expressed RsmF (see below).
For the reaction at 70°C, water and stock buffer were mixed and
left at room temperature for 15 min. Then a substrate, an enzyme
and S-adenosyl methionine were added and incubated at 70°C for
1 h. The 37°C reaction was started by mixing water and buffer
followed by 15 min at room temperature; the substrate was added
and the mixture transferred to 50°C for 5 min. After cooling to
37°C, S-adenosyl methionine and an enzyme were added and the
incubation continued for 1 h. Reactions were stopped by phenol/
chloroform extraction and the rRNA was recovered by ethanol
precipitation before purification of 16S rRNA subfragments as
described above. Control reactions without enzyme or S-adenosyl
methionine were carried out in all instances.
RNase T1 digestion and mass spectrometry
A purified 16S rRNA subfragment (1–2 pmol) was incubated with
2 units RNase T1 (Roche) and 50 mM 3-hydroxypicollinic acid
(3-HPA) in a total volume of 2 mL for 4 h at 37°C. MALDI mass
spectrometry was performed either on an ABI voyager STR in-
strument or a Waters Q-TOF MALDI instrument; MALDI tan-
dem mass spectrometry was done on a Waters Q-TOF MALDI
instrument. All spectra were recorded in positive ion mode using
3-HPA as the matrix. Experimental details were as previously de-
scribed (Douthwaite and Kirpekar 2007).
Protein expression and purification for crystallization
E. coli BL21 (DE3) (Invitrogen) containing pLJ102-RsmF was
grown to midlog phase in LB media at 37°C in the presence of
200 mg/mL ampicillin. Protein expression was induced at 20°C
with 400 mM IPTG. Cells were pelleted after 18 h by centrifuga-
tion at 4000 rpm for 20 min at 4°C and lysed by ultrasonication
on ice in a buffer containing 20 mM Tris-HCl (pH 8.5), 300 mM
NaCl, 5 mM b-mercaptoethanol, 0.1% Triton X-100, and 5%
glycerol. Cell debris and membranes were pelleted by centrifuga-
tion at 11,000 rpm for 30 min at 4°C. The soluble E. coli proteins
were precipitated by heat treatment at 65°C for 30 min and
pelleted by centrifugation at 11,000 rpm at 4°C for 30 min.
Soluble C-terminally hexahistidine-tagged T. thermophilus RsmF
was further purified by affinity chromatography with nickel-
nitrilotriacetic acid resin (Qiagen). Untagged proteins were re-
moved with buffer containing 20 mM Tris-HCl (pH 8.5), 250 mM
NaCl, and 1 mM imidazole (pH 8.5). Recombinant RsmF was
then eluted with the same buffer containing 150 mM imidazole.
The protein was then purified by cation exchange chromatogra-
phy (SP) (GE Healthcare) at pH 8.5, using a linear gradient of
10 mM to 1 M NaCl concentration. RsmF fractions were pooled
and concentrated and applied to a size-exclusion S200 column (GE
Healthcare) equilibrated with a buffer containing 20 mM Tris-HCl
(pH 8.5) and 200 mM NaCl. Purified RsmF was concentrated to
13 mg/mL for crystallization trials. The C-terminal hexahistidine
tag was not removed for crystallization. For the production of
selenomethionyl proteins, the expression construct was trans-
formed into B834 (DE3) cells (Novagen). The bacterial growth
was carried out in defined LeMaster medium (Hendrickson et al.
1990), and the protein was purified using the same protocol as for
the unmodified protein. To form the RsmF-AdoMet complex,
purified RsmF was mixed with 4 mM AdoMet incubated at 60°C
for 15 min and slowly cooled to room temperature before
performing crystallization experiments.
Crystallization of RsmF
All crystals were obtained using the microbatch technique under
oil at 4°C. To obtain the RsmF1 crystal form, 1 mL of protein
solution was mixed with the reservoir solution containing 20%
(w/v) PEG3350 and 200 mM sodium sulfate decahydrate (pH
6.6). Initial crystals grew over the course of 1–2 wk with maxi-
mum dimensions of 0.3 3 0.3 3 0.2 mm. To obtain the RsmF2
crystal form, 1 mL of the RsmF–AdoMet complex was mixed with
the reservoir solution containing 200 mM NaCl, 12% w/v
PEG8000 and 100 mM HEPES-KOH (pH7.5). Initial crystals
grew over the course of 2–3 wk with maximum dimensions of
0.1 3 0.4 3 0.4 mm. To obtain the RsmF3 crystal form, 1 mL of
the RsmF–AdoMet complex was mixed with the reservoir solution
containing 10% w/v PEG1000, 200 mM NaCl, and 100 mM Tris-
HCl (pH 8.5). The initial crystals grew over the course of 1–2 wk
with maximum dimensions of 0.05 3 0.3 3 0.4 mm. To obtain
the RsmF4 crystal form, 1 mL of the RsmF–AdoMet complex
solution was mixed with a reservoir solution containing 160 mM
magnesium chloride hexahydrate, 80 mM Tris-HCl (pH 8.5), and
24% w/v PEG4000. Initial crystals grew over the course of 1–2 wk
with maximum dimensions of 0.05 3 0.3 3 0.3 mm. RsmF1
crystals were gradually dehydrated by increasing the PEG3350 to
30% w/v and then cryoprotected in a mother liquor supplemented
with 25% v/v glycerol and then flash-frozen by being plunged into
liquid nitrogen. RsmF2 crystals were cryoprotected in a mother
liquor supplemented with 20% v/v ethylene glycol and then flash-
frozen by being plunged into liquid nitrogen. RsmF3 crystals
were cryoprotected by gradually increasing the concentration of
PEG1000 to 30% and then flash-frozen by being plunged into
liquid nitrogen. RsmF4 crystals were cryoprotected in a mother
liquor supplemented with 20% glycerol and then flash-frozen by
being plunged into liquid nitrogen.
Data collection
X-ray diffraction data for RsmF1, RsmF2, and RsmF4 crystals
were collected on a MAR CCD detector at the X4C beamline of
the National Synchrotron Light Source in Brookhaven at a wave-
length of 0.979 A˚ and 180°C. Diffraction data for RsmF3 crystals
were collected on an ADSC CCD detector at the X4A beamline of
the National Synchrotron Light Source in Brookhaven at a wave-
length of 0.979 A˚ and 180°C. Diffraction data for RsmF1 in
space group P43 were collected to 1.4 A˚ resolution with cell
Demirci et al.
1594
RNA, Vol. 16, No. 8
dimensions a = 71.0 A˚ , b = 71.0 A˚ , and c = 186.7 A˚ . Diffraction
data to 1.82 A˚ for RsmF2 were collected in space group P2 with
cell dimensions a = 66.0 A˚ , b = 78.3 A˚ , and c = 108.1 A˚ .
Diffraction data to 1.29 A˚ for RsmF3 were collected in space group
P21212 with cell dimensions a = 89.7 A˚ , b = 109.0 A˚ , and c = 51.0
A˚ . Diffraction data to 1.68 A˚ for RsmF4 were collected in space
group P21212 with cell dimensions a = 89.8 A˚ , b = 109.1 A˚ , and c =
50.8 A˚ . A single crystal was used for each data set. The diffraction
images were processed and scaled with the HKL2000 package
(Otwinowski and Minor 1997). The data processing statistics are
summarized in Table 1.
Structure determination and refinement
The RsmF structure was solved by molecular replacement with the
program Phaser (McCoy et al. 2007) from the CCP4 program
suite (Bailey 1994) in space group P43 to 1.4 A˚ resolution (data set
RsmF1). The initial search model was built with the program
Modeller (Eswar et al. 2008) from the catalytic domain of E. coli
YebU (Pdb code 2FRX). After the placement of two RsmF
catalytic domains in the asymmetric unit and the initial re-
finement with Refmac (Murshudov et al. 1997), the model was
further rebuilt with ARP/wARP (Langer et al. 2008). The resulting
model was 90% complete and manually checked and completed
with Coot (Emsley and Cowtan 2004). Final crystallographic re-
finement was performed with the program Phenix (Adams et al.
2002). The other crystal forms were subsequently solved by
molecular replacement. The atomic coordinates from the RsmF4
model were then used for initial refinement of the RsmF–AdoMet
complex structure in space group P21212 (RsmF3). There are two
molecules in the asymmetric unit in data sets RsmF1 and RsmF2,
and one molecule in RsmF3 and RsmF4. The crystallographic
R/Rfree factors are 0.17/0.19, 0.16/0.19, 0.18/0.19, and 0.17/0.19
for the four data sets: RsmF1, RsmF2, RsmF3, and RsmF4,
respectively. The stereochemical quality of the model was assessed
with Procheck (Laskowski et al. 1993). The Ramachandran sta-
tistics (most favored/additionally allowed/generously allowed/
disallowed) are 91.9%/8.1%/0.0%/0.0% for RsmF1, 91.9%/8.1%/
0.0%/0.0% for RsmF2, 93.6%/6.4%/0.0%/0.0% for RsmF3, and
92.5%/7.5%/0.0%/0.0% for RsmF4. The refinement statistics are
summarized in Table 1. Figures were generated using Pymol
(DeLano 2002).
Atomic coordinates
Coordinates and structure factors have been deposited in the
Protein Data Bank with accession codes 3M6U, 3M6V, 3M6W,
and 3M6X for data sets RsmF1, RsmF2, RsmF3, and RsmF4,
respectively.
SUPPLEMENTAL MATERIAL
Supplemental material can be found at http://www.rnajournal.org.
ACKNOWLEDGMENTS
We thank John Schwanof and Randy Abramowitz for access to the
X4A and X4C beamlines at the National Synchrotron Light Source.
This work was supported by grants GM19756 and GM19756-37S1
from the National Institutes of Health.
Received January 14, 2010; accepted April 26, 2010.
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3M6V
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Multi-site-specific 16S rRNA methyltransferase RsmF from Thermus thermophilus in space group P2 in complex with S-Adenosyl-L-Methionine
|
Multi-site-specific 16S rRNA methyltransferase RsmF
from Thermus thermophilus
HASAN DEMIRCI,1,3 LINE H.G. LARSEN,2,3 TRINE HANSEN,2 ANETTE RASMUSSEN,2 ASHWIN CADAMBI,1
STEVEN T. GREGORY,1 FINN KIRPEKAR,2 and GERWALD JOGL1
1Department of Molecular Biology, Cell Biology and Biochemistry, Brown University, Providence, Rhode Island 02912, USA
2Department of Biochemistry and Molecular Biology, University of Southern Denmark, 5230 Odense M, Denmark
ABSTRACT
Cells devote a significant effort toward the production of multiple modified nucleotides in rRNAs, which fine tune the ribosome
function. Here, we report that two methyltransferases, RsmB and RsmF, are responsible for all four 5-methylcytidine (m5C)
modifications in 16S rRNA of Thermus thermophilus. Like Escherichia coli RsmB, T. thermophilus RsmB produces m5C967. In
contrast to E. coli RsmF, which introduces a single m5C1407 modification, T. thermophilus RsmF modifies three positions,
generating m5C1400 and m5C1404 in addition to m5C1407. These three residues are clustered near the decoding site of the
ribosome, but are situated in distinct structural contexts, suggesting a requirement for flexibility in the RsmF active site that is
absent from the E. coli enzyme. Two of these residues, C1400 and C1404, are sufficiently buried in the mature ribosome
structure so as to require extensive unfolding of the rRNA to be accessible to RsmF. In vitro, T. thermophilus RsmF methylates
C1400, C1404, and C1407 in a 30S subunit substrate, but only C1400 and C1404 when naked 16S rRNA is the substrate. The
multispecificity of T. thermophilus RsmF is potentially explained by three crystal structures of the enzyme in a complex with
cofactor S-adenosyl-methionine at up to 1.3 A˚ resolution. In addition to confirming the overall structural similarity to E. coli
RsmF, these structures also reveal that key segments in the active site are likely to be dynamic in solution, thereby expanding
substrate recognition by T. thermophilus RsmF.
Keywords: rRNA methyltransferase; 5-methylcytidine; RsmB; RsmF; RNA mass spectrometry
INTRODUCTION
Ribosomal RNAs (rRNAs) are post-transcriptionally mod-
ified in all three domains of life, and many modifications
are phylogenetically conserved. Most modifications are
located in functionally important regions of the ribosome,
where they probably act to fine tune protein synthesis (Agris
2004; Gustilo et al. 2008). Complete modification maps of
bacterial 16S rRNAs have been determined for only a hand-
ful of species, and among these are the enteric bacterium
Escherichia coli and the extremely thermophilic bacterium
Thermus thermophilus (Guymon et al. 2006). Despite the
large phylogenetic divergence of these two organisms, their
ribosome modification patterns are quite similar. Of the 11
E. coli and 14 T. thermophilus 16S rRNA modifications,
eight are identical. This suggests a set of common functional
requirements conserved since divergence from their last
common ancestor, and also suggests common recognition
mechanisms among their modifying enzymes.
For most ribosome modifications, a single enzyme recog-
nizes and modifies a single site. However, there exist nota-
ble exceptions. Among these are dimethylation of two adja-
cent adenosines in 16S rRNA by KsgA (Helser et al. 1972);
pseudouridylation of three adjacent residues in tRNAs by
TruA (Hur and Stroud 2007); pseudouridylation of several
tRNA residues by Pus1 (Motorin et al. 1998), Pus2 (Behm-
Ansmant et al. 2007), or Pus7 (Behm-Ansmant et al. 2003);
or methylation of four tRNA positions by Saccharomyces
cerevisiae Trm4 (Motorin and Grosjean 1999). Even with
these multi-site-specific enzymes, however, homologs from
various species generally modify the same residues.
E. coli 16S rRNA contains two 5-methylcytidine (m5C)
residues, located in or near the highly conserved decoding
3These authors contributed equally to this work.
Abbreviations: rRNA, ribosomal RNA; AdoMet, S-adenosyl-L-methionine;
m5C, 5-methyl-cytidine; MALDI mass spectrometry, matrix-assisted laser
desorption ionization mass spectrometry.
Reprint requests to: Gerwald Jogl, Department of Molecular Biology,
Cell Biology and Biochemistry, Brown University, Box G-E129, Provi-
dence, RI 02912, USA; e-mail: Gerwald_Jogl@brown.edu; fax: (401)
863-6114; or Finn Kirpekar, Department of Biochemistry and Molecular
Biology, University of Southern Denmark, Campusvej 55, 5230 Odense M,
Denmark; e-mail: f.kir@bmb.sdu.dk; fax: (+45) 65502467.
Article published online ahead of print. Article and publication date
are at http://www.rnajournal.org/cgi/doi/10.1261/rna.2088310.
1584
RNA (2010), 16:1584–1596. Published by Cold Spring Harbor Laboratory Press. Copyright 2010 RNA Society.
center of the 30S subunit (Fig. 1). An m5C967 modification
is produced by RsmB (also called Fmu), while an m5C1407
modification is produced by RsmF, formerly known as
YebU (Andersen and Douthwaite 2006). T. thermophilus
16S rRNA contains m5C967 and m5C1407, as well as two
additional m5C nucleotides, m5C1400 and m5C1404 (E. coli
rRNA numbering used throughout) (Guymon et al. 2006).
While the m5C967 and m5C1407 modifications are pre-
sumably produced by RsmB and RsmF homologs, respec-
tively, the source of the two additional m5C residues has
been unknown. Here we demonstrate that T. thermophilus
RsmF is a multi-site-specific methyltransferase and, in
contrast to the single-site-specific E. coli RsmF, is respon-
sible for the synthesis of three modifications: m5C1407,
m5C1400, and m5C1404. We also demonstrate that RsmB is
responsible for the synthesis of m5C967 in T. thermophilus
as well as is in E. coli, thereby accounting for all four m5C
modifications of 16S rRNA. We present crystal structures
of T. thermophilus RsmF up to 1.3 A˚ resolution that reveal
a dynamic region in the active site that is absent from the E.
coli RsmF structure, providing a possible explanation for
the expanded recognition capacity of the T. thermophilus
methyltransferase.
RESULTS
Identification of T. thermophilus 16S rRNA
m5C-methyltransferases
With the E. coli RsmB and RsmF protein sequences as
queries, we used conventional BLAST searches (Altschul
et al. 1990) to identify potential homologs encoded by the
T. thermophilus HB8 genome (data not shown). Both RsmB
and RsmF have the highest similarity to the T. thermophilus
protein encoded by TTHA1387 (BLAST scores of 106 and
190, respectively) and second-highest similarity to the pro-
tein encoded by TTHA0851 (BLAST scores of 93 and 81,
respectively). The simplest interpretation of these results is
that TTHA1387 encodes RsmF, responsible for methylation
of C1407, leaving TTHA0851 as the most likely candidate
for the gene encoding RsmB, responsible for methylation of
C967. The similarities of the two E. coli enzymes with other
T. thermophilus proteins were far too low to reveal potential
candidates responsible for methylation of C1400 and C1404.
We next constructed T. thermophilus strains in which
either TTHA0851 or TTHA1387 was inactivated by the
homologous recombination and insertion of a heat stable
kanamycin-resistance gene. 16S rRNA was isolated from
these null mutants and subfragments of z50 nucleotides
(nt) around the regions of interest were further purified,
digested with RNase T1, and analyzed by MALDI mass
spectrometry (Fig. 2). Comparison of the TTHA1387 null
mutant to wild-type T. thermophilus HB8 indicates three
clear differences, each corresponding to the disappearance
of a methyl group (z14.0 Da). The RNase T1 digestion
fragment harboring m5C1407, the nucleotide methylated
by RsmF in E. coli, is absent in the null mutant, indicating
that TTHA1387 is indeed rsmF. The predicted RNase T1
fragment reduced by 14.0 Da is obscured by another RNase
T1 fragment that is present in both the wild-type and
TTHA1387 null mutants (Fig. 2B). Unexpectedly, two addi-
tional RNase T1 fragments are also reduced by 14.0 Da.
One of these contains C1400 while the other contains
C1404. This latter RNase T1 fragment from wild-type
T. thermophilus contains three methyl groups, two on
m4Cm1402 and one on m5C1404 (Guymon et al. 2006),
preventing an unambiguous identification of the missing
methyl group. We therefore performed tandem mass spec-
trometry on the 1402CCCG1405 RNase T1 fragment with two
methyl groups from the TTHA1387 null mutant and com-
pared it with the triply methylated wild-type RNase T1
fragment (Fig. 2C). The clear w2 ions, as well as the less
intense z3 ions, display a 14.0 Da mass difference between
the two samples, showing that the methylations on
m4Cm1402 were not affected by inactivation of TTHA1387.
Tandem mass spectrometry was also performed on the
RNase T1 fragments appearing as a consequence of the lack
of methylations on C1400 and C1407 (data not shown).
As expected, the C1400-containing fragment revealed no
FIGURE 1. Secondary structure diagram of the 39 minor domain of
16S rRNA indicating the position of the three RsmF substrate
nucleotides.
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1585
indications of a methyl group, whereas
the RNase T1 fragment with C1407 ex-
hibited a fragmentation pattern corre-
sponding to the expected mass overlap
with an RNase T1 fragment of a different
sequence. In summary, our data lead us
to conclude that TTHA1387 encodes an
RsmF m5C methyltransferase responsible
for synthesizing m5C1400, m5C1404, and
m5C1407 in 16S rRNA of T. thermophilus.
The
above
results
left
us
with
TTHA0851 as the sole candidate for
the gene encoding the m5C967 methyl-
transferase. An approach conceptually
identical to that described above re-
vealed that disruption of TTHA0851
reduced the relevant RNase T1 fragment
by 14.0 Da (Supplemental Fig. 1A).
Since this fragment is methylated at
G966 and C967 in the wild-type strain
(Guymon et al. 2006), tandem mass
spectrometry was again performed (Sup-
plemental Fig. 1B), showing that only
the methyl group on C967 was absent. In
agreement with the suggested nomen-
clature for rRNA modifying enzymes
(Andersen and Douthwaite 2006), we
hereafter refer to TTHA0851 as rsmB
due to substrate specificity identical to
the originally identified enzyme from
E. coli (Gu et al. 1999).
Effect of temperature on growth
of the rsmF null mutant
One possible explanation for methyla-
tion of multiple sites by RsmF is that
such additional methyl groups improve
ribosomal function at elevated temper-
ature. To address this possibility, we ex-
amined the effect of temperature on
growth of the rsmF null mutant. Wild-
type T. thermophilus and the rsmF null
mutant were cocultured at three tem-
peratures and these cocultures were se-
rially subcultured for seven cycles of
24 h each. The proportion of wild-type
and rsmF null mutant cells in each mixed
culture was determined by spreading di-
lutions onto TEM plates with or without
kanamycin. After seven cycles, no differ-
ence in the relative proportions of the
wild-type and rsmF mutant was exhib-
ited at 70°C. However, at 60°C, the rsmF
null mutant constituted only around 5%
FIGURE 2. (Legend on next page)
Demirci et al.
1586
RNA, Vol. 16, No. 8
of the population, and at 80°C the rsmF null mutant was
unable to grow at all. Thus, methylation by RsmF appears to
facilitate growth at temperatures outside the optimal growth
temperature.
RsmF substrate preference
The T. thermophilus rsmB and rsmF genes were each cloned
into an E. coli expression plasmid in order to produce
proteins for X-ray crystallography and in vitro methylation
studies. The expression constructs were equipped with
C-terminal histidine6 tags to facilitate protein purification.
While we achieved a high expression level of RsmF, we were
unable to do so with RsmB despite a series of optimization
attempts. Consequently, in vitro substrate and structure
analyses were performed exclusively with RsmF.
E. coli RsmF requires the 30S ribosomal subunit as a
substrate when the activity is assayed in vitro (Andersen and
Douthwaite 2006). We assayed 70S ribosomes, 30S ribosomal
subunits, and 16S rRNA for the ability to serve as substrates
for methylation by RsmF in vitro. 16S rRNA subfragments of
z50 nt around the target sites were purified after the in vitro
assay and analyzed by mass spectrometry as described above.
In vitro methylation at 70°C showed an interesting but rather
complex substrate pattern. RsmF completely methylates
C1400 when either 16S rRNA or 30S subunits are used as
a substrate. It methylates C1404 to z35% with 16S rRNA
and completely with 30S subunits, and it produces only trace
amounts of methylation of C1407 with 16S rRNA and z75%
with 30S subunits (Fig. 3). There were no indications of
the 70S ribosome being a substrate in vitro. Curiously,
T. thermophilus RsmF expressed in an E. coli rsmF null
mutant almost completely methylated, in vivo, positions
C1400 and C1404, but not C1407 (data not shown).
X-ray crystal structures of RsmF
We determined the structure of T. thermophilus RsmF (456
amino acids) in three different crystal forms and in a com-
plex with cofactor AdoMet to up to 1.3 A˚ resolution (Figs.
4, 5). The structure was solved in space group P43 (data set
RsmF1, 1.4 A˚ resolution) by molecular replacement using
a search model generated with the program Modeller
(Eswar et al. 2008) from the catalytic domain of the RsmF
homolog YebU from E. coli (Pdb 2FRX) (Hallberg et al.
2006). The structures of the AdoMet-bound form in space
group P2 (RsmF2, 1.82 A˚ resolution), of the AdoMet-
bound form (RsmF3, 1.3 A˚ resolution), and of the apo-
form (RsmF4, 1.68 A˚ resolution) in space group P21212
were subsequently solved by molecular replacement with
the refined RsmF1 model. There are two molecules in the
asymmetric unit in space groups P43 and P2 and one mol-
ecule in space group P21212. Electron density is generally
well defined in all crystal forms. The majority of residues
(92.0%, 92.3%, 93.1%, and 92.6%) are in the most favored
region of the Ramachandran plot for RsmF1, RsmF2,
RsmF3, and RsmF4, respectively, and there are no residues
in the disallowed region. The final models consist of
residues 5–178, 194–198, and 201–456 and five additional
residues from the histidine6 affinity tag in both chains of
data set RsmF1; residues 2–456 and five affinity-tag
residues in both chains of data set RsmF2; residues 1–456
and six affinity-tag residues in data set RsmF3; and resi-
dues 1–456 and seven affinity-tag residues in RsmF4. The
N-terminal a-amino group was ordered in data sets RsmF3
and RsmF4 and contained additional electron density,
which we interpreted as N-(dihydroxymethyl)-L-methio-
nine, the hydrated form of N-formyl-methionine. Data
collection and refinement statistics are given in Table 1.
The overall structure of RsmF consists of a central
canonical class I methyltransferase catalytic domain with
additional N-terminal and C-terminal
domains (Figs. 4, 5). The catalytic do-
main is formed by a central seven-
stranded b-sheet that is flanked on both
sides by three helices of varying lengths.
An inserted region between strand b7
and helix a11 contains additional heli-
ces a9 and a10, which interact with the
two N-terminal helices a1 and a2. A
second inserted region following strand
b9 includes the short helices a13 and
FIGURE 2. (A) MALDI mass spectra of an RNase T1-digested 16S rRNA subfragment (pos.
1378–1432) from wild-type cells (upper panel) or from the TTHA1387 (putative rsmF) null
mutant (lower panel). Expected digestion products are labeled; fragments affected by the null
mutation are set in italics. (B) Expansion of the signals affected by the TTHA1387 null mu-
tation. The sequence and methylation status of the RNase T1 products are indicated. (C)
MALDI tandem mass spectrometry of the methylated RNase T1 fragment of 16S rRNA (pos.
1404–1407); wild-type cells (upper panel), TTHA1387 null mutant (lower panel). Mass
spectrometric fragments used to deduce the methylation status are labeled. The position of
the backbone fragments (nomenclature according to McLuckey et al. [1992]) in the sequence is
shown. MH+, precursor ion selected for fragmentation; C, cytosine; mC, methylated cytosine;
C>p, cytidine-2´-39-monophosphate; me, methyl group.
FIGURE 3. In vitro methylation with 30S ribosomal subunits or 16S
rRNA from the T. thermophilus rsmF null mutant as a substrate. Effect
on C1400-, C1404-, and C1407-harboring RNase T1 products. In
vitro methylated products are set in italics. *, artifact signal arising
from the enzyme preparation.
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1587
a14, which interact with helix a12. Furthermore, RsmF
contains three additional smaller domains, an N-terminal
domain consisting of a three-stranded b-sheet and two flank-
ing helices (Fig. 5B, colored in blue), and two C-terminal
domains consisting of four-stranded b-sheets and two or one
helix (Fig. 5B, colored in magenta and red).
Cofactor binding, substrate docking,
and conformational flexibility in the active site
The coordination of AdoMet in the T. thermophilus RsmF
active site is similar to that seen in other class I methyl-
transferases. However, the previously published structure of
E. coli RsmF did not contain the cofactor AdoMet in the
active site, precluding a direct comparison. Both the T.
thermophilus and E. coli RsmF cofactor-binding sites reveal
a new variation for the methyltransferase signature motif I
(Malone et al. 1995), with the highly conserved GxGxG
sequence replaced by 109AAAPG113. The combination of
three alanines and a proline results in a loop conformation
that is very similar to that observed in other methyltrans-
ferases with a GxGxG motif (e.g., RsmC) (Demirci et al.
2008a). In RsmF, the amide hydrogen atom of the last
glycine residue forms a hydrogen bond with the cofactor
carboxy group (Fig. 6). Other key interactions with AdoMet
are well conserved in RsmF. The cofactor adenine ring is
located in a mainly hydrophobic pocket lined by residues
Val134, Pro160, and Leu211. This pocket is open toward the
solvent. The adenine amino group is not specifically recog-
nized and interacts with solvent water molecules. The ribose
hydroxyl groups form hydrogen bonds with Glu133 and
Arg138, and the methionine amino group interacts with
Asp177. The AdoMet cofactor is bound in a cleft in the
RsmF active site, which suggests that substrate cytidine bases
are inserted into the active site in an unstacked conforma-
tion. Inspection of the electrostatic charge distribution re-
veals a large positively charged surface region, which would
be consistent with binding to an RNA surface and modifi-
cation of the substrate base in an unstacked orientation (Fig.
6C). To evaluate the possible orientation of a substrate base
in the active site, we performed computational docking
calculations with the program Dock6 (Lang et al. 2009). The
resulting positions of cytosine and m5C in the presence of
AdoMet in data set RsmF3 are highly similar to each other,
with m5C placed into the active site with its phosphate group
toward a positively charged pocket at the entrance of the
active site cleft (Fig. 6E). The position of the phosphate
group is close to a sulfate molecule that we observed in data
set RsmF1, providing further support for the results of the
docking calculation (Fig. 6F).
Interestingly, we observed that three active site segments
were disordered in data set RsmF1. These segments include
residues 179–193 (including helices a9 and a10 and the
catalytic Cys180), residues 199–200, and the N-terminal
residues 1–5, which interact with the first two segments (Fig.
6E,F, colored in green). We observed electron density for the
intervening residues 194–198, which formed a lattice con-
tact with a neighboring molecule. However, the position of
FIGURE 4. Structure-based sequence alignment of RsmF from T. thermophilus and E. coli. Secondary structure elements of T. thermophilus RsmF
are indicated on top. The color scheme for the secondary structure elements is as in Figure 5A. The position of the variant methyltransferase motif
I is marked with a red box. A flexible region observed in the active site is marked with a green box; residues interacting with the cofactor are
marked with orange boxes.
Demirci et al.
1588
RNA, Vol. 16, No. 8
these five residues was not related to their position in the
other three data sets, suggesting that the extended active
site region between residues 179 and 201 can reorient in the
RsmF structure. This observation suggests that this active
site region is dynamic, which may be important for sub-
strate binding at 72°C, the optimum growth temperature of
T. thermophilus.
DISCUSSION
Substrate recognition mechanisms
We have identified the two enzymes responsible for the
synthesis of the four m5C modifications of T. thermophilus
16S rRNA, and characterized the RsmF methyltransferase
responsible for synthesizing three of these. rRNA modifying
enzymes in bacteria are generally highly specific, with a
one-to-one association between the modifying enzyme and
the modification. A few cases of multitarget ribosome mod-
ifying enzymes have been reported (Helser et al. 1972;
Demirci et al. 2008b), but to our knowledge T. thermophi-
lus RsmF is the first rRNA methyltransferase found to mod-
ify three different nucleotides. Most ribosome modifying
enzymes probably recognize assembly intermediates, and
the data presented here are consistent with that notion.
T. thermophilus RsmF methylates C1400 and C1404 in
vitro using either 16S rRNA or 30S subunits as substrates,
whereas both E. coli (Andersen and Douthwaite 2006) and
T. thermophilus RsmF exclusively utilize 30S subunits as
substrates for methylation of C1407. This may reflect that
C1400 and C1404 methylations do not rely on the asso-
ciation of ribosomal proteins in order to be recognized by
T. thermophilus RsmF. C1407 methylation, in contrast,
depends on both rRNA and the ribosomal protein for the
recognition by RsmF in both T. thermophilus and E. coli.
More puzzling is the observation that T. thermophilus RsmF
does not methylate E. coli ribosomes in vivo on C1407. It
is perhaps worth noting that methylation of C1407 in the
T. thermophilus 30S ribosomal subunit in vitro was less
efficient than methylation of the other two positions,
indicating the need for a particular intermediate assembly
structure or for accessory factors. The only clear in vitro
FIGURE 5. Overall structure of RsmF. (A) Schematic representation of the position of the substrate bases in the 30S ribosomal subunit (Pdb
entry 2WRI) (Gao et al. 2009). C1400, C1404, and C1407 in helix 44 (green) are shown in stick representation with 5-methyl groups as pink
spheres. (B) Schematic stereo representation of the overall structure of RsmF. Secondary structure elements are in orange and yellow for the
catalytic domain, in salmon and blue for additional N-terminal domains, and in magenta and red for C-terminal domains. The flexible region in
the active site including helices a9 and a10 is indicated with an arrow and colored in green. (C) Topology diagram with secondary structure
elements colored as in B.
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1589
methylation observed at 37°C with T. thermophilus RsmF
was on C1400 with 16S rRNA as the substrate (data not
shown), which is not evidently related to the methylation
pattern in vivo in the heterologous system. It seems unlikely
that the aberrant methylation in the heterologous system
reflects species-specific differences in the mature 30S ribo-
somal subunit, given the extreme sequence and structural
conservation of the decoding site. Instead, it may reflect
differences in 30S subunit assembly in the two organisms,
necessary due to the large difference in growth temperature.
The four m5C residues in T. thermophilus 16S rRNA are
clustered in and around the functionally critical decoding
center at or close to sites of contact with tRNA, mRNA, and
EF-G (Ogle et al. 2002; Selmer et al. 2006; Gao et al. 2009).
m5C1400, m5C1404, and m5C1407 are located in the
subunit body, while m5C967 is located in the subunit head,
about 10 A˚ from m5C1400 (Wimberly et al. 2000). Exam-
ination of the 30S subunit crystal structure (Wimberly et al.
2000) indicates that the three bases methylated by RsmF are
situated in three distinct structural contexts, but provides
few clues to a common mode of substrate recognition.
C1400 is an unpaired base protruding from a sharply bent
segment of rRNA at the junction of helices 43 and 44, while
C1404 and C1407 are engaged in Watson–Crick pairs within
helix 44 (Fig. 4A). The C5 positions of the latter two bases
are not obviously accessible, such that RsmF would need to
approach them from the major groove side. While a flipping
of C1407 out of the helix via the minor groove could allow
access to this base, such a mechanism would be problematic
for C1404, whose minor groove side is packed against the
rest of the 30S subunit.
While m5C1404 and m5C1407 are about 11 A˚ apart
(Selmer et al. 2006), m5C1400 is quite distant from both of
these bases (about 21 and 30 A˚ , respectively). The pro-
truded conformation of m5C1400 in the mature 30S sub-
unit is due in part to base-pairing interactions between
adjacent bases and the 1500 region of 16S rRNA (C1399–
G1504 and G1401–C1501). As the 1500 region is one of the
last segments of 16S rRNA to be synthesized, C1400 could
potentially be positioned much closer to C1404 and C1407
in an assembly intermediate, prior to the formation of the
C1399–G1504 and G1401–C1501 base pairs. RsmF could
utilize a single binding mode to then access all three bases,
further facilitated by its flexible active site domain. Meth-
ylation of C1400 in mature 30S subunits would therefore
involve disruption of the adjacent base pairs. A complete
understanding of the recognition mechanism of these
enzymes will require high-resolution structural data on
assembly intermediate-enzyme complexes. Given the large
number of potential subunit assembly intermediates, pre-
cisely defining the physiological substrate for RsmB and
RsmF will be a formidable task.
TABLE 1. Data collection and refinement statistics
RsmF1
RsmF2
RsmF3
RsmF4
Data collectiona
AdoMet
AdoMet
Space group
P43
P2
P21212
P21212
Cell dimensions
a, b, c (A˚ )
71.0, 71.0, 186.7
66.0, 78.3, 108.1
89.7, 109.0, 51.0
89.8, 109.1, 50.8
a, b, g (°)
90, 90, 90
90, 107.1, 90
90, 90, 90
90, 90, 90
Resolution (A˚ )b
30–1.4 (1.55–1.40)
30–1.82 (1.89–1.82)
30–1.30 (1.34–1.30)
30–1.68 (1.74–1.68)
Rmerge
0.065 (0.59)
0.08(0.38)
0.058(0.36)
0.15 (0.49)
I/sI
29.3(2.15)
12.6 (2.04)
24.3 (2.05)
14.2 (1.73)
Completeness (%)
90.1 (72.6)
97.0(86.5)
95.6(66.3)
99.6 (97.8)
Redundancy
8.9 (5.4)
2.8(2.0)
5.2(2.1)
6.4 (3.9)
Refinement
Resolution (A˚ )
30–1.4 (1.42–1.40)
30–1.82 (1.84–1.82)
30–1.30 (1.32–1.30)
30–1.68 (1.69–1.68)
Number of reflections
161,955 (4356)
91,762 (2583)
119,490/2640
109,375 (3244)
Rwork/Rfree
0.169/0.189 (0.227/0.263)
0.162/0.194 (0.222/0.259)
0.177/0.191 (0.233/0.234)
0.173/0.192 (0.217/0.266)
Number of atoms
Protein
6766
7117
3598
3574
Ligand/ion
20
54
27
1
Water
1486
1285
856
665
B-factors
Protein
24.1
20.4
15.9
17.2
Ligand/ion
34.5
21.6
17.5
19.1
Water
38.7
36.3
33.6
33.6
RMSDs
Bond lengths (A˚ )
0.009
0.006
0.005
0.004
Bond angles (°)
1.22
1.04
1.14
0.94
aOne crystal used for each data set.
bThe highest resolution shell is shown in parentheses.
Demirci et al.
1590
RNA, Vol. 16, No. 8
Structural comparison of RsmF with related
methyltransferases
A database search with Dali (Holm et al. 2008) confirmed
the structural similarity of the T. thermophilus and E. coli
(PDB 2FRX) (Hallberg et al. 2006) RsmF homologs, which
superimpose with a root-mean-square deviation (RMSD)
of 1.6 A˚ for 342 Ca atoms (Fig. 7A) and
are the only two structures in the Pro-
tein Database with this domain organi-
zation. Even so, substantial structural
differences are observed in most of the
loop regions and for the long connect-
ing loop between the methyltransferase
domain and the first C-terminal domain.
While in the active site, the positions
of residues in the cofactor-binding site
and of the two cysteine residues are
conserved (Fig. 7B), there are a number
of positively charged residues (Arg30,
Arg190, Arg194, His195, and Arg203) in
the T. thermophilus structure that are
absent from the E. coli enzyme (Fig. 7B).
Three of these are located in the flexible
region, and the combination of a posi-
tive charge and flexibility close to the
active site is suggestive of a functional
contribution of this region to the mul-
tisite specificity of T. thermophilus RsmF
(Figs. 6C,D, 7B). Methylation of three
rRNA positions may require an increase
in the enzyme’s structural dynamics in
order to accommodate the 30S subunit
in slightly different orientations. Similar
observations have been made for other
multi-site-specific methyltransferases in-
cluding KsgA, which modifies two adja-
cent adenosines in the 30S ribosomal
subunit (O’Farrell et al. 2004; Demirci
et al. 2009), and the PrmA ribosomal
protein methyltransferase, which under-
goes dramatic interdomain movements
to modify multiple lysine residues and the
N-terminal a-amino group on the same
substrate protein (Demirci et al. 2007,
2008b).
The second C-terminal domain in
RsmF is related to the RNA-binding
PUA
(pseudouridine
synthase
and
archaeosine transglycosylase) domains
(Perez-Arellano et al. 2007). The RlmI
methyltransferase,
which
produces
m5C1962 in 23S rRNA (Purta et al.
2008) also contains a PUA domain
(Sunita et al. 2008), although it is
N-terminal to the catalytic methyltransferase domain and
in a different orientation. PUA domains contain six
b-strands, which form a central pseudobarrel closed by
a short 310-helix. A comparison of the C-terminal domain
in RsmF with a typical PUA domain in archaeosine trans-
glycosylase (ArcTGT, Pdb entry 1J2B) reveals that the
central fold is similar (53 Ca atoms align with an RMSD
FIGURE 6. Substrate docking and conformational flexibility. (A) Cofactor-binding site in
RsmF. Bound AdoMet is shown in blue sticks. Hydrogen bonds to coordinating residues are
indicated. (B) Final sA-weighted 2mFO-DFC electron density map of the cofactor-binding site
(data set RsmF2) contoured at the 1s level. (C,D) Comparison of the electrostatic surface
charge distribution between RsmF from T. thermophilus and E. coli. The location of the
C-terminal domains and of the flexible region (labeled ‘‘FLEX’’) is indicated with circles.
AdoMet and docked m5C are shown as sticks. AdoMet from the T. thermophilus structure is
shown with the E. coli structure for comparison. (E) The modeled position of m5C docked into
the active site of RsmF. m5C and AdoMet are shown in tan and blue sticks, respectively.
Residues in the flexible region are shown in green. (F) The active site region in data set RsmF1.
A sulfate ion is observed close to the position of the m5C phosphate group. Residues 194–198 in
the flexible region interacting with a neighboring RsmF molecule are indicated with an arrow.
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1591
of 1.8 A˚ ), but that several connecting loop regions are
substantially shorter and two a-helices and one b-strand of
the pseudobarrel are absent. Thus, this PUA-like domain
differs considerably from typical PUA domains. However,
the similarity to RNA-binding PUA domains and the
positive surface charge distribution observed in both RsmF
structures are suggestive for a conserved function of the
PUA-like domain in RNA recognition.
The next most closely related structure is E. coli RsmB
(474 residues, PDB 1SQF) (Foster et al. 2003). A total of
264 Ca atoms can be aligned with an RMSD of 1.5 A˚
between RsmB and RsmF. Both enzymes retain the same
organization for the N-terminal domains and the core
methyltransferase domain. However, an additional 140-
residue N-terminal RNA-binding domain provides sub-
strate specificity to RsmB, whereas the two C-terminal
domains following the core methyltransferase domain (160
residues) are likely to determine substrate recognition by
RsmF. Thus, these two enzymes have evolved substrate
specificity via acquisition of additional,
unrelated RNA recognition domains.
While there are as yet no enzyme–
substrate complexes for rRNA m5C-
methyltransferases, insights into the RsmF
catalytic mechanism can be gleaned from
a comparison with the RlmD and TrmA
m5U methyltransferases in covalent in-
termediate complexes with RNA oligonu-
cleotides (Lee et al. 2005; Alian et al.2008).
DNA and RNA m5C methyltransferases
use a thiol from a catalytic cysteine residue
to attack the six-position of the pyrimi-
dine base to activate the five-position for
methyl group transfer (Liu and Santi
2000). Cys180 and Cys230 in RsmF are
positioned equivalently to the catalytic
Cys324 and the catalytic base Glu358 of
TrmA. The substrate nucleotides insert
into TrmA and RsmF in unrelated di-
rections, consistent with a lack of struc-
tural homology outside the methyltrans-
ferase domains. Nevertheless, the C5
positions of the pyrimidine rings and of
the 5-methyl carbons are quite similar
with respect to the catalytic cysteine resi-
dues and the AdoMet cofactor. The same
structural homology of the active site
geometry can be observed in comparison
with the RlmD methyltransferase (Sup-
plemental Fig. 2; Lee et al. 2005).
A possible origin of T. thermophilus
RsmF
T. thermophilus RsmF shows the highest
similarities with proteins from close relatives, namely,
Thermus aquaticus and two Meiothermus species of the
Thermaceae family. Remarkably, the next highest similarities
(BLAST scores between 267 and 359) are with the NOL1/
NOP2/Sun proteins from the Gram-positive Firmicutes
phylum (Supplemental Fig. 3). This similarity, together
with the fact that the Thermaceae family and most members
of the Firmicutes identified in Supplemental Figure 3 are
thermophilic, suggest that rsmF has undergone horizontal
transfer between the Thermaceae family and members of the
Firmicutes phylum. Thus, we speculate that this version of
the RsmF protein, which catalyzes methylation of three
cytidines, may be adaptive for existence in thermally
challenging environments. The effect of the loss of methyl-
ation by RsmF on growth at different temperatures is
consistent with this notion. Our hypothesis of horizontal
transfer of rsmF predicts that RsmF of other members of the
Thermaceae family and members of the Firmicutes phylum
will also be found to introduce multiple m5C modifications.
FIGURE 7. Comparison with other methyltransferases. (A) Differences between the overall
structures of RsmF from T. thermophilus (orange) and E. coli (green). AdoMet bound in the T.
thermophilus structure is shown as blue sticks. (B) Comparison of the active site region in both
enzymes. Residues in the cofactor-binding site and in the flexible region in the T. thermophilus
enzyme are shown as sticks. (C) Comparison between RsmF (orange) and a substrate complex
structure of the TrmA methyltransferase (cyan/blue). (D) Comparison of the active site region
illustrating the differences in the insertion direction of the substrate base in RsmF (m5C in
light orange) and in TrmA (m5U in cyan).
Demirci et al.
1592
RNA, Vol. 16, No. 8
MATERIALS AND METHODS
Cloning of the T. thermophilus rsmB and rsmF genes
The T. thermophilus HB8 loci TTHA0851 (GenBank accession
number BAD70674) and TTHA1387 (GenBank accession number
BAD71210) were PCR amplified from genomic DNA and purified
via the High Pure PCR Template Preparation Kit (Roche). The
100 mL PCRs contained 150 ng DNA, 10 mM of each primer,
10 mM dNTP, 1 unit Phusion DNA polymerase (Finnzymes), and
1x Phusion HF buffer. Primers for rsmB amplification were 59-CC
CTGGACATATGAGGGCCGG-39 and 59-GGCCAAGATCTTGCC
TGAGAG-39, and the temperature cycling was as follows: 98°C/30
sec; 30X (98°C/10 sec, 59°C/20 sec, and 72°C/36 sec); and 72°C/420s.
Primers for rsmF amplification were 59-GCTAGGGTACACATA
TGCTGCCC-39 and 59-GCACGGGGGTGAGATCTAAGCCC-39,
and the temperature cycling was as follows: 98°C/30 sec; 30X
(98°C/10 sec, 62°C /20 sec, 72°C/42 sec); and 72°C/420 sec. The
desired PCR products were purified from agarose gels using the
GFX PCR purification kit (GE Healthcare). The PCR fragments
were digested with NdeI and BglII and inserted into the expression
vector pLJ102 (Andersen and Douthwaite 2006), generating
isopropyl-1-thio-b-D-galactopyranoside (IPTG)-inducible genes
for the recombinant proteins with a C-terminal histidine6 tag.
The constructs (designated pLJ102-RsmB and pLJ102-RsmF) were
used to transform an rsmF-deletion derivative of E. coli CP79
(Andersen and Douthwaite 2006).
Deletion of the T. thermophilus rsmB and rsmF genes
Constructs for inactivation of the T. thermophilus rsmB and rsmF
genes were made by inserting the gene for a heat tolerant
kanamycin (HTK) nucleotidyltransferase (Hoseki et al. 1999) into
the methyltransferase parts of either pLJ102-RsmB or pLJ102-
RsmF. The htk part of pUC18-htk (Hashimoto et al. 2001) was
amplified by PCR with primers that introduced an upstream AvrII
site and a downstream SacI site into the product for later disrup-
tion of rsmB. For rsmF disruption, the PCR primers introduced
SacI restriction enzyme sites both upstream of and downstream
from the htk gene. These sites were used to insert the PCR prod-
ucts into pLJ102-RsmB and pLJ102-RsmF to form the plasmids
pLJ102-RsmBThtk and pLJ102-RsmFThtk, which were propa-
gated in the E. coli strain Top10 (Invitrogen). T. thermophilus
HB8 was transformed with pLJ102-RsmBThtk or pLJ102-
RsmFThtk selecting for kanamycin resistance as described by
others (Hashimoto et al. 2001; Cameron et al. 2004). Kanamycin-
resistant transformants were restreaked twice. Gene disruptions
were verified by PCR with primers distal to the interrupted rsmB
or rsmF genes on genomic DNA; resulting PCR products were
characterized by sequencing.
Growth competition assays
Wild-type and rsmF null mutant liquid cultures were grown at
70°C to saturation, then equal numbers of cells from each were
mixed and incubated in 5 mL TEM medium at 60°C, 70°C, or
80°C. After growth for 24 h, 100 mL of the 60°C culture, 10 mL of
the 70°C culture, and 1000 mL of the 80°C culture were trans-
ferred to a fresh 5-mL medium and incubated at the respective
temperatures for another 24 h. This was repeated in independent
triplicates for seven cycles. Samples of 1 mL were collected at each
dilution and half was plated on TEM plates without antibiotic and
the other half was plated on TEM plates with 30 mg/mL
kanamycin. The plates were incubated at 70°C.
Purification of T. thermophilus ribosomal subunits
and ribosomes
T. thermophilus culture (1 L) was grown in TEM media (contain-
ing 30 mg/mL of kanamycin when appropriate) with shaking at
70°C to an OD600 = 0.6. Cells were harvested and washed once
with 100 mL of buffer A (10 mM NH4Cl, 20 mM MgCl2, 100 mM
KCl, and 10 mM Tris-HCl [pH 7.5]), then suspended in 10 mL of
buffer A, and disrupted by sonication. The lysate was cleared by
centrifugation twice in a Beckman JA20 rotor at 16,000 rpm for
10 min at 4°C. Crude ribosomes were collected by centrifugation
in a Beckman Ti50 rotor at 19,000 rpm for 19 h at 4°C, and
dissolved in buffer A. 70S ribosomes were obtained by centrifu-
gation of 100 A260 units of crude ribosomes through a 10%–40%
sucrose gradient (200 mM NH4Cl, 20 mM MgCl2, 20 mM Tris-
HCl [pH 7.5]) in a Beckman SW28 rotor at 20,000 rpm for 18 h at
4°C. Fractions containing intact 70S ribosomes were pooled and
concentrated by centrifugation in a Beckman Ti50 rotor at 40,000
rpm for 22 h at 4°C, washed, and dissolved in buffer A, and stored
at 80°C.
50S and 30S ribosomal subunits were obtained by adjusting
100 A260 units of crude ribosomes (10 mM NH4Cl, 2 mM MgCl2,
100 mM KCl, and 10 mM Tris-HCl [pH 7.5]) and passing
through a 5%–20% sucrose gradient (200 mM NH4Cl, 1 mM
MgCl2 and 20 mM Tris-HCl [pH 7.5]) in a Sorvall AH-629 rotor
at 20,000 rpm for 18 h at 4°C. After pooling of the relevant
fractions, the subunits were adjusted to 10 mM MgCl2 and
pelleted by centrifugation in a Beckman Ti50 rotor at 40,000
rpm for 22 h at 4°C, washed with and dissolved in 10 mM NH4Cl,
10 mM MgCl2, 100 mM KCl, and 10 mM Tris-HCl (pH 7.5), and
stored at 80°C.
Isolation of 16S rRNA and subfragments
from T. thermophilus and E. coli
Water (400 mL) was added to 100 mL of 30S ribosomal subunits
and the rRNA was extracted with 500 mL phenol, phenol/
chloroform, and chloroform. rRNA was ethanol precipitated
and dissolved in water. Purification of 16S rRNA subfragments
was performed as previously described (Andersen et al. 2004).
Briefly, 16S rRNA was hybridized to an excess of oligodeoxynu-
cleotide complementary to either the region 944–990 or the region
1378–1432. Single-stranded nucleic acids were digested with
Mung Bean Nuclease and RNase A. The resulting mixture was
separated on a polyacrylamide gel. Bands were visualized by ethid-
ium bromide staining, excised, and eluted.
E. coli CP79 with the endogenous rsmF inactivated, but com-
plemented with the T. thermophilus homolog on the plasmid
pLJ102-RsmF, were grown at 37°C to an OD450 = 0.45 in 200 mL
of LB medium containing 100 mg/L of ampicillin. RsmF expres-
sion was induced by addition of IPTG to 1 mM, and incubation
for another 3 h. Cells were harvested by centrifugation at 4°C,
washed in 100 mL TMN buffer (50 mM Tris-HCl [pH 7.8],
10 mM magnesium acetate, 100 mM NH4Cl), and resuspended in
2 mL TMN buffer prior to lysis by sonication (7 3 30 sec on ice)
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1593
and removal of debris by centrifugation (10 min/14,000 rpm/4°C/
microcentrifuge). Total RNA was recovered from the supernatant
by phenol extraction and ethanol precipitation. A 16S rRNA
subfragment was isolated as described above using an oligodeoxy-
nucleotide complementary to the region 1378–1432.
In vitro methylation
Reactions contained 50 pmol of 16S rRNA, 30S subunits, or 70S
ribosomes from the T. thermophilus TTHA1387 null mutant as the
substrate in a total volume of 100 mL (containing 100 mM NH4Cl,
10 mM MgCl2, 40 mM Hepes [pH 7.5]), 6 mM b-mercaptoetha-
nol, and 10% glycerol (prepared as a two times concentrated stock
solution), 1.5 mM S-adenosyl methionine, and 2 mg of recombi-
nantly expressed RsmF (see below).
For the reaction at 70°C, water and stock buffer were mixed and
left at room temperature for 15 min. Then a substrate, an enzyme
and S-adenosyl methionine were added and incubated at 70°C for
1 h. The 37°C reaction was started by mixing water and buffer
followed by 15 min at room temperature; the substrate was added
and the mixture transferred to 50°C for 5 min. After cooling to
37°C, S-adenosyl methionine and an enzyme were added and the
incubation continued for 1 h. Reactions were stopped by phenol/
chloroform extraction and the rRNA was recovered by ethanol
precipitation before purification of 16S rRNA subfragments as
described above. Control reactions without enzyme or S-adenosyl
methionine were carried out in all instances.
RNase T1 digestion and mass spectrometry
A purified 16S rRNA subfragment (1–2 pmol) was incubated with
2 units RNase T1 (Roche) and 50 mM 3-hydroxypicollinic acid
(3-HPA) in a total volume of 2 mL for 4 h at 37°C. MALDI mass
spectrometry was performed either on an ABI voyager STR in-
strument or a Waters Q-TOF MALDI instrument; MALDI tan-
dem mass spectrometry was done on a Waters Q-TOF MALDI
instrument. All spectra were recorded in positive ion mode using
3-HPA as the matrix. Experimental details were as previously de-
scribed (Douthwaite and Kirpekar 2007).
Protein expression and purification for crystallization
E. coli BL21 (DE3) (Invitrogen) containing pLJ102-RsmF was
grown to midlog phase in LB media at 37°C in the presence of
200 mg/mL ampicillin. Protein expression was induced at 20°C
with 400 mM IPTG. Cells were pelleted after 18 h by centrifuga-
tion at 4000 rpm for 20 min at 4°C and lysed by ultrasonication
on ice in a buffer containing 20 mM Tris-HCl (pH 8.5), 300 mM
NaCl, 5 mM b-mercaptoethanol, 0.1% Triton X-100, and 5%
glycerol. Cell debris and membranes were pelleted by centrifuga-
tion at 11,000 rpm for 30 min at 4°C. The soluble E. coli proteins
were precipitated by heat treatment at 65°C for 30 min and
pelleted by centrifugation at 11,000 rpm at 4°C for 30 min.
Soluble C-terminally hexahistidine-tagged T. thermophilus RsmF
was further purified by affinity chromatography with nickel-
nitrilotriacetic acid resin (Qiagen). Untagged proteins were re-
moved with buffer containing 20 mM Tris-HCl (pH 8.5), 250 mM
NaCl, and 1 mM imidazole (pH 8.5). Recombinant RsmF was
then eluted with the same buffer containing 150 mM imidazole.
The protein was then purified by cation exchange chromatogra-
phy (SP) (GE Healthcare) at pH 8.5, using a linear gradient of
10 mM to 1 M NaCl concentration. RsmF fractions were pooled
and concentrated and applied to a size-exclusion S200 column (GE
Healthcare) equilibrated with a buffer containing 20 mM Tris-HCl
(pH 8.5) and 200 mM NaCl. Purified RsmF was concentrated to
13 mg/mL for crystallization trials. The C-terminal hexahistidine
tag was not removed for crystallization. For the production of
selenomethionyl proteins, the expression construct was trans-
formed into B834 (DE3) cells (Novagen). The bacterial growth
was carried out in defined LeMaster medium (Hendrickson et al.
1990), and the protein was purified using the same protocol as for
the unmodified protein. To form the RsmF-AdoMet complex,
purified RsmF was mixed with 4 mM AdoMet incubated at 60°C
for 15 min and slowly cooled to room temperature before
performing crystallization experiments.
Crystallization of RsmF
All crystals were obtained using the microbatch technique under
oil at 4°C. To obtain the RsmF1 crystal form, 1 mL of protein
solution was mixed with the reservoir solution containing 20%
(w/v) PEG3350 and 200 mM sodium sulfate decahydrate (pH
6.6). Initial crystals grew over the course of 1–2 wk with maxi-
mum dimensions of 0.3 3 0.3 3 0.2 mm. To obtain the RsmF2
crystal form, 1 mL of the RsmF–AdoMet complex was mixed with
the reservoir solution containing 200 mM NaCl, 12% w/v
PEG8000 and 100 mM HEPES-KOH (pH7.5). Initial crystals
grew over the course of 2–3 wk with maximum dimensions of
0.1 3 0.4 3 0.4 mm. To obtain the RsmF3 crystal form, 1 mL of
the RsmF–AdoMet complex was mixed with the reservoir solution
containing 10% w/v PEG1000, 200 mM NaCl, and 100 mM Tris-
HCl (pH 8.5). The initial crystals grew over the course of 1–2 wk
with maximum dimensions of 0.05 3 0.3 3 0.4 mm. To obtain
the RsmF4 crystal form, 1 mL of the RsmF–AdoMet complex
solution was mixed with a reservoir solution containing 160 mM
magnesium chloride hexahydrate, 80 mM Tris-HCl (pH 8.5), and
24% w/v PEG4000. Initial crystals grew over the course of 1–2 wk
with maximum dimensions of 0.05 3 0.3 3 0.3 mm. RsmF1
crystals were gradually dehydrated by increasing the PEG3350 to
30% w/v and then cryoprotected in a mother liquor supplemented
with 25% v/v glycerol and then flash-frozen by being plunged into
liquid nitrogen. RsmF2 crystals were cryoprotected in a mother
liquor supplemented with 20% v/v ethylene glycol and then flash-
frozen by being plunged into liquid nitrogen. RsmF3 crystals
were cryoprotected by gradually increasing the concentration of
PEG1000 to 30% and then flash-frozen by being plunged into
liquid nitrogen. RsmF4 crystals were cryoprotected in a mother
liquor supplemented with 20% glycerol and then flash-frozen by
being plunged into liquid nitrogen.
Data collection
X-ray diffraction data for RsmF1, RsmF2, and RsmF4 crystals
were collected on a MAR CCD detector at the X4C beamline of
the National Synchrotron Light Source in Brookhaven at a wave-
length of 0.979 A˚ and 180°C. Diffraction data for RsmF3 crystals
were collected on an ADSC CCD detector at the X4A beamline of
the National Synchrotron Light Source in Brookhaven at a wave-
length of 0.979 A˚ and 180°C. Diffraction data for RsmF1 in
space group P43 were collected to 1.4 A˚ resolution with cell
Demirci et al.
1594
RNA, Vol. 16, No. 8
dimensions a = 71.0 A˚ , b = 71.0 A˚ , and c = 186.7 A˚ . Diffraction
data to 1.82 A˚ for RsmF2 were collected in space group P2 with
cell dimensions a = 66.0 A˚ , b = 78.3 A˚ , and c = 108.1 A˚ .
Diffraction data to 1.29 A˚ for RsmF3 were collected in space group
P21212 with cell dimensions a = 89.7 A˚ , b = 109.0 A˚ , and c = 51.0
A˚ . Diffraction data to 1.68 A˚ for RsmF4 were collected in space
group P21212 with cell dimensions a = 89.8 A˚ , b = 109.1 A˚ , and c =
50.8 A˚ . A single crystal was used for each data set. The diffraction
images were processed and scaled with the HKL2000 package
(Otwinowski and Minor 1997). The data processing statistics are
summarized in Table 1.
Structure determination and refinement
The RsmF structure was solved by molecular replacement with the
program Phaser (McCoy et al. 2007) from the CCP4 program
suite (Bailey 1994) in space group P43 to 1.4 A˚ resolution (data set
RsmF1). The initial search model was built with the program
Modeller (Eswar et al. 2008) from the catalytic domain of E. coli
YebU (Pdb code 2FRX). After the placement of two RsmF
catalytic domains in the asymmetric unit and the initial re-
finement with Refmac (Murshudov et al. 1997), the model was
further rebuilt with ARP/wARP (Langer et al. 2008). The resulting
model was 90% complete and manually checked and completed
with Coot (Emsley and Cowtan 2004). Final crystallographic re-
finement was performed with the program Phenix (Adams et al.
2002). The other crystal forms were subsequently solved by
molecular replacement. The atomic coordinates from the RsmF4
model were then used for initial refinement of the RsmF–AdoMet
complex structure in space group P21212 (RsmF3). There are two
molecules in the asymmetric unit in data sets RsmF1 and RsmF2,
and one molecule in RsmF3 and RsmF4. The crystallographic
R/Rfree factors are 0.17/0.19, 0.16/0.19, 0.18/0.19, and 0.17/0.19
for the four data sets: RsmF1, RsmF2, RsmF3, and RsmF4,
respectively. The stereochemical quality of the model was assessed
with Procheck (Laskowski et al. 1993). The Ramachandran sta-
tistics (most favored/additionally allowed/generously allowed/
disallowed) are 91.9%/8.1%/0.0%/0.0% for RsmF1, 91.9%/8.1%/
0.0%/0.0% for RsmF2, 93.6%/6.4%/0.0%/0.0% for RsmF3, and
92.5%/7.5%/0.0%/0.0% for RsmF4. The refinement statistics are
summarized in Table 1. Figures were generated using Pymol
(DeLano 2002).
Atomic coordinates
Coordinates and structure factors have been deposited in the
Protein Data Bank with accession codes 3M6U, 3M6V, 3M6W,
and 3M6X for data sets RsmF1, RsmF2, RsmF3, and RsmF4,
respectively.
SUPPLEMENTAL MATERIAL
Supplemental material can be found at http://www.rnajournal.org.
ACKNOWLEDGMENTS
We thank John Schwanof and Randy Abramowitz for access to the
X4A and X4C beamlines at the National Synchrotron Light Source.
This work was supported by grants GM19756 and GM19756-37S1
from the National Institutes of Health.
Received January 14, 2010; accepted April 26, 2010.
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3M6W
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Multi-site-specific 16S rRNA methyltransferase RsmF from Thermus thermophilus in space group P21212 in complex with S-Adenosyl-L-Methionine
|
Multi-site-specific 16S rRNA methyltransferase RsmF
from Thermus thermophilus
HASAN DEMIRCI,1,3 LINE H.G. LARSEN,2,3 TRINE HANSEN,2 ANETTE RASMUSSEN,2 ASHWIN CADAMBI,1
STEVEN T. GREGORY,1 FINN KIRPEKAR,2 and GERWALD JOGL1
1Department of Molecular Biology, Cell Biology and Biochemistry, Brown University, Providence, Rhode Island 02912, USA
2Department of Biochemistry and Molecular Biology, University of Southern Denmark, 5230 Odense M, Denmark
ABSTRACT
Cells devote a significant effort toward the production of multiple modified nucleotides in rRNAs, which fine tune the ribosome
function. Here, we report that two methyltransferases, RsmB and RsmF, are responsible for all four 5-methylcytidine (m5C)
modifications in 16S rRNA of Thermus thermophilus. Like Escherichia coli RsmB, T. thermophilus RsmB produces m5C967. In
contrast to E. coli RsmF, which introduces a single m5C1407 modification, T. thermophilus RsmF modifies three positions,
generating m5C1400 and m5C1404 in addition to m5C1407. These three residues are clustered near the decoding site of the
ribosome, but are situated in distinct structural contexts, suggesting a requirement for flexibility in the RsmF active site that is
absent from the E. coli enzyme. Two of these residues, C1400 and C1404, are sufficiently buried in the mature ribosome
structure so as to require extensive unfolding of the rRNA to be accessible to RsmF. In vitro, T. thermophilus RsmF methylates
C1400, C1404, and C1407 in a 30S subunit substrate, but only C1400 and C1404 when naked 16S rRNA is the substrate. The
multispecificity of T. thermophilus RsmF is potentially explained by three crystal structures of the enzyme in a complex with
cofactor S-adenosyl-methionine at up to 1.3 A˚ resolution. In addition to confirming the overall structural similarity to E. coli
RsmF, these structures also reveal that key segments in the active site are likely to be dynamic in solution, thereby expanding
substrate recognition by T. thermophilus RsmF.
Keywords: rRNA methyltransferase; 5-methylcytidine; RsmB; RsmF; RNA mass spectrometry
INTRODUCTION
Ribosomal RNAs (rRNAs) are post-transcriptionally mod-
ified in all three domains of life, and many modifications
are phylogenetically conserved. Most modifications are
located in functionally important regions of the ribosome,
where they probably act to fine tune protein synthesis (Agris
2004; Gustilo et al. 2008). Complete modification maps of
bacterial 16S rRNAs have been determined for only a hand-
ful of species, and among these are the enteric bacterium
Escherichia coli and the extremely thermophilic bacterium
Thermus thermophilus (Guymon et al. 2006). Despite the
large phylogenetic divergence of these two organisms, their
ribosome modification patterns are quite similar. Of the 11
E. coli and 14 T. thermophilus 16S rRNA modifications,
eight are identical. This suggests a set of common functional
requirements conserved since divergence from their last
common ancestor, and also suggests common recognition
mechanisms among their modifying enzymes.
For most ribosome modifications, a single enzyme recog-
nizes and modifies a single site. However, there exist nota-
ble exceptions. Among these are dimethylation of two adja-
cent adenosines in 16S rRNA by KsgA (Helser et al. 1972);
pseudouridylation of three adjacent residues in tRNAs by
TruA (Hur and Stroud 2007); pseudouridylation of several
tRNA residues by Pus1 (Motorin et al. 1998), Pus2 (Behm-
Ansmant et al. 2007), or Pus7 (Behm-Ansmant et al. 2003);
or methylation of four tRNA positions by Saccharomyces
cerevisiae Trm4 (Motorin and Grosjean 1999). Even with
these multi-site-specific enzymes, however, homologs from
various species generally modify the same residues.
E. coli 16S rRNA contains two 5-methylcytidine (m5C)
residues, located in or near the highly conserved decoding
3These authors contributed equally to this work.
Abbreviations: rRNA, ribosomal RNA; AdoMet, S-adenosyl-L-methionine;
m5C, 5-methyl-cytidine; MALDI mass spectrometry, matrix-assisted laser
desorption ionization mass spectrometry.
Reprint requests to: Gerwald Jogl, Department of Molecular Biology,
Cell Biology and Biochemistry, Brown University, Box G-E129, Provi-
dence, RI 02912, USA; e-mail: Gerwald_Jogl@brown.edu; fax: (401)
863-6114; or Finn Kirpekar, Department of Biochemistry and Molecular
Biology, University of Southern Denmark, Campusvej 55, 5230 Odense M,
Denmark; e-mail: f.kir@bmb.sdu.dk; fax: (+45) 65502467.
Article published online ahead of print. Article and publication date
are at http://www.rnajournal.org/cgi/doi/10.1261/rna.2088310.
1584
RNA (2010), 16:1584–1596. Published by Cold Spring Harbor Laboratory Press. Copyright 2010 RNA Society.
center of the 30S subunit (Fig. 1). An m5C967 modification
is produced by RsmB (also called Fmu), while an m5C1407
modification is produced by RsmF, formerly known as
YebU (Andersen and Douthwaite 2006). T. thermophilus
16S rRNA contains m5C967 and m5C1407, as well as two
additional m5C nucleotides, m5C1400 and m5C1404 (E. coli
rRNA numbering used throughout) (Guymon et al. 2006).
While the m5C967 and m5C1407 modifications are pre-
sumably produced by RsmB and RsmF homologs, respec-
tively, the source of the two additional m5C residues has
been unknown. Here we demonstrate that T. thermophilus
RsmF is a multi-site-specific methyltransferase and, in
contrast to the single-site-specific E. coli RsmF, is respon-
sible for the synthesis of three modifications: m5C1407,
m5C1400, and m5C1404. We also demonstrate that RsmB is
responsible for the synthesis of m5C967 in T. thermophilus
as well as is in E. coli, thereby accounting for all four m5C
modifications of 16S rRNA. We present crystal structures
of T. thermophilus RsmF up to 1.3 A˚ resolution that reveal
a dynamic region in the active site that is absent from the E.
coli RsmF structure, providing a possible explanation for
the expanded recognition capacity of the T. thermophilus
methyltransferase.
RESULTS
Identification of T. thermophilus 16S rRNA
m5C-methyltransferases
With the E. coli RsmB and RsmF protein sequences as
queries, we used conventional BLAST searches (Altschul
et al. 1990) to identify potential homologs encoded by the
T. thermophilus HB8 genome (data not shown). Both RsmB
and RsmF have the highest similarity to the T. thermophilus
protein encoded by TTHA1387 (BLAST scores of 106 and
190, respectively) and second-highest similarity to the pro-
tein encoded by TTHA0851 (BLAST scores of 93 and 81,
respectively). The simplest interpretation of these results is
that TTHA1387 encodes RsmF, responsible for methylation
of C1407, leaving TTHA0851 as the most likely candidate
for the gene encoding RsmB, responsible for methylation of
C967. The similarities of the two E. coli enzymes with other
T. thermophilus proteins were far too low to reveal potential
candidates responsible for methylation of C1400 and C1404.
We next constructed T. thermophilus strains in which
either TTHA0851 or TTHA1387 was inactivated by the
homologous recombination and insertion of a heat stable
kanamycin-resistance gene. 16S rRNA was isolated from
these null mutants and subfragments of z50 nucleotides
(nt) around the regions of interest were further purified,
digested with RNase T1, and analyzed by MALDI mass
spectrometry (Fig. 2). Comparison of the TTHA1387 null
mutant to wild-type T. thermophilus HB8 indicates three
clear differences, each corresponding to the disappearance
of a methyl group (z14.0 Da). The RNase T1 digestion
fragment harboring m5C1407, the nucleotide methylated
by RsmF in E. coli, is absent in the null mutant, indicating
that TTHA1387 is indeed rsmF. The predicted RNase T1
fragment reduced by 14.0 Da is obscured by another RNase
T1 fragment that is present in both the wild-type and
TTHA1387 null mutants (Fig. 2B). Unexpectedly, two addi-
tional RNase T1 fragments are also reduced by 14.0 Da.
One of these contains C1400 while the other contains
C1404. This latter RNase T1 fragment from wild-type
T. thermophilus contains three methyl groups, two on
m4Cm1402 and one on m5C1404 (Guymon et al. 2006),
preventing an unambiguous identification of the missing
methyl group. We therefore performed tandem mass spec-
trometry on the 1402CCCG1405 RNase T1 fragment with two
methyl groups from the TTHA1387 null mutant and com-
pared it with the triply methylated wild-type RNase T1
fragment (Fig. 2C). The clear w2 ions, as well as the less
intense z3 ions, display a 14.0 Da mass difference between
the two samples, showing that the methylations on
m4Cm1402 were not affected by inactivation of TTHA1387.
Tandem mass spectrometry was also performed on the
RNase T1 fragments appearing as a consequence of the lack
of methylations on C1400 and C1407 (data not shown).
As expected, the C1400-containing fragment revealed no
FIGURE 1. Secondary structure diagram of the 39 minor domain of
16S rRNA indicating the position of the three RsmF substrate
nucleotides.
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1585
indications of a methyl group, whereas
the RNase T1 fragment with C1407 ex-
hibited a fragmentation pattern corre-
sponding to the expected mass overlap
with an RNase T1 fragment of a different
sequence. In summary, our data lead us
to conclude that TTHA1387 encodes an
RsmF m5C methyltransferase responsible
for synthesizing m5C1400, m5C1404, and
m5C1407 in 16S rRNA of T. thermophilus.
The
above
results
left
us
with
TTHA0851 as the sole candidate for
the gene encoding the m5C967 methyl-
transferase. An approach conceptually
identical to that described above re-
vealed that disruption of TTHA0851
reduced the relevant RNase T1 fragment
by 14.0 Da (Supplemental Fig. 1A).
Since this fragment is methylated at
G966 and C967 in the wild-type strain
(Guymon et al. 2006), tandem mass
spectrometry was again performed (Sup-
plemental Fig. 1B), showing that only
the methyl group on C967 was absent. In
agreement with the suggested nomen-
clature for rRNA modifying enzymes
(Andersen and Douthwaite 2006), we
hereafter refer to TTHA0851 as rsmB
due to substrate specificity identical to
the originally identified enzyme from
E. coli (Gu et al. 1999).
Effect of temperature on growth
of the rsmF null mutant
One possible explanation for methyla-
tion of multiple sites by RsmF is that
such additional methyl groups improve
ribosomal function at elevated temper-
ature. To address this possibility, we ex-
amined the effect of temperature on
growth of the rsmF null mutant. Wild-
type T. thermophilus and the rsmF null
mutant were cocultured at three tem-
peratures and these cocultures were se-
rially subcultured for seven cycles of
24 h each. The proportion of wild-type
and rsmF null mutant cells in each mixed
culture was determined by spreading di-
lutions onto TEM plates with or without
kanamycin. After seven cycles, no differ-
ence in the relative proportions of the
wild-type and rsmF mutant was exhib-
ited at 70°C. However, at 60°C, the rsmF
null mutant constituted only around 5%
FIGURE 2. (Legend on next page)
Demirci et al.
1586
RNA, Vol. 16, No. 8
of the population, and at 80°C the rsmF null mutant was
unable to grow at all. Thus, methylation by RsmF appears to
facilitate growth at temperatures outside the optimal growth
temperature.
RsmF substrate preference
The T. thermophilus rsmB and rsmF genes were each cloned
into an E. coli expression plasmid in order to produce
proteins for X-ray crystallography and in vitro methylation
studies. The expression constructs were equipped with
C-terminal histidine6 tags to facilitate protein purification.
While we achieved a high expression level of RsmF, we were
unable to do so with RsmB despite a series of optimization
attempts. Consequently, in vitro substrate and structure
analyses were performed exclusively with RsmF.
E. coli RsmF requires the 30S ribosomal subunit as a
substrate when the activity is assayed in vitro (Andersen and
Douthwaite 2006). We assayed 70S ribosomes, 30S ribosomal
subunits, and 16S rRNA for the ability to serve as substrates
for methylation by RsmF in vitro. 16S rRNA subfragments of
z50 nt around the target sites were purified after the in vitro
assay and analyzed by mass spectrometry as described above.
In vitro methylation at 70°C showed an interesting but rather
complex substrate pattern. RsmF completely methylates
C1400 when either 16S rRNA or 30S subunits are used as
a substrate. It methylates C1404 to z35% with 16S rRNA
and completely with 30S subunits, and it produces only trace
amounts of methylation of C1407 with 16S rRNA and z75%
with 30S subunits (Fig. 3). There were no indications of
the 70S ribosome being a substrate in vitro. Curiously,
T. thermophilus RsmF expressed in an E. coli rsmF null
mutant almost completely methylated, in vivo, positions
C1400 and C1404, but not C1407 (data not shown).
X-ray crystal structures of RsmF
We determined the structure of T. thermophilus RsmF (456
amino acids) in three different crystal forms and in a com-
plex with cofactor AdoMet to up to 1.3 A˚ resolution (Figs.
4, 5). The structure was solved in space group P43 (data set
RsmF1, 1.4 A˚ resolution) by molecular replacement using
a search model generated with the program Modeller
(Eswar et al. 2008) from the catalytic domain of the RsmF
homolog YebU from E. coli (Pdb 2FRX) (Hallberg et al.
2006). The structures of the AdoMet-bound form in space
group P2 (RsmF2, 1.82 A˚ resolution), of the AdoMet-
bound form (RsmF3, 1.3 A˚ resolution), and of the apo-
form (RsmF4, 1.68 A˚ resolution) in space group P21212
were subsequently solved by molecular replacement with
the refined RsmF1 model. There are two molecules in the
asymmetric unit in space groups P43 and P2 and one mol-
ecule in space group P21212. Electron density is generally
well defined in all crystal forms. The majority of residues
(92.0%, 92.3%, 93.1%, and 92.6%) are in the most favored
region of the Ramachandran plot for RsmF1, RsmF2,
RsmF3, and RsmF4, respectively, and there are no residues
in the disallowed region. The final models consist of
residues 5–178, 194–198, and 201–456 and five additional
residues from the histidine6 affinity tag in both chains of
data set RsmF1; residues 2–456 and five affinity-tag
residues in both chains of data set RsmF2; residues 1–456
and six affinity-tag residues in data set RsmF3; and resi-
dues 1–456 and seven affinity-tag residues in RsmF4. The
N-terminal a-amino group was ordered in data sets RsmF3
and RsmF4 and contained additional electron density,
which we interpreted as N-(dihydroxymethyl)-L-methio-
nine, the hydrated form of N-formyl-methionine. Data
collection and refinement statistics are given in Table 1.
The overall structure of RsmF consists of a central
canonical class I methyltransferase catalytic domain with
additional N-terminal and C-terminal
domains (Figs. 4, 5). The catalytic do-
main is formed by a central seven-
stranded b-sheet that is flanked on both
sides by three helices of varying lengths.
An inserted region between strand b7
and helix a11 contains additional heli-
ces a9 and a10, which interact with the
two N-terminal helices a1 and a2. A
second inserted region following strand
b9 includes the short helices a13 and
FIGURE 2. (A) MALDI mass spectra of an RNase T1-digested 16S rRNA subfragment (pos.
1378–1432) from wild-type cells (upper panel) or from the TTHA1387 (putative rsmF) null
mutant (lower panel). Expected digestion products are labeled; fragments affected by the null
mutation are set in italics. (B) Expansion of the signals affected by the TTHA1387 null mu-
tation. The sequence and methylation status of the RNase T1 products are indicated. (C)
MALDI tandem mass spectrometry of the methylated RNase T1 fragment of 16S rRNA (pos.
1404–1407); wild-type cells (upper panel), TTHA1387 null mutant (lower panel). Mass
spectrometric fragments used to deduce the methylation status are labeled. The position of
the backbone fragments (nomenclature according to McLuckey et al. [1992]) in the sequence is
shown. MH+, precursor ion selected for fragmentation; C, cytosine; mC, methylated cytosine;
C>p, cytidine-2´-39-monophosphate; me, methyl group.
FIGURE 3. In vitro methylation with 30S ribosomal subunits or 16S
rRNA from the T. thermophilus rsmF null mutant as a substrate. Effect
on C1400-, C1404-, and C1407-harboring RNase T1 products. In
vitro methylated products are set in italics. *, artifact signal arising
from the enzyme preparation.
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1587
a14, which interact with helix a12. Furthermore, RsmF
contains three additional smaller domains, an N-terminal
domain consisting of a three-stranded b-sheet and two flank-
ing helices (Fig. 5B, colored in blue), and two C-terminal
domains consisting of four-stranded b-sheets and two or one
helix (Fig. 5B, colored in magenta and red).
Cofactor binding, substrate docking,
and conformational flexibility in the active site
The coordination of AdoMet in the T. thermophilus RsmF
active site is similar to that seen in other class I methyl-
transferases. However, the previously published structure of
E. coli RsmF did not contain the cofactor AdoMet in the
active site, precluding a direct comparison. Both the T.
thermophilus and E. coli RsmF cofactor-binding sites reveal
a new variation for the methyltransferase signature motif I
(Malone et al. 1995), with the highly conserved GxGxG
sequence replaced by 109AAAPG113. The combination of
three alanines and a proline results in a loop conformation
that is very similar to that observed in other methyltrans-
ferases with a GxGxG motif (e.g., RsmC) (Demirci et al.
2008a). In RsmF, the amide hydrogen atom of the last
glycine residue forms a hydrogen bond with the cofactor
carboxy group (Fig. 6). Other key interactions with AdoMet
are well conserved in RsmF. The cofactor adenine ring is
located in a mainly hydrophobic pocket lined by residues
Val134, Pro160, and Leu211. This pocket is open toward the
solvent. The adenine amino group is not specifically recog-
nized and interacts with solvent water molecules. The ribose
hydroxyl groups form hydrogen bonds with Glu133 and
Arg138, and the methionine amino group interacts with
Asp177. The AdoMet cofactor is bound in a cleft in the
RsmF active site, which suggests that substrate cytidine bases
are inserted into the active site in an unstacked conforma-
tion. Inspection of the electrostatic charge distribution re-
veals a large positively charged surface region, which would
be consistent with binding to an RNA surface and modifi-
cation of the substrate base in an unstacked orientation (Fig.
6C). To evaluate the possible orientation of a substrate base
in the active site, we performed computational docking
calculations with the program Dock6 (Lang et al. 2009). The
resulting positions of cytosine and m5C in the presence of
AdoMet in data set RsmF3 are highly similar to each other,
with m5C placed into the active site with its phosphate group
toward a positively charged pocket at the entrance of the
active site cleft (Fig. 6E). The position of the phosphate
group is close to a sulfate molecule that we observed in data
set RsmF1, providing further support for the results of the
docking calculation (Fig. 6F).
Interestingly, we observed that three active site segments
were disordered in data set RsmF1. These segments include
residues 179–193 (including helices a9 and a10 and the
catalytic Cys180), residues 199–200, and the N-terminal
residues 1–5, which interact with the first two segments (Fig.
6E,F, colored in green). We observed electron density for the
intervening residues 194–198, which formed a lattice con-
tact with a neighboring molecule. However, the position of
FIGURE 4. Structure-based sequence alignment of RsmF from T. thermophilus and E. coli. Secondary structure elements of T. thermophilus RsmF
are indicated on top. The color scheme for the secondary structure elements is as in Figure 5A. The position of the variant methyltransferase motif
I is marked with a red box. A flexible region observed in the active site is marked with a green box; residues interacting with the cofactor are
marked with orange boxes.
Demirci et al.
1588
RNA, Vol. 16, No. 8
these five residues was not related to their position in the
other three data sets, suggesting that the extended active
site region between residues 179 and 201 can reorient in the
RsmF structure. This observation suggests that this active
site region is dynamic, which may be important for sub-
strate binding at 72°C, the optimum growth temperature of
T. thermophilus.
DISCUSSION
Substrate recognition mechanisms
We have identified the two enzymes responsible for the
synthesis of the four m5C modifications of T. thermophilus
16S rRNA, and characterized the RsmF methyltransferase
responsible for synthesizing three of these. rRNA modifying
enzymes in bacteria are generally highly specific, with a
one-to-one association between the modifying enzyme and
the modification. A few cases of multitarget ribosome mod-
ifying enzymes have been reported (Helser et al. 1972;
Demirci et al. 2008b), but to our knowledge T. thermophi-
lus RsmF is the first rRNA methyltransferase found to mod-
ify three different nucleotides. Most ribosome modifying
enzymes probably recognize assembly intermediates, and
the data presented here are consistent with that notion.
T. thermophilus RsmF methylates C1400 and C1404 in
vitro using either 16S rRNA or 30S subunits as substrates,
whereas both E. coli (Andersen and Douthwaite 2006) and
T. thermophilus RsmF exclusively utilize 30S subunits as
substrates for methylation of C1407. This may reflect that
C1400 and C1404 methylations do not rely on the asso-
ciation of ribosomal proteins in order to be recognized by
T. thermophilus RsmF. C1407 methylation, in contrast,
depends on both rRNA and the ribosomal protein for the
recognition by RsmF in both T. thermophilus and E. coli.
More puzzling is the observation that T. thermophilus RsmF
does not methylate E. coli ribosomes in vivo on C1407. It
is perhaps worth noting that methylation of C1407 in the
T. thermophilus 30S ribosomal subunit in vitro was less
efficient than methylation of the other two positions,
indicating the need for a particular intermediate assembly
structure or for accessory factors. The only clear in vitro
FIGURE 5. Overall structure of RsmF. (A) Schematic representation of the position of the substrate bases in the 30S ribosomal subunit (Pdb
entry 2WRI) (Gao et al. 2009). C1400, C1404, and C1407 in helix 44 (green) are shown in stick representation with 5-methyl groups as pink
spheres. (B) Schematic stereo representation of the overall structure of RsmF. Secondary structure elements are in orange and yellow for the
catalytic domain, in salmon and blue for additional N-terminal domains, and in magenta and red for C-terminal domains. The flexible region in
the active site including helices a9 and a10 is indicated with an arrow and colored in green. (C) Topology diagram with secondary structure
elements colored as in B.
T. thermophilus 16S rRNA methyltransferase RsmF
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1589
methylation observed at 37°C with T. thermophilus RsmF
was on C1400 with 16S rRNA as the substrate (data not
shown), which is not evidently related to the methylation
pattern in vivo in the heterologous system. It seems unlikely
that the aberrant methylation in the heterologous system
reflects species-specific differences in the mature 30S ribo-
somal subunit, given the extreme sequence and structural
conservation of the decoding site. Instead, it may reflect
differences in 30S subunit assembly in the two organisms,
necessary due to the large difference in growth temperature.
The four m5C residues in T. thermophilus 16S rRNA are
clustered in and around the functionally critical decoding
center at or close to sites of contact with tRNA, mRNA, and
EF-G (Ogle et al. 2002; Selmer et al. 2006; Gao et al. 2009).
m5C1400, m5C1404, and m5C1407 are located in the
subunit body, while m5C967 is located in the subunit head,
about 10 A˚ from m5C1400 (Wimberly et al. 2000). Exam-
ination of the 30S subunit crystal structure (Wimberly et al.
2000) indicates that the three bases methylated by RsmF are
situated in three distinct structural contexts, but provides
few clues to a common mode of substrate recognition.
C1400 is an unpaired base protruding from a sharply bent
segment of rRNA at the junction of helices 43 and 44, while
C1404 and C1407 are engaged in Watson–Crick pairs within
helix 44 (Fig. 4A). The C5 positions of the latter two bases
are not obviously accessible, such that RsmF would need to
approach them from the major groove side. While a flipping
of C1407 out of the helix via the minor groove could allow
access to this base, such a mechanism would be problematic
for C1404, whose minor groove side is packed against the
rest of the 30S subunit.
While m5C1404 and m5C1407 are about 11 A˚ apart
(Selmer et al. 2006), m5C1400 is quite distant from both of
these bases (about 21 and 30 A˚ , respectively). The pro-
truded conformation of m5C1400 in the mature 30S sub-
unit is due in part to base-pairing interactions between
adjacent bases and the 1500 region of 16S rRNA (C1399–
G1504 and G1401–C1501). As the 1500 region is one of the
last segments of 16S rRNA to be synthesized, C1400 could
potentially be positioned much closer to C1404 and C1407
in an assembly intermediate, prior to the formation of the
C1399–G1504 and G1401–C1501 base pairs. RsmF could
utilize a single binding mode to then access all three bases,
further facilitated by its flexible active site domain. Meth-
ylation of C1400 in mature 30S subunits would therefore
involve disruption of the adjacent base pairs. A complete
understanding of the recognition mechanism of these
enzymes will require high-resolution structural data on
assembly intermediate-enzyme complexes. Given the large
number of potential subunit assembly intermediates, pre-
cisely defining the physiological substrate for RsmB and
RsmF will be a formidable task.
TABLE 1. Data collection and refinement statistics
RsmF1
RsmF2
RsmF3
RsmF4
Data collectiona
AdoMet
AdoMet
Space group
P43
P2
P21212
P21212
Cell dimensions
a, b, c (A˚ )
71.0, 71.0, 186.7
66.0, 78.3, 108.1
89.7, 109.0, 51.0
89.8, 109.1, 50.8
a, b, g (°)
90, 90, 90
90, 107.1, 90
90, 90, 90
90, 90, 90
Resolution (A˚ )b
30–1.4 (1.55–1.40)
30–1.82 (1.89–1.82)
30–1.30 (1.34–1.30)
30–1.68 (1.74–1.68)
Rmerge
0.065 (0.59)
0.08(0.38)
0.058(0.36)
0.15 (0.49)
I/sI
29.3(2.15)
12.6 (2.04)
24.3 (2.05)
14.2 (1.73)
Completeness (%)
90.1 (72.6)
97.0(86.5)
95.6(66.3)
99.6 (97.8)
Redundancy
8.9 (5.4)
2.8(2.0)
5.2(2.1)
6.4 (3.9)
Refinement
Resolution (A˚ )
30–1.4 (1.42–1.40)
30–1.82 (1.84–1.82)
30–1.30 (1.32–1.30)
30–1.68 (1.69–1.68)
Number of reflections
161,955 (4356)
91,762 (2583)
119,490/2640
109,375 (3244)
Rwork/Rfree
0.169/0.189 (0.227/0.263)
0.162/0.194 (0.222/0.259)
0.177/0.191 (0.233/0.234)
0.173/0.192 (0.217/0.266)
Number of atoms
Protein
6766
7117
3598
3574
Ligand/ion
20
54
27
1
Water
1486
1285
856
665
B-factors
Protein
24.1
20.4
15.9
17.2
Ligand/ion
34.5
21.6
17.5
19.1
Water
38.7
36.3
33.6
33.6
RMSDs
Bond lengths (A˚ )
0.009
0.006
0.005
0.004
Bond angles (°)
1.22
1.04
1.14
0.94
aOne crystal used for each data set.
bThe highest resolution shell is shown in parentheses.
Demirci et al.
1590
RNA, Vol. 16, No. 8
Structural comparison of RsmF with related
methyltransferases
A database search with Dali (Holm et al. 2008) confirmed
the structural similarity of the T. thermophilus and E. coli
(PDB 2FRX) (Hallberg et al. 2006) RsmF homologs, which
superimpose with a root-mean-square deviation (RMSD)
of 1.6 A˚ for 342 Ca atoms (Fig. 7A) and
are the only two structures in the Pro-
tein Database with this domain organi-
zation. Even so, substantial structural
differences are observed in most of the
loop regions and for the long connect-
ing loop between the methyltransferase
domain and the first C-terminal domain.
While in the active site, the positions
of residues in the cofactor-binding site
and of the two cysteine residues are
conserved (Fig. 7B), there are a number
of positively charged residues (Arg30,
Arg190, Arg194, His195, and Arg203) in
the T. thermophilus structure that are
absent from the E. coli enzyme (Fig. 7B).
Three of these are located in the flexible
region, and the combination of a posi-
tive charge and flexibility close to the
active site is suggestive of a functional
contribution of this region to the mul-
tisite specificity of T. thermophilus RsmF
(Figs. 6C,D, 7B). Methylation of three
rRNA positions may require an increase
in the enzyme’s structural dynamics in
order to accommodate the 30S subunit
in slightly different orientations. Similar
observations have been made for other
multi-site-specific methyltransferases in-
cluding KsgA, which modifies two adja-
cent adenosines in the 30S ribosomal
subunit (O’Farrell et al. 2004; Demirci
et al. 2009), and the PrmA ribosomal
protein methyltransferase, which under-
goes dramatic interdomain movements
to modify multiple lysine residues and the
N-terminal a-amino group on the same
substrate protein (Demirci et al. 2007,
2008b).
The second C-terminal domain in
RsmF is related to the RNA-binding
PUA
(pseudouridine
synthase
and
archaeosine transglycosylase) domains
(Perez-Arellano et al. 2007). The RlmI
methyltransferase,
which
produces
m5C1962 in 23S rRNA (Purta et al.
2008) also contains a PUA domain
(Sunita et al. 2008), although it is
N-terminal to the catalytic methyltransferase domain and
in a different orientation. PUA domains contain six
b-strands, which form a central pseudobarrel closed by
a short 310-helix. A comparison of the C-terminal domain
in RsmF with a typical PUA domain in archaeosine trans-
glycosylase (ArcTGT, Pdb entry 1J2B) reveals that the
central fold is similar (53 Ca atoms align with an RMSD
FIGURE 6. Substrate docking and conformational flexibility. (A) Cofactor-binding site in
RsmF. Bound AdoMet is shown in blue sticks. Hydrogen bonds to coordinating residues are
indicated. (B) Final sA-weighted 2mFO-DFC electron density map of the cofactor-binding site
(data set RsmF2) contoured at the 1s level. (C,D) Comparison of the electrostatic surface
charge distribution between RsmF from T. thermophilus and E. coli. The location of the
C-terminal domains and of the flexible region (labeled ‘‘FLEX’’) is indicated with circles.
AdoMet and docked m5C are shown as sticks. AdoMet from the T. thermophilus structure is
shown with the E. coli structure for comparison. (E) The modeled position of m5C docked into
the active site of RsmF. m5C and AdoMet are shown in tan and blue sticks, respectively.
Residues in the flexible region are shown in green. (F) The active site region in data set RsmF1.
A sulfate ion is observed close to the position of the m5C phosphate group. Residues 194–198 in
the flexible region interacting with a neighboring RsmF molecule are indicated with an arrow.
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1591
of 1.8 A˚ ), but that several connecting loop regions are
substantially shorter and two a-helices and one b-strand of
the pseudobarrel are absent. Thus, this PUA-like domain
differs considerably from typical PUA domains. However,
the similarity to RNA-binding PUA domains and the
positive surface charge distribution observed in both RsmF
structures are suggestive for a conserved function of the
PUA-like domain in RNA recognition.
The next most closely related structure is E. coli RsmB
(474 residues, PDB 1SQF) (Foster et al. 2003). A total of
264 Ca atoms can be aligned with an RMSD of 1.5 A˚
between RsmB and RsmF. Both enzymes retain the same
organization for the N-terminal domains and the core
methyltransferase domain. However, an additional 140-
residue N-terminal RNA-binding domain provides sub-
strate specificity to RsmB, whereas the two C-terminal
domains following the core methyltransferase domain (160
residues) are likely to determine substrate recognition by
RsmF. Thus, these two enzymes have evolved substrate
specificity via acquisition of additional,
unrelated RNA recognition domains.
While there are as yet no enzyme–
substrate complexes for rRNA m5C-
methyltransferases, insights into the RsmF
catalytic mechanism can be gleaned from
a comparison with the RlmD and TrmA
m5U methyltransferases in covalent in-
termediate complexes with RNA oligonu-
cleotides (Lee et al. 2005; Alian et al.2008).
DNA and RNA m5C methyltransferases
use a thiol from a catalytic cysteine residue
to attack the six-position of the pyrimi-
dine base to activate the five-position for
methyl group transfer (Liu and Santi
2000). Cys180 and Cys230 in RsmF are
positioned equivalently to the catalytic
Cys324 and the catalytic base Glu358 of
TrmA. The substrate nucleotides insert
into TrmA and RsmF in unrelated di-
rections, consistent with a lack of struc-
tural homology outside the methyltrans-
ferase domains. Nevertheless, the C5
positions of the pyrimidine rings and of
the 5-methyl carbons are quite similar
with respect to the catalytic cysteine resi-
dues and the AdoMet cofactor. The same
structural homology of the active site
geometry can be observed in comparison
with the RlmD methyltransferase (Sup-
plemental Fig. 2; Lee et al. 2005).
A possible origin of T. thermophilus
RsmF
T. thermophilus RsmF shows the highest
similarities with proteins from close relatives, namely,
Thermus aquaticus and two Meiothermus species of the
Thermaceae family. Remarkably, the next highest similarities
(BLAST scores between 267 and 359) are with the NOL1/
NOP2/Sun proteins from the Gram-positive Firmicutes
phylum (Supplemental Fig. 3). This similarity, together
with the fact that the Thermaceae family and most members
of the Firmicutes identified in Supplemental Figure 3 are
thermophilic, suggest that rsmF has undergone horizontal
transfer between the Thermaceae family and members of the
Firmicutes phylum. Thus, we speculate that this version of
the RsmF protein, which catalyzes methylation of three
cytidines, may be adaptive for existence in thermally
challenging environments. The effect of the loss of methyl-
ation by RsmF on growth at different temperatures is
consistent with this notion. Our hypothesis of horizontal
transfer of rsmF predicts that RsmF of other members of the
Thermaceae family and members of the Firmicutes phylum
will also be found to introduce multiple m5C modifications.
FIGURE 7. Comparison with other methyltransferases. (A) Differences between the overall
structures of RsmF from T. thermophilus (orange) and E. coli (green). AdoMet bound in the T.
thermophilus structure is shown as blue sticks. (B) Comparison of the active site region in both
enzymes. Residues in the cofactor-binding site and in the flexible region in the T. thermophilus
enzyme are shown as sticks. (C) Comparison between RsmF (orange) and a substrate complex
structure of the TrmA methyltransferase (cyan/blue). (D) Comparison of the active site region
illustrating the differences in the insertion direction of the substrate base in RsmF (m5C in
light orange) and in TrmA (m5U in cyan).
Demirci et al.
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RNA, Vol. 16, No. 8
MATERIALS AND METHODS
Cloning of the T. thermophilus rsmB and rsmF genes
The T. thermophilus HB8 loci TTHA0851 (GenBank accession
number BAD70674) and TTHA1387 (GenBank accession number
BAD71210) were PCR amplified from genomic DNA and purified
via the High Pure PCR Template Preparation Kit (Roche). The
100 mL PCRs contained 150 ng DNA, 10 mM of each primer,
10 mM dNTP, 1 unit Phusion DNA polymerase (Finnzymes), and
1x Phusion HF buffer. Primers for rsmB amplification were 59-CC
CTGGACATATGAGGGCCGG-39 and 59-GGCCAAGATCTTGCC
TGAGAG-39, and the temperature cycling was as follows: 98°C/30
sec; 30X (98°C/10 sec, 59°C/20 sec, and 72°C/36 sec); and 72°C/420s.
Primers for rsmF amplification were 59-GCTAGGGTACACATA
TGCTGCCC-39 and 59-GCACGGGGGTGAGATCTAAGCCC-39,
and the temperature cycling was as follows: 98°C/30 sec; 30X
(98°C/10 sec, 62°C /20 sec, 72°C/42 sec); and 72°C/420 sec. The
desired PCR products were purified from agarose gels using the
GFX PCR purification kit (GE Healthcare). The PCR fragments
were digested with NdeI and BglII and inserted into the expression
vector pLJ102 (Andersen and Douthwaite 2006), generating
isopropyl-1-thio-b-D-galactopyranoside (IPTG)-inducible genes
for the recombinant proteins with a C-terminal histidine6 tag.
The constructs (designated pLJ102-RsmB and pLJ102-RsmF) were
used to transform an rsmF-deletion derivative of E. coli CP79
(Andersen and Douthwaite 2006).
Deletion of the T. thermophilus rsmB and rsmF genes
Constructs for inactivation of the T. thermophilus rsmB and rsmF
genes were made by inserting the gene for a heat tolerant
kanamycin (HTK) nucleotidyltransferase (Hoseki et al. 1999) into
the methyltransferase parts of either pLJ102-RsmB or pLJ102-
RsmF. The htk part of pUC18-htk (Hashimoto et al. 2001) was
amplified by PCR with primers that introduced an upstream AvrII
site and a downstream SacI site into the product for later disrup-
tion of rsmB. For rsmF disruption, the PCR primers introduced
SacI restriction enzyme sites both upstream of and downstream
from the htk gene. These sites were used to insert the PCR prod-
ucts into pLJ102-RsmB and pLJ102-RsmF to form the plasmids
pLJ102-RsmBThtk and pLJ102-RsmFThtk, which were propa-
gated in the E. coli strain Top10 (Invitrogen). T. thermophilus
HB8 was transformed with pLJ102-RsmBThtk or pLJ102-
RsmFThtk selecting for kanamycin resistance as described by
others (Hashimoto et al. 2001; Cameron et al. 2004). Kanamycin-
resistant transformants were restreaked twice. Gene disruptions
were verified by PCR with primers distal to the interrupted rsmB
or rsmF genes on genomic DNA; resulting PCR products were
characterized by sequencing.
Growth competition assays
Wild-type and rsmF null mutant liquid cultures were grown at
70°C to saturation, then equal numbers of cells from each were
mixed and incubated in 5 mL TEM medium at 60°C, 70°C, or
80°C. After growth for 24 h, 100 mL of the 60°C culture, 10 mL of
the 70°C culture, and 1000 mL of the 80°C culture were trans-
ferred to a fresh 5-mL medium and incubated at the respective
temperatures for another 24 h. This was repeated in independent
triplicates for seven cycles. Samples of 1 mL were collected at each
dilution and half was plated on TEM plates without antibiotic and
the other half was plated on TEM plates with 30 mg/mL
kanamycin. The plates were incubated at 70°C.
Purification of T. thermophilus ribosomal subunits
and ribosomes
T. thermophilus culture (1 L) was grown in TEM media (contain-
ing 30 mg/mL of kanamycin when appropriate) with shaking at
70°C to an OD600 = 0.6. Cells were harvested and washed once
with 100 mL of buffer A (10 mM NH4Cl, 20 mM MgCl2, 100 mM
KCl, and 10 mM Tris-HCl [pH 7.5]), then suspended in 10 mL of
buffer A, and disrupted by sonication. The lysate was cleared by
centrifugation twice in a Beckman JA20 rotor at 16,000 rpm for
10 min at 4°C. Crude ribosomes were collected by centrifugation
in a Beckman Ti50 rotor at 19,000 rpm for 19 h at 4°C, and
dissolved in buffer A. 70S ribosomes were obtained by centrifu-
gation of 100 A260 units of crude ribosomes through a 10%–40%
sucrose gradient (200 mM NH4Cl, 20 mM MgCl2, 20 mM Tris-
HCl [pH 7.5]) in a Beckman SW28 rotor at 20,000 rpm for 18 h at
4°C. Fractions containing intact 70S ribosomes were pooled and
concentrated by centrifugation in a Beckman Ti50 rotor at 40,000
rpm for 22 h at 4°C, washed, and dissolved in buffer A, and stored
at 80°C.
50S and 30S ribosomal subunits were obtained by adjusting
100 A260 units of crude ribosomes (10 mM NH4Cl, 2 mM MgCl2,
100 mM KCl, and 10 mM Tris-HCl [pH 7.5]) and passing
through a 5%–20% sucrose gradient (200 mM NH4Cl, 1 mM
MgCl2 and 20 mM Tris-HCl [pH 7.5]) in a Sorvall AH-629 rotor
at 20,000 rpm for 18 h at 4°C. After pooling of the relevant
fractions, the subunits were adjusted to 10 mM MgCl2 and
pelleted by centrifugation in a Beckman Ti50 rotor at 40,000
rpm for 22 h at 4°C, washed with and dissolved in 10 mM NH4Cl,
10 mM MgCl2, 100 mM KCl, and 10 mM Tris-HCl (pH 7.5), and
stored at 80°C.
Isolation of 16S rRNA and subfragments
from T. thermophilus and E. coli
Water (400 mL) was added to 100 mL of 30S ribosomal subunits
and the rRNA was extracted with 500 mL phenol, phenol/
chloroform, and chloroform. rRNA was ethanol precipitated
and dissolved in water. Purification of 16S rRNA subfragments
was performed as previously described (Andersen et al. 2004).
Briefly, 16S rRNA was hybridized to an excess of oligodeoxynu-
cleotide complementary to either the region 944–990 or the region
1378–1432. Single-stranded nucleic acids were digested with
Mung Bean Nuclease and RNase A. The resulting mixture was
separated on a polyacrylamide gel. Bands were visualized by ethid-
ium bromide staining, excised, and eluted.
E. coli CP79 with the endogenous rsmF inactivated, but com-
plemented with the T. thermophilus homolog on the plasmid
pLJ102-RsmF, were grown at 37°C to an OD450 = 0.45 in 200 mL
of LB medium containing 100 mg/L of ampicillin. RsmF expres-
sion was induced by addition of IPTG to 1 mM, and incubation
for another 3 h. Cells were harvested by centrifugation at 4°C,
washed in 100 mL TMN buffer (50 mM Tris-HCl [pH 7.8],
10 mM magnesium acetate, 100 mM NH4Cl), and resuspended in
2 mL TMN buffer prior to lysis by sonication (7 3 30 sec on ice)
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1593
and removal of debris by centrifugation (10 min/14,000 rpm/4°C/
microcentrifuge). Total RNA was recovered from the supernatant
by phenol extraction and ethanol precipitation. A 16S rRNA
subfragment was isolated as described above using an oligodeoxy-
nucleotide complementary to the region 1378–1432.
In vitro methylation
Reactions contained 50 pmol of 16S rRNA, 30S subunits, or 70S
ribosomes from the T. thermophilus TTHA1387 null mutant as the
substrate in a total volume of 100 mL (containing 100 mM NH4Cl,
10 mM MgCl2, 40 mM Hepes [pH 7.5]), 6 mM b-mercaptoetha-
nol, and 10% glycerol (prepared as a two times concentrated stock
solution), 1.5 mM S-adenosyl methionine, and 2 mg of recombi-
nantly expressed RsmF (see below).
For the reaction at 70°C, water and stock buffer were mixed and
left at room temperature for 15 min. Then a substrate, an enzyme
and S-adenosyl methionine were added and incubated at 70°C for
1 h. The 37°C reaction was started by mixing water and buffer
followed by 15 min at room temperature; the substrate was added
and the mixture transferred to 50°C for 5 min. After cooling to
37°C, S-adenosyl methionine and an enzyme were added and the
incubation continued for 1 h. Reactions were stopped by phenol/
chloroform extraction and the rRNA was recovered by ethanol
precipitation before purification of 16S rRNA subfragments as
described above. Control reactions without enzyme or S-adenosyl
methionine were carried out in all instances.
RNase T1 digestion and mass spectrometry
A purified 16S rRNA subfragment (1–2 pmol) was incubated with
2 units RNase T1 (Roche) and 50 mM 3-hydroxypicollinic acid
(3-HPA) in a total volume of 2 mL for 4 h at 37°C. MALDI mass
spectrometry was performed either on an ABI voyager STR in-
strument or a Waters Q-TOF MALDI instrument; MALDI tan-
dem mass spectrometry was done on a Waters Q-TOF MALDI
instrument. All spectra were recorded in positive ion mode using
3-HPA as the matrix. Experimental details were as previously de-
scribed (Douthwaite and Kirpekar 2007).
Protein expression and purification for crystallization
E. coli BL21 (DE3) (Invitrogen) containing pLJ102-RsmF was
grown to midlog phase in LB media at 37°C in the presence of
200 mg/mL ampicillin. Protein expression was induced at 20°C
with 400 mM IPTG. Cells were pelleted after 18 h by centrifuga-
tion at 4000 rpm for 20 min at 4°C and lysed by ultrasonication
on ice in a buffer containing 20 mM Tris-HCl (pH 8.5), 300 mM
NaCl, 5 mM b-mercaptoethanol, 0.1% Triton X-100, and 5%
glycerol. Cell debris and membranes were pelleted by centrifuga-
tion at 11,000 rpm for 30 min at 4°C. The soluble E. coli proteins
were precipitated by heat treatment at 65°C for 30 min and
pelleted by centrifugation at 11,000 rpm at 4°C for 30 min.
Soluble C-terminally hexahistidine-tagged T. thermophilus RsmF
was further purified by affinity chromatography with nickel-
nitrilotriacetic acid resin (Qiagen). Untagged proteins were re-
moved with buffer containing 20 mM Tris-HCl (pH 8.5), 250 mM
NaCl, and 1 mM imidazole (pH 8.5). Recombinant RsmF was
then eluted with the same buffer containing 150 mM imidazole.
The protein was then purified by cation exchange chromatogra-
phy (SP) (GE Healthcare) at pH 8.5, using a linear gradient of
10 mM to 1 M NaCl concentration. RsmF fractions were pooled
and concentrated and applied to a size-exclusion S200 column (GE
Healthcare) equilibrated with a buffer containing 20 mM Tris-HCl
(pH 8.5) and 200 mM NaCl. Purified RsmF was concentrated to
13 mg/mL for crystallization trials. The C-terminal hexahistidine
tag was not removed for crystallization. For the production of
selenomethionyl proteins, the expression construct was trans-
formed into B834 (DE3) cells (Novagen). The bacterial growth
was carried out in defined LeMaster medium (Hendrickson et al.
1990), and the protein was purified using the same protocol as for
the unmodified protein. To form the RsmF-AdoMet complex,
purified RsmF was mixed with 4 mM AdoMet incubated at 60°C
for 15 min and slowly cooled to room temperature before
performing crystallization experiments.
Crystallization of RsmF
All crystals were obtained using the microbatch technique under
oil at 4°C. To obtain the RsmF1 crystal form, 1 mL of protein
solution was mixed with the reservoir solution containing 20%
(w/v) PEG3350 and 200 mM sodium sulfate decahydrate (pH
6.6). Initial crystals grew over the course of 1–2 wk with maxi-
mum dimensions of 0.3 3 0.3 3 0.2 mm. To obtain the RsmF2
crystal form, 1 mL of the RsmF–AdoMet complex was mixed with
the reservoir solution containing 200 mM NaCl, 12% w/v
PEG8000 and 100 mM HEPES-KOH (pH7.5). Initial crystals
grew over the course of 2–3 wk with maximum dimensions of
0.1 3 0.4 3 0.4 mm. To obtain the RsmF3 crystal form, 1 mL of
the RsmF–AdoMet complex was mixed with the reservoir solution
containing 10% w/v PEG1000, 200 mM NaCl, and 100 mM Tris-
HCl (pH 8.5). The initial crystals grew over the course of 1–2 wk
with maximum dimensions of 0.05 3 0.3 3 0.4 mm. To obtain
the RsmF4 crystal form, 1 mL of the RsmF–AdoMet complex
solution was mixed with a reservoir solution containing 160 mM
magnesium chloride hexahydrate, 80 mM Tris-HCl (pH 8.5), and
24% w/v PEG4000. Initial crystals grew over the course of 1–2 wk
with maximum dimensions of 0.05 3 0.3 3 0.3 mm. RsmF1
crystals were gradually dehydrated by increasing the PEG3350 to
30% w/v and then cryoprotected in a mother liquor supplemented
with 25% v/v glycerol and then flash-frozen by being plunged into
liquid nitrogen. RsmF2 crystals were cryoprotected in a mother
liquor supplemented with 20% v/v ethylene glycol and then flash-
frozen by being plunged into liquid nitrogen. RsmF3 crystals
were cryoprotected by gradually increasing the concentration of
PEG1000 to 30% and then flash-frozen by being plunged into
liquid nitrogen. RsmF4 crystals were cryoprotected in a mother
liquor supplemented with 20% glycerol and then flash-frozen by
being plunged into liquid nitrogen.
Data collection
X-ray diffraction data for RsmF1, RsmF2, and RsmF4 crystals
were collected on a MAR CCD detector at the X4C beamline of
the National Synchrotron Light Source in Brookhaven at a wave-
length of 0.979 A˚ and 180°C. Diffraction data for RsmF3 crystals
were collected on an ADSC CCD detector at the X4A beamline of
the National Synchrotron Light Source in Brookhaven at a wave-
length of 0.979 A˚ and 180°C. Diffraction data for RsmF1 in
space group P43 were collected to 1.4 A˚ resolution with cell
Demirci et al.
1594
RNA, Vol. 16, No. 8
dimensions a = 71.0 A˚ , b = 71.0 A˚ , and c = 186.7 A˚ . Diffraction
data to 1.82 A˚ for RsmF2 were collected in space group P2 with
cell dimensions a = 66.0 A˚ , b = 78.3 A˚ , and c = 108.1 A˚ .
Diffraction data to 1.29 A˚ for RsmF3 were collected in space group
P21212 with cell dimensions a = 89.7 A˚ , b = 109.0 A˚ , and c = 51.0
A˚ . Diffraction data to 1.68 A˚ for RsmF4 were collected in space
group P21212 with cell dimensions a = 89.8 A˚ , b = 109.1 A˚ , and c =
50.8 A˚ . A single crystal was used for each data set. The diffraction
images were processed and scaled with the HKL2000 package
(Otwinowski and Minor 1997). The data processing statistics are
summarized in Table 1.
Structure determination and refinement
The RsmF structure was solved by molecular replacement with the
program Phaser (McCoy et al. 2007) from the CCP4 program
suite (Bailey 1994) in space group P43 to 1.4 A˚ resolution (data set
RsmF1). The initial search model was built with the program
Modeller (Eswar et al. 2008) from the catalytic domain of E. coli
YebU (Pdb code 2FRX). After the placement of two RsmF
catalytic domains in the asymmetric unit and the initial re-
finement with Refmac (Murshudov et al. 1997), the model was
further rebuilt with ARP/wARP (Langer et al. 2008). The resulting
model was 90% complete and manually checked and completed
with Coot (Emsley and Cowtan 2004). Final crystallographic re-
finement was performed with the program Phenix (Adams et al.
2002). The other crystal forms were subsequently solved by
molecular replacement. The atomic coordinates from the RsmF4
model were then used for initial refinement of the RsmF–AdoMet
complex structure in space group P21212 (RsmF3). There are two
molecules in the asymmetric unit in data sets RsmF1 and RsmF2,
and one molecule in RsmF3 and RsmF4. The crystallographic
R/Rfree factors are 0.17/0.19, 0.16/0.19, 0.18/0.19, and 0.17/0.19
for the four data sets: RsmF1, RsmF2, RsmF3, and RsmF4,
respectively. The stereochemical quality of the model was assessed
with Procheck (Laskowski et al. 1993). The Ramachandran sta-
tistics (most favored/additionally allowed/generously allowed/
disallowed) are 91.9%/8.1%/0.0%/0.0% for RsmF1, 91.9%/8.1%/
0.0%/0.0% for RsmF2, 93.6%/6.4%/0.0%/0.0% for RsmF3, and
92.5%/7.5%/0.0%/0.0% for RsmF4. The refinement statistics are
summarized in Table 1. Figures were generated using Pymol
(DeLano 2002).
Atomic coordinates
Coordinates and structure factors have been deposited in the
Protein Data Bank with accession codes 3M6U, 3M6V, 3M6W,
and 3M6X for data sets RsmF1, RsmF2, RsmF3, and RsmF4,
respectively.
SUPPLEMENTAL MATERIAL
Supplemental material can be found at http://www.rnajournal.org.
ACKNOWLEDGMENTS
We thank John Schwanof and Randy Abramowitz for access to the
X4A and X4C beamlines at the National Synchrotron Light Source.
This work was supported by grants GM19756 and GM19756-37S1
from the National Institutes of Health.
Received January 14, 2010; accepted April 26, 2010.
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|
3M6X
|
Multi-site-specific 16S rRNA methyltransferase RsmF from Thermus thermophilus in space group P21212
|
Multi-site-specific 16S rRNA methyltransferase RsmF
from Thermus thermophilus
HASAN DEMIRCI,1,3 LINE H.G. LARSEN,2,3 TRINE HANSEN,2 ANETTE RASMUSSEN,2 ASHWIN CADAMBI,1
STEVEN T. GREGORY,1 FINN KIRPEKAR,2 and GERWALD JOGL1
1Department of Molecular Biology, Cell Biology and Biochemistry, Brown University, Providence, Rhode Island 02912, USA
2Department of Biochemistry and Molecular Biology, University of Southern Denmark, 5230 Odense M, Denmark
ABSTRACT
Cells devote a significant effort toward the production of multiple modified nucleotides in rRNAs, which fine tune the ribosome
function. Here, we report that two methyltransferases, RsmB and RsmF, are responsible for all four 5-methylcytidine (m5C)
modifications in 16S rRNA of Thermus thermophilus. Like Escherichia coli RsmB, T. thermophilus RsmB produces m5C967. In
contrast to E. coli RsmF, which introduces a single m5C1407 modification, T. thermophilus RsmF modifies three positions,
generating m5C1400 and m5C1404 in addition to m5C1407. These three residues are clustered near the decoding site of the
ribosome, but are situated in distinct structural contexts, suggesting a requirement for flexibility in the RsmF active site that is
absent from the E. coli enzyme. Two of these residues, C1400 and C1404, are sufficiently buried in the mature ribosome
structure so as to require extensive unfolding of the rRNA to be accessible to RsmF. In vitro, T. thermophilus RsmF methylates
C1400, C1404, and C1407 in a 30S subunit substrate, but only C1400 and C1404 when naked 16S rRNA is the substrate. The
multispecificity of T. thermophilus RsmF is potentially explained by three crystal structures of the enzyme in a complex with
cofactor S-adenosyl-methionine at up to 1.3 A˚ resolution. In addition to confirming the overall structural similarity to E. coli
RsmF, these structures also reveal that key segments in the active site are likely to be dynamic in solution, thereby expanding
substrate recognition by T. thermophilus RsmF.
Keywords: rRNA methyltransferase; 5-methylcytidine; RsmB; RsmF; RNA mass spectrometry
INTRODUCTION
Ribosomal RNAs (rRNAs) are post-transcriptionally mod-
ified in all three domains of life, and many modifications
are phylogenetically conserved. Most modifications are
located in functionally important regions of the ribosome,
where they probably act to fine tune protein synthesis (Agris
2004; Gustilo et al. 2008). Complete modification maps of
bacterial 16S rRNAs have been determined for only a hand-
ful of species, and among these are the enteric bacterium
Escherichia coli and the extremely thermophilic bacterium
Thermus thermophilus (Guymon et al. 2006). Despite the
large phylogenetic divergence of these two organisms, their
ribosome modification patterns are quite similar. Of the 11
E. coli and 14 T. thermophilus 16S rRNA modifications,
eight are identical. This suggests a set of common functional
requirements conserved since divergence from their last
common ancestor, and also suggests common recognition
mechanisms among their modifying enzymes.
For most ribosome modifications, a single enzyme recog-
nizes and modifies a single site. However, there exist nota-
ble exceptions. Among these are dimethylation of two adja-
cent adenosines in 16S rRNA by KsgA (Helser et al. 1972);
pseudouridylation of three adjacent residues in tRNAs by
TruA (Hur and Stroud 2007); pseudouridylation of several
tRNA residues by Pus1 (Motorin et al. 1998), Pus2 (Behm-
Ansmant et al. 2007), or Pus7 (Behm-Ansmant et al. 2003);
or methylation of four tRNA positions by Saccharomyces
cerevisiae Trm4 (Motorin and Grosjean 1999). Even with
these multi-site-specific enzymes, however, homologs from
various species generally modify the same residues.
E. coli 16S rRNA contains two 5-methylcytidine (m5C)
residues, located in or near the highly conserved decoding
3These authors contributed equally to this work.
Abbreviations: rRNA, ribosomal RNA; AdoMet, S-adenosyl-L-methionine;
m5C, 5-methyl-cytidine; MALDI mass spectrometry, matrix-assisted laser
desorption ionization mass spectrometry.
Reprint requests to: Gerwald Jogl, Department of Molecular Biology,
Cell Biology and Biochemistry, Brown University, Box G-E129, Provi-
dence, RI 02912, USA; e-mail: Gerwald_Jogl@brown.edu; fax: (401)
863-6114; or Finn Kirpekar, Department of Biochemistry and Molecular
Biology, University of Southern Denmark, Campusvej 55, 5230 Odense M,
Denmark; e-mail: f.kir@bmb.sdu.dk; fax: (+45) 65502467.
Article published online ahead of print. Article and publication date
are at http://www.rnajournal.org/cgi/doi/10.1261/rna.2088310.
1584
RNA (2010), 16:1584–1596. Published by Cold Spring Harbor Laboratory Press. Copyright 2010 RNA Society.
center of the 30S subunit (Fig. 1). An m5C967 modification
is produced by RsmB (also called Fmu), while an m5C1407
modification is produced by RsmF, formerly known as
YebU (Andersen and Douthwaite 2006). T. thermophilus
16S rRNA contains m5C967 and m5C1407, as well as two
additional m5C nucleotides, m5C1400 and m5C1404 (E. coli
rRNA numbering used throughout) (Guymon et al. 2006).
While the m5C967 and m5C1407 modifications are pre-
sumably produced by RsmB and RsmF homologs, respec-
tively, the source of the two additional m5C residues has
been unknown. Here we demonstrate that T. thermophilus
RsmF is a multi-site-specific methyltransferase and, in
contrast to the single-site-specific E. coli RsmF, is respon-
sible for the synthesis of three modifications: m5C1407,
m5C1400, and m5C1404. We also demonstrate that RsmB is
responsible for the synthesis of m5C967 in T. thermophilus
as well as is in E. coli, thereby accounting for all four m5C
modifications of 16S rRNA. We present crystal structures
of T. thermophilus RsmF up to 1.3 A˚ resolution that reveal
a dynamic region in the active site that is absent from the E.
coli RsmF structure, providing a possible explanation for
the expanded recognition capacity of the T. thermophilus
methyltransferase.
RESULTS
Identification of T. thermophilus 16S rRNA
m5C-methyltransferases
With the E. coli RsmB and RsmF protein sequences as
queries, we used conventional BLAST searches (Altschul
et al. 1990) to identify potential homologs encoded by the
T. thermophilus HB8 genome (data not shown). Both RsmB
and RsmF have the highest similarity to the T. thermophilus
protein encoded by TTHA1387 (BLAST scores of 106 and
190, respectively) and second-highest similarity to the pro-
tein encoded by TTHA0851 (BLAST scores of 93 and 81,
respectively). The simplest interpretation of these results is
that TTHA1387 encodes RsmF, responsible for methylation
of C1407, leaving TTHA0851 as the most likely candidate
for the gene encoding RsmB, responsible for methylation of
C967. The similarities of the two E. coli enzymes with other
T. thermophilus proteins were far too low to reveal potential
candidates responsible for methylation of C1400 and C1404.
We next constructed T. thermophilus strains in which
either TTHA0851 or TTHA1387 was inactivated by the
homologous recombination and insertion of a heat stable
kanamycin-resistance gene. 16S rRNA was isolated from
these null mutants and subfragments of z50 nucleotides
(nt) around the regions of interest were further purified,
digested with RNase T1, and analyzed by MALDI mass
spectrometry (Fig. 2). Comparison of the TTHA1387 null
mutant to wild-type T. thermophilus HB8 indicates three
clear differences, each corresponding to the disappearance
of a methyl group (z14.0 Da). The RNase T1 digestion
fragment harboring m5C1407, the nucleotide methylated
by RsmF in E. coli, is absent in the null mutant, indicating
that TTHA1387 is indeed rsmF. The predicted RNase T1
fragment reduced by 14.0 Da is obscured by another RNase
T1 fragment that is present in both the wild-type and
TTHA1387 null mutants (Fig. 2B). Unexpectedly, two addi-
tional RNase T1 fragments are also reduced by 14.0 Da.
One of these contains C1400 while the other contains
C1404. This latter RNase T1 fragment from wild-type
T. thermophilus contains three methyl groups, two on
m4Cm1402 and one on m5C1404 (Guymon et al. 2006),
preventing an unambiguous identification of the missing
methyl group. We therefore performed tandem mass spec-
trometry on the 1402CCCG1405 RNase T1 fragment with two
methyl groups from the TTHA1387 null mutant and com-
pared it with the triply methylated wild-type RNase T1
fragment (Fig. 2C). The clear w2 ions, as well as the less
intense z3 ions, display a 14.0 Da mass difference between
the two samples, showing that the methylations on
m4Cm1402 were not affected by inactivation of TTHA1387.
Tandem mass spectrometry was also performed on the
RNase T1 fragments appearing as a consequence of the lack
of methylations on C1400 and C1407 (data not shown).
As expected, the C1400-containing fragment revealed no
FIGURE 1. Secondary structure diagram of the 39 minor domain of
16S rRNA indicating the position of the three RsmF substrate
nucleotides.
T. thermophilus 16S rRNA methyltransferase RsmF
www.rnajournal.org
1585
indications of a methyl group, whereas
the RNase T1 fragment with C1407 ex-
hibited a fragmentation pattern corre-
sponding to the expected mass overlap
with an RNase T1 fragment of a different
sequence. In summary, our data lead us
to conclude that TTHA1387 encodes an
RsmF m5C methyltransferase responsible
for synthesizing m5C1400, m5C1404, and
m5C1407 in 16S rRNA of T. thermophilus.
The
above
results
left
us
with
TTHA0851 as the sole candidate for
the gene encoding the m5C967 methyl-
transferase. An approach conceptually
identical to that described above re-
vealed that disruption of TTHA0851
reduced the relevant RNase T1 fragment
by 14.0 Da (Supplemental Fig. 1A).
Since this fragment is methylated at
G966 and C967 in the wild-type strain
(Guymon et al. 2006), tandem mass
spectrometry was again performed (Sup-
plemental Fig. 1B), showing that only
the methyl group on C967 was absent. In
agreement with the suggested nomen-
clature for rRNA modifying enzymes
(Andersen and Douthwaite 2006), we
hereafter refer to TTHA0851 as rsmB
due to substrate specificity identical to
the originally identified enzyme from
E. coli (Gu et al. 1999).
Effect of temperature on growth
of the rsmF null mutant
One possible explanation for methyla-
tion of multiple sites by RsmF is that
such additional methyl groups improve
ribosomal function at elevated temper-
ature. To address this possibility, we ex-
amined the effect of temperature on
growth of the rsmF null mutant. Wild-
type T. thermophilus and the rsmF null
mutant were cocultured at three tem-
peratures and these cocultures were se-
rially subcultured for seven cycles of
24 h each. The proportion of wild-type
and rsmF null mutant cells in each mixed
culture was determined by spreading di-
lutions onto TEM plates with or without
kanamycin. After seven cycles, no differ-
ence in the relative proportions of the
wild-type and rsmF mutant was exhib-
ited at 70°C. However, at 60°C, the rsmF
null mutant constituted only around 5%
FIGURE 2. (Legend on next page)
Demirci et al.
1586
RNA, Vol. 16, No. 8
of the population, and at 80°C the rsmF null mutant was
unable to grow at all. Thus, methylation by RsmF appears to
facilitate growth at temperatures outside the optimal growth
temperature.
RsmF substrate preference
The T. thermophilus rsmB and rsmF genes were each cloned
into an E. coli expression plasmid in order to produce
proteins for X-ray crystallography and in vitro methylation
studies. The expression constructs were equipped with
C-terminal histidine6 tags to facilitate protein purification.
While we achieved a high expression level of RsmF, we were
unable to do so with RsmB despite a series of optimization
attempts. Consequently, in vitro substrate and structure
analyses were performed exclusively with RsmF.
E. coli RsmF requires the 30S ribosomal subunit as a
substrate when the activity is assayed in vitro (Andersen and
Douthwaite 2006). We assayed 70S ribosomes, 30S ribosomal
subunits, and 16S rRNA for the ability to serve as substrates
for methylation by RsmF in vitro. 16S rRNA subfragments of
z50 nt around the target sites were purified after the in vitro
assay and analyzed by mass spectrometry as described above.
In vitro methylation at 70°C showed an interesting but rather
complex substrate pattern. RsmF completely methylates
C1400 when either 16S rRNA or 30S subunits are used as
a substrate. It methylates C1404 to z35% with 16S rRNA
and completely with 30S subunits, and it produces only trace
amounts of methylation of C1407 with 16S rRNA and z75%
with 30S subunits (Fig. 3). There were no indications of
the 70S ribosome being a substrate in vitro. Curiously,
T. thermophilus RsmF expressed in an E. coli rsmF null
mutant almost completely methylated, in vivo, positions
C1400 and C1404, but not C1407 (data not shown).
X-ray crystal structures of RsmF
We determined the structure of T. thermophilus RsmF (456
amino acids) in three different crystal forms and in a com-
plex with cofactor AdoMet to up to 1.3 A˚ resolution (Figs.
4, 5). The structure was solved in space group P43 (data set
RsmF1, 1.4 A˚ resolution) by molecular replacement using
a search model generated with the program Modeller
(Eswar et al. 2008) from the catalytic domain of the RsmF
homolog YebU from E. coli (Pdb 2FRX) (Hallberg et al.
2006). The structures of the AdoMet-bound form in space
group P2 (RsmF2, 1.82 A˚ resolution), of the AdoMet-
bound form (RsmF3, 1.3 A˚ resolution), and of the apo-
form (RsmF4, 1.68 A˚ resolution) in space group P21212
were subsequently solved by molecular replacement with
the refined RsmF1 model. There are two molecules in the
asymmetric unit in space groups P43 and P2 and one mol-
ecule in space group P21212. Electron density is generally
well defined in all crystal forms. The majority of residues
(92.0%, 92.3%, 93.1%, and 92.6%) are in the most favored
region of the Ramachandran plot for RsmF1, RsmF2,
RsmF3, and RsmF4, respectively, and there are no residues
in the disallowed region. The final models consist of
residues 5–178, 194–198, and 201–456 and five additional
residues from the histidine6 affinity tag in both chains of
data set RsmF1; residues 2–456 and five affinity-tag
residues in both chains of data set RsmF2; residues 1–456
and six affinity-tag residues in data set RsmF3; and resi-
dues 1–456 and seven affinity-tag residues in RsmF4. The
N-terminal a-amino group was ordered in data sets RsmF3
and RsmF4 and contained additional electron density,
which we interpreted as N-(dihydroxymethyl)-L-methio-
nine, the hydrated form of N-formyl-methionine. Data
collection and refinement statistics are given in Table 1.
The overall structure of RsmF consists of a central
canonical class I methyltransferase catalytic domain with
additional N-terminal and C-terminal
domains (Figs. 4, 5). The catalytic do-
main is formed by a central seven-
stranded b-sheet that is flanked on both
sides by three helices of varying lengths.
An inserted region between strand b7
and helix a11 contains additional heli-
ces a9 and a10, which interact with the
two N-terminal helices a1 and a2. A
second inserted region following strand
b9 includes the short helices a13 and
FIGURE 2. (A) MALDI mass spectra of an RNase T1-digested 16S rRNA subfragment (pos.
1378–1432) from wild-type cells (upper panel) or from the TTHA1387 (putative rsmF) null
mutant (lower panel). Expected digestion products are labeled; fragments affected by the null
mutation are set in italics. (B) Expansion of the signals affected by the TTHA1387 null mu-
tation. The sequence and methylation status of the RNase T1 products are indicated. (C)
MALDI tandem mass spectrometry of the methylated RNase T1 fragment of 16S rRNA (pos.
1404–1407); wild-type cells (upper panel), TTHA1387 null mutant (lower panel). Mass
spectrometric fragments used to deduce the methylation status are labeled. The position of
the backbone fragments (nomenclature according to McLuckey et al. [1992]) in the sequence is
shown. MH+, precursor ion selected for fragmentation; C, cytosine; mC, methylated cytosine;
C>p, cytidine-2´-39-monophosphate; me, methyl group.
FIGURE 3. In vitro methylation with 30S ribosomal subunits or 16S
rRNA from the T. thermophilus rsmF null mutant as a substrate. Effect
on C1400-, C1404-, and C1407-harboring RNase T1 products. In
vitro methylated products are set in italics. *, artifact signal arising
from the enzyme preparation.
T. thermophilus 16S rRNA methyltransferase RsmF
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1587
a14, which interact with helix a12. Furthermore, RsmF
contains three additional smaller domains, an N-terminal
domain consisting of a three-stranded b-sheet and two flank-
ing helices (Fig. 5B, colored in blue), and two C-terminal
domains consisting of four-stranded b-sheets and two or one
helix (Fig. 5B, colored in magenta and red).
Cofactor binding, substrate docking,
and conformational flexibility in the active site
The coordination of AdoMet in the T. thermophilus RsmF
active site is similar to that seen in other class I methyl-
transferases. However, the previously published structure of
E. coli RsmF did not contain the cofactor AdoMet in the
active site, precluding a direct comparison. Both the T.
thermophilus and E. coli RsmF cofactor-binding sites reveal
a new variation for the methyltransferase signature motif I
(Malone et al. 1995), with the highly conserved GxGxG
sequence replaced by 109AAAPG113. The combination of
three alanines and a proline results in a loop conformation
that is very similar to that observed in other methyltrans-
ferases with a GxGxG motif (e.g., RsmC) (Demirci et al.
2008a). In RsmF, the amide hydrogen atom of the last
glycine residue forms a hydrogen bond with the cofactor
carboxy group (Fig. 6). Other key interactions with AdoMet
are well conserved in RsmF. The cofactor adenine ring is
located in a mainly hydrophobic pocket lined by residues
Val134, Pro160, and Leu211. This pocket is open toward the
solvent. The adenine amino group is not specifically recog-
nized and interacts with solvent water molecules. The ribose
hydroxyl groups form hydrogen bonds with Glu133 and
Arg138, and the methionine amino group interacts with
Asp177. The AdoMet cofactor is bound in a cleft in the
RsmF active site, which suggests that substrate cytidine bases
are inserted into the active site in an unstacked conforma-
tion. Inspection of the electrostatic charge distribution re-
veals a large positively charged surface region, which would
be consistent with binding to an RNA surface and modifi-
cation of the substrate base in an unstacked orientation (Fig.
6C). To evaluate the possible orientation of a substrate base
in the active site, we performed computational docking
calculations with the program Dock6 (Lang et al. 2009). The
resulting positions of cytosine and m5C in the presence of
AdoMet in data set RsmF3 are highly similar to each other,
with m5C placed into the active site with its phosphate group
toward a positively charged pocket at the entrance of the
active site cleft (Fig. 6E). The position of the phosphate
group is close to a sulfate molecule that we observed in data
set RsmF1, providing further support for the results of the
docking calculation (Fig. 6F).
Interestingly, we observed that three active site segments
were disordered in data set RsmF1. These segments include
residues 179–193 (including helices a9 and a10 and the
catalytic Cys180), residues 199–200, and the N-terminal
residues 1–5, which interact with the first two segments (Fig.
6E,F, colored in green). We observed electron density for the
intervening residues 194–198, which formed a lattice con-
tact with a neighboring molecule. However, the position of
FIGURE 4. Structure-based sequence alignment of RsmF from T. thermophilus and E. coli. Secondary structure elements of T. thermophilus RsmF
are indicated on top. The color scheme for the secondary structure elements is as in Figure 5A. The position of the variant methyltransferase motif
I is marked with a red box. A flexible region observed in the active site is marked with a green box; residues interacting with the cofactor are
marked with orange boxes.
Demirci et al.
1588
RNA, Vol. 16, No. 8
these five residues was not related to their position in the
other three data sets, suggesting that the extended active
site region between residues 179 and 201 can reorient in the
RsmF structure. This observation suggests that this active
site region is dynamic, which may be important for sub-
strate binding at 72°C, the optimum growth temperature of
T. thermophilus.
DISCUSSION
Substrate recognition mechanisms
We have identified the two enzymes responsible for the
synthesis of the four m5C modifications of T. thermophilus
16S rRNA, and characterized the RsmF methyltransferase
responsible for synthesizing three of these. rRNA modifying
enzymes in bacteria are generally highly specific, with a
one-to-one association between the modifying enzyme and
the modification. A few cases of multitarget ribosome mod-
ifying enzymes have been reported (Helser et al. 1972;
Demirci et al. 2008b), but to our knowledge T. thermophi-
lus RsmF is the first rRNA methyltransferase found to mod-
ify three different nucleotides. Most ribosome modifying
enzymes probably recognize assembly intermediates, and
the data presented here are consistent with that notion.
T. thermophilus RsmF methylates C1400 and C1404 in
vitro using either 16S rRNA or 30S subunits as substrates,
whereas both E. coli (Andersen and Douthwaite 2006) and
T. thermophilus RsmF exclusively utilize 30S subunits as
substrates for methylation of C1407. This may reflect that
C1400 and C1404 methylations do not rely on the asso-
ciation of ribosomal proteins in order to be recognized by
T. thermophilus RsmF. C1407 methylation, in contrast,
depends on both rRNA and the ribosomal protein for the
recognition by RsmF in both T. thermophilus and E. coli.
More puzzling is the observation that T. thermophilus RsmF
does not methylate E. coli ribosomes in vivo on C1407. It
is perhaps worth noting that methylation of C1407 in the
T. thermophilus 30S ribosomal subunit in vitro was less
efficient than methylation of the other two positions,
indicating the need for a particular intermediate assembly
structure or for accessory factors. The only clear in vitro
FIGURE 5. Overall structure of RsmF. (A) Schematic representation of the position of the substrate bases in the 30S ribosomal subunit (Pdb
entry 2WRI) (Gao et al. 2009). C1400, C1404, and C1407 in helix 44 (green) are shown in stick representation with 5-methyl groups as pink
spheres. (B) Schematic stereo representation of the overall structure of RsmF. Secondary structure elements are in orange and yellow for the
catalytic domain, in salmon and blue for additional N-terminal domains, and in magenta and red for C-terminal domains. The flexible region in
the active site including helices a9 and a10 is indicated with an arrow and colored in green. (C) Topology diagram with secondary structure
elements colored as in B.
T. thermophilus 16S rRNA methyltransferase RsmF
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1589
methylation observed at 37°C with T. thermophilus RsmF
was on C1400 with 16S rRNA as the substrate (data not
shown), which is not evidently related to the methylation
pattern in vivo in the heterologous system. It seems unlikely
that the aberrant methylation in the heterologous system
reflects species-specific differences in the mature 30S ribo-
somal subunit, given the extreme sequence and structural
conservation of the decoding site. Instead, it may reflect
differences in 30S subunit assembly in the two organisms,
necessary due to the large difference in growth temperature.
The four m5C residues in T. thermophilus 16S rRNA are
clustered in and around the functionally critical decoding
center at or close to sites of contact with tRNA, mRNA, and
EF-G (Ogle et al. 2002; Selmer et al. 2006; Gao et al. 2009).
m5C1400, m5C1404, and m5C1407 are located in the
subunit body, while m5C967 is located in the subunit head,
about 10 A˚ from m5C1400 (Wimberly et al. 2000). Exam-
ination of the 30S subunit crystal structure (Wimberly et al.
2000) indicates that the three bases methylated by RsmF are
situated in three distinct structural contexts, but provides
few clues to a common mode of substrate recognition.
C1400 is an unpaired base protruding from a sharply bent
segment of rRNA at the junction of helices 43 and 44, while
C1404 and C1407 are engaged in Watson–Crick pairs within
helix 44 (Fig. 4A). The C5 positions of the latter two bases
are not obviously accessible, such that RsmF would need to
approach them from the major groove side. While a flipping
of C1407 out of the helix via the minor groove could allow
access to this base, such a mechanism would be problematic
for C1404, whose minor groove side is packed against the
rest of the 30S subunit.
While m5C1404 and m5C1407 are about 11 A˚ apart
(Selmer et al. 2006), m5C1400 is quite distant from both of
these bases (about 21 and 30 A˚ , respectively). The pro-
truded conformation of m5C1400 in the mature 30S sub-
unit is due in part to base-pairing interactions between
adjacent bases and the 1500 region of 16S rRNA (C1399–
G1504 and G1401–C1501). As the 1500 region is one of the
last segments of 16S rRNA to be synthesized, C1400 could
potentially be positioned much closer to C1404 and C1407
in an assembly intermediate, prior to the formation of the
C1399–G1504 and G1401–C1501 base pairs. RsmF could
utilize a single binding mode to then access all three bases,
further facilitated by its flexible active site domain. Meth-
ylation of C1400 in mature 30S subunits would therefore
involve disruption of the adjacent base pairs. A complete
understanding of the recognition mechanism of these
enzymes will require high-resolution structural data on
assembly intermediate-enzyme complexes. Given the large
number of potential subunit assembly intermediates, pre-
cisely defining the physiological substrate for RsmB and
RsmF will be a formidable task.
TABLE 1. Data collection and refinement statistics
RsmF1
RsmF2
RsmF3
RsmF4
Data collectiona
AdoMet
AdoMet
Space group
P43
P2
P21212
P21212
Cell dimensions
a, b, c (A˚ )
71.0, 71.0, 186.7
66.0, 78.3, 108.1
89.7, 109.0, 51.0
89.8, 109.1, 50.8
a, b, g (°)
90, 90, 90
90, 107.1, 90
90, 90, 90
90, 90, 90
Resolution (A˚ )b
30–1.4 (1.55–1.40)
30–1.82 (1.89–1.82)
30–1.30 (1.34–1.30)
30–1.68 (1.74–1.68)
Rmerge
0.065 (0.59)
0.08(0.38)
0.058(0.36)
0.15 (0.49)
I/sI
29.3(2.15)
12.6 (2.04)
24.3 (2.05)
14.2 (1.73)
Completeness (%)
90.1 (72.6)
97.0(86.5)
95.6(66.3)
99.6 (97.8)
Redundancy
8.9 (5.4)
2.8(2.0)
5.2(2.1)
6.4 (3.9)
Refinement
Resolution (A˚ )
30–1.4 (1.42–1.40)
30–1.82 (1.84–1.82)
30–1.30 (1.32–1.30)
30–1.68 (1.69–1.68)
Number of reflections
161,955 (4356)
91,762 (2583)
119,490/2640
109,375 (3244)
Rwork/Rfree
0.169/0.189 (0.227/0.263)
0.162/0.194 (0.222/0.259)
0.177/0.191 (0.233/0.234)
0.173/0.192 (0.217/0.266)
Number of atoms
Protein
6766
7117
3598
3574
Ligand/ion
20
54
27
1
Water
1486
1285
856
665
B-factors
Protein
24.1
20.4
15.9
17.2
Ligand/ion
34.5
21.6
17.5
19.1
Water
38.7
36.3
33.6
33.6
RMSDs
Bond lengths (A˚ )
0.009
0.006
0.005
0.004
Bond angles (°)
1.22
1.04
1.14
0.94
aOne crystal used for each data set.
bThe highest resolution shell is shown in parentheses.
Demirci et al.
1590
RNA, Vol. 16, No. 8
Structural comparison of RsmF with related
methyltransferases
A database search with Dali (Holm et al. 2008) confirmed
the structural similarity of the T. thermophilus and E. coli
(PDB 2FRX) (Hallberg et al. 2006) RsmF homologs, which
superimpose with a root-mean-square deviation (RMSD)
of 1.6 A˚ for 342 Ca atoms (Fig. 7A) and
are the only two structures in the Pro-
tein Database with this domain organi-
zation. Even so, substantial structural
differences are observed in most of the
loop regions and for the long connect-
ing loop between the methyltransferase
domain and the first C-terminal domain.
While in the active site, the positions
of residues in the cofactor-binding site
and of the two cysteine residues are
conserved (Fig. 7B), there are a number
of positively charged residues (Arg30,
Arg190, Arg194, His195, and Arg203) in
the T. thermophilus structure that are
absent from the E. coli enzyme (Fig. 7B).
Three of these are located in the flexible
region, and the combination of a posi-
tive charge and flexibility close to the
active site is suggestive of a functional
contribution of this region to the mul-
tisite specificity of T. thermophilus RsmF
(Figs. 6C,D, 7B). Methylation of three
rRNA positions may require an increase
in the enzyme’s structural dynamics in
order to accommodate the 30S subunit
in slightly different orientations. Similar
observations have been made for other
multi-site-specific methyltransferases in-
cluding KsgA, which modifies two adja-
cent adenosines in the 30S ribosomal
subunit (O’Farrell et al. 2004; Demirci
et al. 2009), and the PrmA ribosomal
protein methyltransferase, which under-
goes dramatic interdomain movements
to modify multiple lysine residues and the
N-terminal a-amino group on the same
substrate protein (Demirci et al. 2007,
2008b).
The second C-terminal domain in
RsmF is related to the RNA-binding
PUA
(pseudouridine
synthase
and
archaeosine transglycosylase) domains
(Perez-Arellano et al. 2007). The RlmI
methyltransferase,
which
produces
m5C1962 in 23S rRNA (Purta et al.
2008) also contains a PUA domain
(Sunita et al. 2008), although it is
N-terminal to the catalytic methyltransferase domain and
in a different orientation. PUA domains contain six
b-strands, which form a central pseudobarrel closed by
a short 310-helix. A comparison of the C-terminal domain
in RsmF with a typical PUA domain in archaeosine trans-
glycosylase (ArcTGT, Pdb entry 1J2B) reveals that the
central fold is similar (53 Ca atoms align with an RMSD
FIGURE 6. Substrate docking and conformational flexibility. (A) Cofactor-binding site in
RsmF. Bound AdoMet is shown in blue sticks. Hydrogen bonds to coordinating residues are
indicated. (B) Final sA-weighted 2mFO-DFC electron density map of the cofactor-binding site
(data set RsmF2) contoured at the 1s level. (C,D) Comparison of the electrostatic surface
charge distribution between RsmF from T. thermophilus and E. coli. The location of the
C-terminal domains and of the flexible region (labeled ‘‘FLEX’’) is indicated with circles.
AdoMet and docked m5C are shown as sticks. AdoMet from the T. thermophilus structure is
shown with the E. coli structure for comparison. (E) The modeled position of m5C docked into
the active site of RsmF. m5C and AdoMet are shown in tan and blue sticks, respectively.
Residues in the flexible region are shown in green. (F) The active site region in data set RsmF1.
A sulfate ion is observed close to the position of the m5C phosphate group. Residues 194–198 in
the flexible region interacting with a neighboring RsmF molecule are indicated with an arrow.
T. thermophilus 16S rRNA methyltransferase RsmF
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1591
of 1.8 A˚ ), but that several connecting loop regions are
substantially shorter and two a-helices and one b-strand of
the pseudobarrel are absent. Thus, this PUA-like domain
differs considerably from typical PUA domains. However,
the similarity to RNA-binding PUA domains and the
positive surface charge distribution observed in both RsmF
structures are suggestive for a conserved function of the
PUA-like domain in RNA recognition.
The next most closely related structure is E. coli RsmB
(474 residues, PDB 1SQF) (Foster et al. 2003). A total of
264 Ca atoms can be aligned with an RMSD of 1.5 A˚
between RsmB and RsmF. Both enzymes retain the same
organization for the N-terminal domains and the core
methyltransferase domain. However, an additional 140-
residue N-terminal RNA-binding domain provides sub-
strate specificity to RsmB, whereas the two C-terminal
domains following the core methyltransferase domain (160
residues) are likely to determine substrate recognition by
RsmF. Thus, these two enzymes have evolved substrate
specificity via acquisition of additional,
unrelated RNA recognition domains.
While there are as yet no enzyme–
substrate complexes for rRNA m5C-
methyltransferases, insights into the RsmF
catalytic mechanism can be gleaned from
a comparison with the RlmD and TrmA
m5U methyltransferases in covalent in-
termediate complexes with RNA oligonu-
cleotides (Lee et al. 2005; Alian et al.2008).
DNA and RNA m5C methyltransferases
use a thiol from a catalytic cysteine residue
to attack the six-position of the pyrimi-
dine base to activate the five-position for
methyl group transfer (Liu and Santi
2000). Cys180 and Cys230 in RsmF are
positioned equivalently to the catalytic
Cys324 and the catalytic base Glu358 of
TrmA. The substrate nucleotides insert
into TrmA and RsmF in unrelated di-
rections, consistent with a lack of struc-
tural homology outside the methyltrans-
ferase domains. Nevertheless, the C5
positions of the pyrimidine rings and of
the 5-methyl carbons are quite similar
with respect to the catalytic cysteine resi-
dues and the AdoMet cofactor. The same
structural homology of the active site
geometry can be observed in comparison
with the RlmD methyltransferase (Sup-
plemental Fig. 2; Lee et al. 2005).
A possible origin of T. thermophilus
RsmF
T. thermophilus RsmF shows the highest
similarities with proteins from close relatives, namely,
Thermus aquaticus and two Meiothermus species of the
Thermaceae family. Remarkably, the next highest similarities
(BLAST scores between 267 and 359) are with the NOL1/
NOP2/Sun proteins from the Gram-positive Firmicutes
phylum (Supplemental Fig. 3). This similarity, together
with the fact that the Thermaceae family and most members
of the Firmicutes identified in Supplemental Figure 3 are
thermophilic, suggest that rsmF has undergone horizontal
transfer between the Thermaceae family and members of the
Firmicutes phylum. Thus, we speculate that this version of
the RsmF protein, which catalyzes methylation of three
cytidines, may be adaptive for existence in thermally
challenging environments. The effect of the loss of methyl-
ation by RsmF on growth at different temperatures is
consistent with this notion. Our hypothesis of horizontal
transfer of rsmF predicts that RsmF of other members of the
Thermaceae family and members of the Firmicutes phylum
will also be found to introduce multiple m5C modifications.
FIGURE 7. Comparison with other methyltransferases. (A) Differences between the overall
structures of RsmF from T. thermophilus (orange) and E. coli (green). AdoMet bound in the T.
thermophilus structure is shown as blue sticks. (B) Comparison of the active site region in both
enzymes. Residues in the cofactor-binding site and in the flexible region in the T. thermophilus
enzyme are shown as sticks. (C) Comparison between RsmF (orange) and a substrate complex
structure of the TrmA methyltransferase (cyan/blue). (D) Comparison of the active site region
illustrating the differences in the insertion direction of the substrate base in RsmF (m5C in
light orange) and in TrmA (m5U in cyan).
Demirci et al.
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RNA, Vol. 16, No. 8
MATERIALS AND METHODS
Cloning of the T. thermophilus rsmB and rsmF genes
The T. thermophilus HB8 loci TTHA0851 (GenBank accession
number BAD70674) and TTHA1387 (GenBank accession number
BAD71210) were PCR amplified from genomic DNA and purified
via the High Pure PCR Template Preparation Kit (Roche). The
100 mL PCRs contained 150 ng DNA, 10 mM of each primer,
10 mM dNTP, 1 unit Phusion DNA polymerase (Finnzymes), and
1x Phusion HF buffer. Primers for rsmB amplification were 59-CC
CTGGACATATGAGGGCCGG-39 and 59-GGCCAAGATCTTGCC
TGAGAG-39, and the temperature cycling was as follows: 98°C/30
sec; 30X (98°C/10 sec, 59°C/20 sec, and 72°C/36 sec); and 72°C/420s.
Primers for rsmF amplification were 59-GCTAGGGTACACATA
TGCTGCCC-39 and 59-GCACGGGGGTGAGATCTAAGCCC-39,
and the temperature cycling was as follows: 98°C/30 sec; 30X
(98°C/10 sec, 62°C /20 sec, 72°C/42 sec); and 72°C/420 sec. The
desired PCR products were purified from agarose gels using the
GFX PCR purification kit (GE Healthcare). The PCR fragments
were digested with NdeI and BglII and inserted into the expression
vector pLJ102 (Andersen and Douthwaite 2006), generating
isopropyl-1-thio-b-D-galactopyranoside (IPTG)-inducible genes
for the recombinant proteins with a C-terminal histidine6 tag.
The constructs (designated pLJ102-RsmB and pLJ102-RsmF) were
used to transform an rsmF-deletion derivative of E. coli CP79
(Andersen and Douthwaite 2006).
Deletion of the T. thermophilus rsmB and rsmF genes
Constructs for inactivation of the T. thermophilus rsmB and rsmF
genes were made by inserting the gene for a heat tolerant
kanamycin (HTK) nucleotidyltransferase (Hoseki et al. 1999) into
the methyltransferase parts of either pLJ102-RsmB or pLJ102-
RsmF. The htk part of pUC18-htk (Hashimoto et al. 2001) was
amplified by PCR with primers that introduced an upstream AvrII
site and a downstream SacI site into the product for later disrup-
tion of rsmB. For rsmF disruption, the PCR primers introduced
SacI restriction enzyme sites both upstream of and downstream
from the htk gene. These sites were used to insert the PCR prod-
ucts into pLJ102-RsmB and pLJ102-RsmF to form the plasmids
pLJ102-RsmBThtk and pLJ102-RsmFThtk, which were propa-
gated in the E. coli strain Top10 (Invitrogen). T. thermophilus
HB8 was transformed with pLJ102-RsmBThtk or pLJ102-
RsmFThtk selecting for kanamycin resistance as described by
others (Hashimoto et al. 2001; Cameron et al. 2004). Kanamycin-
resistant transformants were restreaked twice. Gene disruptions
were verified by PCR with primers distal to the interrupted rsmB
or rsmF genes on genomic DNA; resulting PCR products were
characterized by sequencing.
Growth competition assays
Wild-type and rsmF null mutant liquid cultures were grown at
70°C to saturation, then equal numbers of cells from each were
mixed and incubated in 5 mL TEM medium at 60°C, 70°C, or
80°C. After growth for 24 h, 100 mL of the 60°C culture, 10 mL of
the 70°C culture, and 1000 mL of the 80°C culture were trans-
ferred to a fresh 5-mL medium and incubated at the respective
temperatures for another 24 h. This was repeated in independent
triplicates for seven cycles. Samples of 1 mL were collected at each
dilution and half was plated on TEM plates without antibiotic and
the other half was plated on TEM plates with 30 mg/mL
kanamycin. The plates were incubated at 70°C.
Purification of T. thermophilus ribosomal subunits
and ribosomes
T. thermophilus culture (1 L) was grown in TEM media (contain-
ing 30 mg/mL of kanamycin when appropriate) with shaking at
70°C to an OD600 = 0.6. Cells were harvested and washed once
with 100 mL of buffer A (10 mM NH4Cl, 20 mM MgCl2, 100 mM
KCl, and 10 mM Tris-HCl [pH 7.5]), then suspended in 10 mL of
buffer A, and disrupted by sonication. The lysate was cleared by
centrifugation twice in a Beckman JA20 rotor at 16,000 rpm for
10 min at 4°C. Crude ribosomes were collected by centrifugation
in a Beckman Ti50 rotor at 19,000 rpm for 19 h at 4°C, and
dissolved in buffer A. 70S ribosomes were obtained by centrifu-
gation of 100 A260 units of crude ribosomes through a 10%–40%
sucrose gradient (200 mM NH4Cl, 20 mM MgCl2, 20 mM Tris-
HCl [pH 7.5]) in a Beckman SW28 rotor at 20,000 rpm for 18 h at
4°C. Fractions containing intact 70S ribosomes were pooled and
concentrated by centrifugation in a Beckman Ti50 rotor at 40,000
rpm for 22 h at 4°C, washed, and dissolved in buffer A, and stored
at 80°C.
50S and 30S ribosomal subunits were obtained by adjusting
100 A260 units of crude ribosomes (10 mM NH4Cl, 2 mM MgCl2,
100 mM KCl, and 10 mM Tris-HCl [pH 7.5]) and passing
through a 5%–20% sucrose gradient (200 mM NH4Cl, 1 mM
MgCl2 and 20 mM Tris-HCl [pH 7.5]) in a Sorvall AH-629 rotor
at 20,000 rpm for 18 h at 4°C. After pooling of the relevant
fractions, the subunits were adjusted to 10 mM MgCl2 and
pelleted by centrifugation in a Beckman Ti50 rotor at 40,000
rpm for 22 h at 4°C, washed with and dissolved in 10 mM NH4Cl,
10 mM MgCl2, 100 mM KCl, and 10 mM Tris-HCl (pH 7.5), and
stored at 80°C.
Isolation of 16S rRNA and subfragments
from T. thermophilus and E. coli
Water (400 mL) was added to 100 mL of 30S ribosomal subunits
and the rRNA was extracted with 500 mL phenol, phenol/
chloroform, and chloroform. rRNA was ethanol precipitated
and dissolved in water. Purification of 16S rRNA subfragments
was performed as previously described (Andersen et al. 2004).
Briefly, 16S rRNA was hybridized to an excess of oligodeoxynu-
cleotide complementary to either the region 944–990 or the region
1378–1432. Single-stranded nucleic acids were digested with
Mung Bean Nuclease and RNase A. The resulting mixture was
separated on a polyacrylamide gel. Bands were visualized by ethid-
ium bromide staining, excised, and eluted.
E. coli CP79 with the endogenous rsmF inactivated, but com-
plemented with the T. thermophilus homolog on the plasmid
pLJ102-RsmF, were grown at 37°C to an OD450 = 0.45 in 200 mL
of LB medium containing 100 mg/L of ampicillin. RsmF expres-
sion was induced by addition of IPTG to 1 mM, and incubation
for another 3 h. Cells were harvested by centrifugation at 4°C,
washed in 100 mL TMN buffer (50 mM Tris-HCl [pH 7.8],
10 mM magnesium acetate, 100 mM NH4Cl), and resuspended in
2 mL TMN buffer prior to lysis by sonication (7 3 30 sec on ice)
T. thermophilus 16S rRNA methyltransferase RsmF
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1593
and removal of debris by centrifugation (10 min/14,000 rpm/4°C/
microcentrifuge). Total RNA was recovered from the supernatant
by phenol extraction and ethanol precipitation. A 16S rRNA
subfragment was isolated as described above using an oligodeoxy-
nucleotide complementary to the region 1378–1432.
In vitro methylation
Reactions contained 50 pmol of 16S rRNA, 30S subunits, or 70S
ribosomes from the T. thermophilus TTHA1387 null mutant as the
substrate in a total volume of 100 mL (containing 100 mM NH4Cl,
10 mM MgCl2, 40 mM Hepes [pH 7.5]), 6 mM b-mercaptoetha-
nol, and 10% glycerol (prepared as a two times concentrated stock
solution), 1.5 mM S-adenosyl methionine, and 2 mg of recombi-
nantly expressed RsmF (see below).
For the reaction at 70°C, water and stock buffer were mixed and
left at room temperature for 15 min. Then a substrate, an enzyme
and S-adenosyl methionine were added and incubated at 70°C for
1 h. The 37°C reaction was started by mixing water and buffer
followed by 15 min at room temperature; the substrate was added
and the mixture transferred to 50°C for 5 min. After cooling to
37°C, S-adenosyl methionine and an enzyme were added and the
incubation continued for 1 h. Reactions were stopped by phenol/
chloroform extraction and the rRNA was recovered by ethanol
precipitation before purification of 16S rRNA subfragments as
described above. Control reactions without enzyme or S-adenosyl
methionine were carried out in all instances.
RNase T1 digestion and mass spectrometry
A purified 16S rRNA subfragment (1–2 pmol) was incubated with
2 units RNase T1 (Roche) and 50 mM 3-hydroxypicollinic acid
(3-HPA) in a total volume of 2 mL for 4 h at 37°C. MALDI mass
spectrometry was performed either on an ABI voyager STR in-
strument or a Waters Q-TOF MALDI instrument; MALDI tan-
dem mass spectrometry was done on a Waters Q-TOF MALDI
instrument. All spectra were recorded in positive ion mode using
3-HPA as the matrix. Experimental details were as previously de-
scribed (Douthwaite and Kirpekar 2007).
Protein expression and purification for crystallization
E. coli BL21 (DE3) (Invitrogen) containing pLJ102-RsmF was
grown to midlog phase in LB media at 37°C in the presence of
200 mg/mL ampicillin. Protein expression was induced at 20°C
with 400 mM IPTG. Cells were pelleted after 18 h by centrifuga-
tion at 4000 rpm for 20 min at 4°C and lysed by ultrasonication
on ice in a buffer containing 20 mM Tris-HCl (pH 8.5), 300 mM
NaCl, 5 mM b-mercaptoethanol, 0.1% Triton X-100, and 5%
glycerol. Cell debris and membranes were pelleted by centrifuga-
tion at 11,000 rpm for 30 min at 4°C. The soluble E. coli proteins
were precipitated by heat treatment at 65°C for 30 min and
pelleted by centrifugation at 11,000 rpm at 4°C for 30 min.
Soluble C-terminally hexahistidine-tagged T. thermophilus RsmF
was further purified by affinity chromatography with nickel-
nitrilotriacetic acid resin (Qiagen). Untagged proteins were re-
moved with buffer containing 20 mM Tris-HCl (pH 8.5), 250 mM
NaCl, and 1 mM imidazole (pH 8.5). Recombinant RsmF was
then eluted with the same buffer containing 150 mM imidazole.
The protein was then purified by cation exchange chromatogra-
phy (SP) (GE Healthcare) at pH 8.5, using a linear gradient of
10 mM to 1 M NaCl concentration. RsmF fractions were pooled
and concentrated and applied to a size-exclusion S200 column (GE
Healthcare) equilibrated with a buffer containing 20 mM Tris-HCl
(pH 8.5) and 200 mM NaCl. Purified RsmF was concentrated to
13 mg/mL for crystallization trials. The C-terminal hexahistidine
tag was not removed for crystallization. For the production of
selenomethionyl proteins, the expression construct was trans-
formed into B834 (DE3) cells (Novagen). The bacterial growth
was carried out in defined LeMaster medium (Hendrickson et al.
1990), and the protein was purified using the same protocol as for
the unmodified protein. To form the RsmF-AdoMet complex,
purified RsmF was mixed with 4 mM AdoMet incubated at 60°C
for 15 min and slowly cooled to room temperature before
performing crystallization experiments.
Crystallization of RsmF
All crystals were obtained using the microbatch technique under
oil at 4°C. To obtain the RsmF1 crystal form, 1 mL of protein
solution was mixed with the reservoir solution containing 20%
(w/v) PEG3350 and 200 mM sodium sulfate decahydrate (pH
6.6). Initial crystals grew over the course of 1–2 wk with maxi-
mum dimensions of 0.3 3 0.3 3 0.2 mm. To obtain the RsmF2
crystal form, 1 mL of the RsmF–AdoMet complex was mixed with
the reservoir solution containing 200 mM NaCl, 12% w/v
PEG8000 and 100 mM HEPES-KOH (pH7.5). Initial crystals
grew over the course of 2–3 wk with maximum dimensions of
0.1 3 0.4 3 0.4 mm. To obtain the RsmF3 crystal form, 1 mL of
the RsmF–AdoMet complex was mixed with the reservoir solution
containing 10% w/v PEG1000, 200 mM NaCl, and 100 mM Tris-
HCl (pH 8.5). The initial crystals grew over the course of 1–2 wk
with maximum dimensions of 0.05 3 0.3 3 0.4 mm. To obtain
the RsmF4 crystal form, 1 mL of the RsmF–AdoMet complex
solution was mixed with a reservoir solution containing 160 mM
magnesium chloride hexahydrate, 80 mM Tris-HCl (pH 8.5), and
24% w/v PEG4000. Initial crystals grew over the course of 1–2 wk
with maximum dimensions of 0.05 3 0.3 3 0.3 mm. RsmF1
crystals were gradually dehydrated by increasing the PEG3350 to
30% w/v and then cryoprotected in a mother liquor supplemented
with 25% v/v glycerol and then flash-frozen by being plunged into
liquid nitrogen. RsmF2 crystals were cryoprotected in a mother
liquor supplemented with 20% v/v ethylene glycol and then flash-
frozen by being plunged into liquid nitrogen. RsmF3 crystals
were cryoprotected by gradually increasing the concentration of
PEG1000 to 30% and then flash-frozen by being plunged into
liquid nitrogen. RsmF4 crystals were cryoprotected in a mother
liquor supplemented with 20% glycerol and then flash-frozen by
being plunged into liquid nitrogen.
Data collection
X-ray diffraction data for RsmF1, RsmF2, and RsmF4 crystals
were collected on a MAR CCD detector at the X4C beamline of
the National Synchrotron Light Source in Brookhaven at a wave-
length of 0.979 A˚ and 180°C. Diffraction data for RsmF3 crystals
were collected on an ADSC CCD detector at the X4A beamline of
the National Synchrotron Light Source in Brookhaven at a wave-
length of 0.979 A˚ and 180°C. Diffraction data for RsmF1 in
space group P43 were collected to 1.4 A˚ resolution with cell
Demirci et al.
1594
RNA, Vol. 16, No. 8
dimensions a = 71.0 A˚ , b = 71.0 A˚ , and c = 186.7 A˚ . Diffraction
data to 1.82 A˚ for RsmF2 were collected in space group P2 with
cell dimensions a = 66.0 A˚ , b = 78.3 A˚ , and c = 108.1 A˚ .
Diffraction data to 1.29 A˚ for RsmF3 were collected in space group
P21212 with cell dimensions a = 89.7 A˚ , b = 109.0 A˚ , and c = 51.0
A˚ . Diffraction data to 1.68 A˚ for RsmF4 were collected in space
group P21212 with cell dimensions a = 89.8 A˚ , b = 109.1 A˚ , and c =
50.8 A˚ . A single crystal was used for each data set. The diffraction
images were processed and scaled with the HKL2000 package
(Otwinowski and Minor 1997). The data processing statistics are
summarized in Table 1.
Structure determination and refinement
The RsmF structure was solved by molecular replacement with the
program Phaser (McCoy et al. 2007) from the CCP4 program
suite (Bailey 1994) in space group P43 to 1.4 A˚ resolution (data set
RsmF1). The initial search model was built with the program
Modeller (Eswar et al. 2008) from the catalytic domain of E. coli
YebU (Pdb code 2FRX). After the placement of two RsmF
catalytic domains in the asymmetric unit and the initial re-
finement with Refmac (Murshudov et al. 1997), the model was
further rebuilt with ARP/wARP (Langer et al. 2008). The resulting
model was 90% complete and manually checked and completed
with Coot (Emsley and Cowtan 2004). Final crystallographic re-
finement was performed with the program Phenix (Adams et al.
2002). The other crystal forms were subsequently solved by
molecular replacement. The atomic coordinates from the RsmF4
model were then used for initial refinement of the RsmF–AdoMet
complex structure in space group P21212 (RsmF3). There are two
molecules in the asymmetric unit in data sets RsmF1 and RsmF2,
and one molecule in RsmF3 and RsmF4. The crystallographic
R/Rfree factors are 0.17/0.19, 0.16/0.19, 0.18/0.19, and 0.17/0.19
for the four data sets: RsmF1, RsmF2, RsmF3, and RsmF4,
respectively. The stereochemical quality of the model was assessed
with Procheck (Laskowski et al. 1993). The Ramachandran sta-
tistics (most favored/additionally allowed/generously allowed/
disallowed) are 91.9%/8.1%/0.0%/0.0% for RsmF1, 91.9%/8.1%/
0.0%/0.0% for RsmF2, 93.6%/6.4%/0.0%/0.0% for RsmF3, and
92.5%/7.5%/0.0%/0.0% for RsmF4. The refinement statistics are
summarized in Table 1. Figures were generated using Pymol
(DeLano 2002).
Atomic coordinates
Coordinates and structure factors have been deposited in the
Protein Data Bank with accession codes 3M6U, 3M6V, 3M6W,
and 3M6X for data sets RsmF1, RsmF2, RsmF3, and RsmF4,
respectively.
SUPPLEMENTAL MATERIAL
Supplemental material can be found at http://www.rnajournal.org.
ACKNOWLEDGMENTS
We thank John Schwanof and Randy Abramowitz for access to the
X4A and X4C beamlines at the National Synchrotron Light Source.
This work was supported by grants GM19756 and GM19756-37S1
from the National Institutes of Health.
Received January 14, 2010; accepted April 26, 2010.
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|
3M6Z
|
Crystal structure of an N-terminal 44 kDa fragment of topoisomerase V in the presence of guanidium hydrochloride
|
Structures of minimal catalytic fragments of topoisomerase V
reveals conformational changes relevant for DNA binding
Rakhi Rajan*, Bhupesh Taneja*,†, and Alfonso Mondragón*,‡
* Department of Biochemistry, Molecular Biology and Cell Biology, Northwestern University, 2205
Tech Dr, Evanston, IL 60208
Summary
Topoisomerase V is an archaeal type I topoisomerase that is unique among topoisomerases due to
presence of both topoisomerase and DNA repair activities in the same protein. It is organized as an
N-terminal topoisomerase domain followed by 24 tandem helix hairpin helix (HhH) motifs.
Structural studies have shown that the active site is buried by the (HhH) motifs. Here we show that
the N-terminal domain can relax DNA in the absence of any HhH motifs and that the HhH motifs
are required for stable protein-DNA complex formation. Crystal structures of various
topoisomerase V fragments show changes in the relative orientation of the domains mediated by a
long bent linker helix, and these movements are essential for the DNA to enter the active site.
Phosphate ions bound to the protein near the active site helped model DNA in the topoisomerase
domain and shows how topoisomerase V may interact with DNA.
Introduction
DNA topoisomerases are enzymes found in all forms of life (bacteria, eukarya, and archaea)
and they regulate the topological state of DNA inside the cell. They form a transient break in
a single or double stranded DNA and allow the passage of another single or double DNA
strand through the break, before resealing the break (Champoux, 2001) (Schoeffler and
Berger, 2008). As a result of this, topoisomerases can relax supercoiled DNA, help in the
segregation of DNA strands following replication, and lead to the formation and resolution
of knots and catenates (Gellert, 1981). Topoisomerases participate in many aspects of DNA
metabolism, such as replication, recombination, and transcription (Champoux, 2001). In
addition, they are targets of various anti-cancerous drugs and anti-bacterial agents (Maxwell,
1999; Pommier, 1998; Rothenberg, 1997; Wang et al., 1997).
DNA topoisomerases are broadly classified into two types, type I and type II enzymes. Type
I enzymes cleave a single strand of a DNA molecule and pass another single or double
stranded DNA through the break before resealing the opening. Type II enzymes cleave both
‡Corresponding author: Phone: 847-491-7726, Fax: 847-467-6489, a-mondragon@northwestern.edu.
†Present address: Institute of Genomics and Integrative Biology, CSIR, Delhi, India
Protein data bank accession codes
The final structure factors and coordinates of Topo-31, Topo-44 Form I, Form II, and Form III have been deposited in the Protein Data
Bank with accession codes 3M7G, 3M7D, 3M6K, and 3M6Z respectively.
Supplementary data
Supplementary data are available at Structure Journal Online.
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Author Manuscript
Structure. Author manuscript; available in PMC 2011 July 14.
Published in final edited form as:
Structure. 2010 July 14; 18(7): 829–838. doi:10.1016/j.str.2010.03.006.
NIH-PA Author Manuscript
NIH-PA Author Manuscript
NIH-PA Author Manuscript
strands of a double stranded DNA in concert and pass another double stranded DNA through
the break. Type I enzymes use the torsional energy stored in the supercoiled DNA to drive
DNA relaxation and hence they do not require high energy cofactors, such as ATP, for their
activity (Baker et al., 2009) Type II enzymes, on the other hand, require ATP and Mg2+ for
their activity. Type I topoisomerases are further subdivided into three subtypes: IA, IB, and
IC (Forterre et al., 2007). Type IA and IB enzymes have been studied extensively (Baker et
al., 2009) and there is ample information available about their general mechanism of DNA
relaxation and the mode of DNA binding. Type IC, on the other hand, is a relatively new
subtype. Currently topoisomerase V is the only member of this family and it has been
identified only in the Methanopyrus genus. Previously, topoisomerase V had been
considered as a type IB enzyme based on its biochemical characteristics (Slesarev et al.,
1993), but the crystal structure of an N-terminal 61 kDa of topoisomerase V (Topo-61)
(Taneja et al., 2006) revealed a completely new fold without similarity to other
topoisomerases or any other known protein. Furthermore, the orientation of the putative
active site residues is also different from other type I topoisomerases, suggesting a different
mechanism of cleavage and religation of DNA. These observations, together with the lack of
sequence similarity, indicated that topoisomerase V defines a new subtype of type I enzymes
(Forterre, 2006).
Topoisomerase V was identified in Methanopyrus kandleri, an extremophile isolated from a
deep-water ‘black smoker’ chimney in the Gulf of California (Huber et al., 1989). The
enzyme is active at very high temperatures (122°C) and high salt concentrations (0.65 M
NaCl and 3.1 M potassium glutamate). The unusual characteristic of topoisomerase V is that
it has both topoisomerase and DNA repair activities in the same polypeptide (Belova et al.,
2001). Based on the sequence analysis of topoisomerase V, it has been predicted that the
protein contains 24 helix-hairpin-helix (HhH) DNA binding motifs arranged as 12 (HhH)2
domains around the N-terminal topoisomerase domain (Belova et al., 2002) (Figure 1A).
Some of these (HhH)2 domains are involved in the apurinic/apyrimidinic (AP) site-
processing activity, but the exact location of the repair active site is not known yet.
Topoisomerase V can relax both positively and negatively supercoiled DNA without the
need for metal cations or high energy cofactors. Single molecule experiments have shown
that topoisomerase V relaxes DNA by a constrained swiveling mechanism, relaxing around
12 turns of DNA per relaxation cycle (Taneja et al., 2007). Type IB enzymes, which also use
a constrained swiveling mechanism for DNA relaxation, relax around 19 turns of DNA per
relaxation cycle (Koster et al., 2005).
The structure of Topo-61 showed that the topoisomerase domain is mainly alpha helical and
that the first four (HhH)2 domains curl around the topoisomerase domain (Taneja et al.,
2006) (Figure 1B). The topoisomerase and (HhH)2 domains are joined by a long bent helix,
termed the “linker helix”. Three of the five putative active site residues are present in a
helix-turn-helix (HTH) domain and the other two are present in an intervening loop and a
helix. The active site residues are buried by the first (HhH)2 domain and it has been
suggested that large conformational changes will be needed for the DNA to access the active
site of topoisomerase V (Taneja et al., 2007). Here we present data that shows that the N-
terminal 31 kDa fragment of topoisomerase V (Topo-31) has topoisomerase activity,
consistent with previous predictions based on the structure. In addition, we show that the
Topo-44 fragment (N-terminal 44 kDa fragment of topoisomerase V) can form a stable
protein-DNA complex, emphasizing the need of the (HhH)2 domains for binding DNA. We
determined a crystal structure of (Topo-31) fragment, which has only the topoisomerase
domain, and three different crystal structures of the Topo-44 fragment, which includes the
topoisomerase domain and three tandem HhH motifs. In all structures, the topoisomerase
domain is very similar. In contrast, the structures of Topo-44 show conformational changes
in the linker helix resulting in variable orientations of the (HhH)2 domains when compared
Rajan et al.
Page 2
Structure. Author manuscript; available in PMC 2011 July 14.
NIH-PA Author Manuscript
NIH-PA Author Manuscript
NIH-PA Author Manuscript
to the Topo-61 structure. Phosphate ions are present in the vicinity of the topoisomerase
active site in two of the Topo-44 structures. Some of the catalytic residues interact with the
phosphate ions and may mimic contacts with DNA. These observations suggest that the
movement of the (HhH)2 domains is mediated by the linker helix and helps expose the
topoisomerase active site to facilitate DNA binding. In addition, the location of the
phosphate ions suggests a possible path for the DNA and the way the active site residues
interact with it.
Results
The topoisomerase domain can relax DNA
DNA relaxation assays using different topoisomerase V fragments showed that the
topoisomerase domain alone is capable of relaxing DNA. Topoisomerase V fragments with
different numbers of (HhH)2 domains, Topo-31, Topo-44, and Topo-78, were studied using
relaxation assays. Topo-31 has no (HhH)2 domains, Topo-44 has one full and one partial
(HhH)2 domain, while Topo-78 has eight full (HhH)2 domains, including a putative DNA
repair domain. In addition to standard conditions, the effect of different pH conditions and
presence of magnesium ions were also tested. The experiments show that Topo-31 is
capable of relaxing DNA, despite the absence of the (HhH)2 domains (Figure 2B). A pH
profile analysis for the DNA relaxation assays showed that Topo-78 relaxes DNA over a
wider pH range (pH 5 to 9), while Topo-31 and Topo-44 relax DNA optimally at pH 5
(Figure 2A, 2B, 2C). In addition, magnesium is not required for the reaction, but stimulates
it at all pH values (Figure 2B, 2C). Topo-78 can relax DNA to the same extent with lower
amounts of protein (0.1 μg/reaction) compared to Topo-44 (~1.5 μg/reaction) and Topo-31
(~9 μg/reaction). This could be due to the enhanced DNA binding facilitated by the (HhH)2
domains. Together, these results suggest that, even though the (HhH)2 domains are
dispensable for topoisomerase activity, they enhance DNA relaxation activity. In addition,
the pH dependence of the DNA relaxation activity indicates that the reaction is likely to
involve side chains with ionizable groups in the low pH range, such as glutamates. Finally,
the magnesium independence of the reactions confirms that even the smallest fragments do
not require metals for activity, although magnesium has a stimulatory effect. This may be
due to favorable interactions of the cations with DNA.
The (HhH)2 domains enhance DNA binding affinity
EMSA experiments with different fragments of topoisomerase V and DNA showed that
(HhH)2 domains could help in the formation of a stable protein-DNA complex. Various
topoisomerase V fragments (Topo-31, Topo-44, and Topo-78) and single and double
stranded DNA were analyzed by EMSA experiments. Topo-44 and Topo-78 formed stable
complexes with a 39mer double stranded DNA (Figure 2D), while no DNA binding was
observed for the Topo-31 fragment (data not shown). These observations indicate that
(HhH)2 domains are necessary for a stable protein-DNA complex and that as few as one and
half (HhH)2 domains are enough for formation of a stable protein-DNA complex. EMSA
with single stranded DNA showed that Topo-31 and Topo-44 cannot bind to single stranded
DNA, while Topo-78 can bind to single stranded DNA (data not shown).
Overall Structures
The topoisomerase domain of topoisomerase V is a helical-rich compact domain that has no
structural similarity to any other known protein. The only recognizable structural element is
a HTH that contains some of the active site residues. Not surprisingly, the topoisomerase
domain of the four structures (Topo-31, Topo-44 (Forms I, II, and III)) superimpose very
well on each other and also to that from the Topo-61 structure. In the Topo-31 structure, two
surface loops, residues 39-49 and 120-124, adopt a different conformation compared to the
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Topo-61 and Topo-44 structures. These two loops are not always visible in the Topo-44
structures, suggesting that they are mobile regions. The r.m.s.d. for the superposition of the
topoisomerase core domain of all the new structures on to the Topo-61 structure range from
0.2 Å to 0.7 Å if the two mobile surface loops are not included (Figure 3A). In general, the
topoisomerase domain remains unchanged and is identical in all structures. The (HhH)2
domains also remain largely unchanged, with r.m.s.d. for the superposition of only the
(HhH)2 domains from the three Topo-44 crystal forms and equivalent domains in the
Topo-61 structure ranging from 0.31 Å to 0.56 Å.
The five crystallographically independent structures of Topo-44 (Form I, Form II A and B
monomers, and Form III A and B monomers) were compared with each other and to the two
crystallographically independent Topo-61 monomers to understand the conformational
changes in the protein. The r.m.s.d. for the superposition of all the Topo-44 structures
(residues 3-375) on to the Topo-61 fragment or on each other vary between 0.9 Å and 2.7 Å,
with the majority above 1.5 Å, showing that in general the structures have slightly different
conformations. As mentioned above, the different domains behave as rigid or almost rigid
subunits and the only change in the structure is the relative orientation between the
topoisomerase and the (HhH)2 domains. The change in orientation of the domains starts at
the linker helix (residues 269-295), which acts as a hinge region, and follows into the
(HhH)2 domains. At the start of the linker helix, the structures superimpose very well for all
five Topo-44 and two Topo-61 structures. In the middle of the linker helix there is a kink
after which the linker helix from all the structures shows different orientations (Figure 3B).
The flexibility of the linker helix is also evident by the fact that the linker helix in the B
subunit of Form III crystals appears in two alternate conformations. The change in the
relative orientation of the (HhH)2 and topoisomerase domains (Figure 3C and 3D), suggests
that these domains can adopt different orientations and these movements might be necessary
for the DNA to access the active site.
The topoisomerase domain has a positively charged groove adjacent to the active site
The structure of the Topo-31 as well as the structures of the Topo-44 fragment reveals the
presence of a positively charged groove in the protein that encompasses the active site
region (shown later in Figure 6C). This charged groove had been observed before in the
structure of the Topo-61 fragment, although several (HhH)2 motifs partially obstruct it
(Taneja et al., 2006). The structure of the Topo-31 confirms the presence of the groove even
in the absence of the (HhH)2 motifs. The groove is long and can be deep in some areas. It
includes regions of the HTH motifs and extends all the way to the linker helix. All the
residues forming the active site pentad point towards the groove. The active site tyrosine,
Tyr226, is found near one of the ends of the groove, a region where it widens. The positively
charged character of the groove and its presence by the active site strongly suggest that it
may be involved in DNA binding.
Phosphate ions bind in the groove near the topoisomerase active site
An interesting observation stemming from the Form II and Form III Topo-44 structures is
the presence of phosphate ions near the positively charged DNA binding groove. All three
Topo-44 crystal forms were crystallized in the presence of phosphate-citrate buffer, but only
Form II and Form III structures showed phosphate ions bound to the protein, which were
assigned based on electron density consistent with a tetrahedral phosphate ion (Figure 4A).
Form II and Form III crystals include 1–1.2 M guanidium hydrochloride in the
crystallization solution. The high resolution Form III structure shows clear density for three
guanidium ions bound to the protein, two very well ordered and one with weak density. The
presence of guanidium hydrochloride in the crystals appears to trigger a conformational
change allowing the binding of phosphate ions to the protein. It is interesting to note that
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Form I crystals did not show any bound phosphate albeit its presence in the crystallization
condition. This could be due to the absence of guanidium hydrochloride to trigger the
binding of phosphate ions as observed in Form II and Form III structures. There are three
phosphate ions in the B subunit and none in the A subunit of the Form II Topo-44 structure.
Two of the phosphates are in the topoisomerase active site and one of them forms close
contacts with the putative active site residues in the topoisomerase domain (Figure 4B).
Form III crystal has seven phosphate ions, three in each subunit and one between both the
subunits. In the Form III structure, the phosphate ion near the active site Tyr226 is absent,
but it shows several new locations for phosphate ions, especially in the positively charged
groove containing the topoisomerase active site (Figure 5A). An overlay of the A and B
subunits of the Topo-44 Form III structure with the B subunit of Topo-44 Form II structure
shows eight unique phosphate ions (Figure 5A). It clearly shows that there are more
phosphate ions bound in the positively charged groove compared to other regions of the
protein.
Taking into account all structures, there are five unique phosphate ion binding sites in the
putative DNA binding groove and an additional one near its end and close to the start of the
linker helix. Several pairs of phosphates in the groove are separated by a distance of around
7 Å (Figure 5B), which would be consistent with the phosphate-phosphate distance in
adjacent nucleotides in a DNA double helix. One of the phosphates (P1) is found near the
active site tyrosine and is coordinated by Tyr226 and Arg131, two residues that have been
implicated in cleavage and religation of the DNA (Taneja et al., 2006), and by Glu215,
whose charge may be important for interactions with DNA (R.R. and A.M., unpublished
observations). The side chains of the tyrosine and the glutamate residues are in contact with
Arg144 and His200, the other putative active site residues, and these interactions may help
to orient them for the catalytic reaction. Adjacent to P1, there is a second phosphate (P2) at a
distance of 7.5 Å which is trapped between the topoisomerase domain and an HhH motif. P2
is coordinated by Arg131, an active site residue, in addition to Arg108 from the
topoisomerase domain and Arg293 and serines 322 and 324 from the second HhH motif
(Figure 6C). Three more phosphates are found in the groove (P3, P4, and P5) coordinated
mainly by positively charged residues, such as Arg37, Lys47, Arg108, Lys134, and Arg135
from the topoisomerase domain and also residues from the linker helix such as Tyr289 and
Arg293. In general, some of the side chains can contact more than one phosphate. The
distance between P3 and P4 and P4 and P5 is 6.8 Å and 6.5 Å respectively. A final
phosphate (P6) is located at the start of the linker helix and on the edge of the groove
(Figure 5A).
Discussion
Topoisomerase V is active at very high temperatures (122°C) and high salt concentrations.
DNA relaxation assays with various topoisomerase V fragments (Topo-44 and Topo-61)
show that a temperature above 60° C is required for optimal activity, although longer
fragments of topoisomerase V can relax DNA at lower temperatures (Taneja et al., 2007).
Topo-44 was first identified by limited proteolytic digestion of the full length topoisomerase
V protein (Belova et al., 2002) at 80°C. In contrast, Topo-61 is the shortest fragment
showing topoisomerase activity when the proteolytic reaction is performed at 37°C (Belova
et al., 2002). The N-terminal Topo-31 fragment, which contains neither HhH motifs nor the
linker helix, was identified as the smallest region spanning the topoisomerase domain from
the crystal structure of Topo-61 fragment (Taneja et al., 2006) and it was suggested that it
could represent the minimal domain capable of relaxing DNA. Relaxation experiments with
this minimal domain show that this is indeed the case, although the activity is not as robust
as with longer fragments. As expected, Topo-31 does not require magnesium for activity,
but magnesium enhances it, as is the case for type IB topoisomerases, which also uses a
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swiveling mechanism for DNA relaxation (Stewart et al., 1996). The optimal pH for activity
for the Topo-31 and the Topo-44 fragments is around 5. This pH dependence is not observed
for the Topo-78 fragment. The DNA relaxation by shorter fragments of topoisomerase V at
pH 5 could point to the involvement of some ionizable side chains in the relaxation activity.
It could also be simply due to the effects of various side chains on DNA binding. Further
experiments with different active site mutations in both longer and shorter fragments of
topoisomerase V will be required to probe the pH dependence of the relaxation reaction by
shorter topoisomerase V fragments.
Gel shift experiments show that Topo-44 and also longer fragments (Topo-78) can bind
double stranded DNA. Surprisingly, Topo-31 does not show DNA binding activity in these
assays even though it is still capable of relaxing DNA. It appears that the presence of the
(HhH)2 domains stabilizes the DNA/protein complex. One possibility is that the (HhH)2
domains could play a similar role to the cap domain present in type IB enzymes, which helps
to encircle the DNA during the swiveling reaction (Redinbo et al., 1998). In addition, both
short fragments of topoisomerase V do not bind single stranded DNA, whereas Topo-78 can
form a stable complex with single stranded DNA (data not shown). (HhH)2 domains binding
to single stranded DNA has been observed before. For instance, the N-terminal 8 kDa of
mammalian polymerase β, which contains a single HhH motif, binds to single stranded DNA
through both helices (Kumar et al., 1990; Liu et al., 1994). The exact mode of single
stranded DNA binding by Topo-78 or the possible role in relaxation or repair activities is not
yet clear.
The structure of Topo-61 showed that the topoisomerase active site of topoisomerase V is
buried by one of the (HhH)2 domains suggesting that conformational changes are essential
for the protein to bind DNA. The present structures of Topo-44 reinforce this observation
and show that the (HhH)2 domains can change their position relative to the topoisomerase
domain and that this change is mediated by the movement of the linker helix. The (HhH)2
domains act as rigid individual units, as evidenced by the fact that in different structures
they show the same structure and relative orientation of the two HhH motifs. The
topoisomerase domain also appears to be rigid showing the same structure even in the total
absence of the rest of the protein. The linker helix (residues 269-295), which is a long bent
helix, serves as a hinge for the movement of the (HhH)2 domains away from the rigid
topoisomerase domain, possibly by responding to interactions with double stranded DNA.
This movement has to be quite large. The Topo-44 structures in the absence of DNA capture
the regions that move, but do not show the full extent of the movement or indicate the way
the HhH motifs interact with DNA.
As mentioned before, topoisomerase V binds double stranded DNA and has a groove wide
enough to accommodate double strand DNA (Figure 6C). The presence of an HTH domain
normally associated with DNA binding, the positively charged nature of it, and several
phosphates bound along it suggest that this groove could be involved in DNA binding. In
addition, the active site is found in this groove and some residues form part of the HTH
domain. Previously, DNA was modeled bound to the topoisomerase domain (Taneja et al.,
2006) based on the structures of HTH domains in complex with DNA but there was no
evidence to support it. Using the phosphates present in the groove in the current structures, it
is possible to refine this model. A superposition of the B subunit of Form II and the A and B
subunits of Form III Topo-44 structures shows five different phosphate ions in the positively
charged groove which are separated by a distance of around 7 Å, consistent with the
distance of consecutive phosphates in B DNA of ~6.4 Å. A sixth phosphate ion is found
outside the groove near the linker helix. A double stranded DNA molecule was modeled into
the groove based on the positions of the phosphate ions (Figure 6). Only five out of the six
phosphates could be placed on the DNA molecule, as one of them was inconsistent with a
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double stranded molecule. Phosphate ions P3, P4, and P5 would correspond to three
adjacent phosphates in one DNA strand, while P1, located near the active site, would belong
to the opposite strand. A final phosphate (P6) is away from the groove and near the linker
helix (Figure 6A, 6B). The fit to the four inner phosphates is excellent and the DNA can be
accommodated in the groove of the Topo-31 structure without the need for any major
rearrangements of the protein backbone. The fifth phosphate (P6) does not fit as well and a
better fit would require movement of either the protein or the DNA, but the change would be
relatively modest. Several side chains would need to move, but these changes would also be
minor. The major change needed to accommodate the DNA in the structures with the
(HhH)2 domains present is the movement of the (HhH)2 domains away from the
topoisomerase domain (Figure 6B). The movement of (HhH)2 domains should be feasible as
is evident from the Topo-44 structures showing different orientations of the (HhH)2
domains. The location of the (HhH)2 domains after DNA binding is not evident, but one
possibility is that they would help enclose the DNA to form a clamp around it, similar to the
arrangement in type IB enzymes.
In the model of the topoisomerase domain in complex with DNA, the active site residues are
in close contact with the backbone of DNA. The catalytic Tyr226 is pointing towards the
phosphate of the DNA backbone; Arg131 and Arg144 are positioned to stabilize the protein-
DNA covalent complex. Surprisingly Glu215 also appears to interact directly with the DNA
phosphate backbone. The other active site residues like His200 and Lys 218 are also near the
DNA. The active site is located near the end of the groove, where it widens. At this end, the
DNA fits loosely in the groove, which is spacious to accommodate the movement of the
strands. The ‘constrained swiveling’ mechanism employed by type IB and IC enzymes
necessitates rotation of one strand about the other after forming the covalent protein-DNA
intermediate. The position of the active site at the wider end of the putative DNA binding
groove would facilitate the rotation of the DNA strand at this end, while holding the rest of
the DNA in place through extensive interactions along the groove.
Even though type IB and IC enzymes have a similar overall mechanism of action, the
structures of fragments of topoisomerase V suggest many differences. Type IB enzymes
have two domains which come together to form a C-shaped clamp around the DNA (Perry et
al., 2006; Redinbo et al., 1998; Stewart et al., 1998) The protein has an open stage where
these domains are separate and this helps in the entry and release of the DNA from the
protein active site. A wide DNA binding cavity is not observed in the topoisomerase V
structures. Instead, the structures show a positively charged groove which is always present
in the protein and does not require domain rearrangements to form. DNA can access this
groove after a conformational change involving the movement of the (HhH)2 domains
exposing the active site. The (HhH)2 domains could help enclose DNA during the swiveling
of the DNA, forming a similar enclosure to the one observed for type IB enzymes. It is not
known whether all HhH motifs contact DNA simultaneously, but this appears unlikely
without a major rearrangement of the motifs. It is likely that only some of the HhH motifs
contact DNA at any given time or that some of the motifs do not have the capacity to bind
DNA. Finally, similar to type IB enzymes (Cheng and Shuman, 1998), the putative domain
enclosing the DNA is dispensable for activity, although it enhances the relaxation activity
markedly. Thus, it is likely that type IB and IC enzymes have several overall similarities in
the way that they interact with DNA, but the atomic details are markedly different.
There are still many details of the atomic mechanism of type IC topoisomerases that need to
be understood. The present functional and structural studies provide new information about
topoisomerase V including the observations that the Topo-31 is the minimal fragment
capable of DNA relaxation, the (HhH)2 domains enhance binding of the protein to DNA, the
changes in relative orientation of the domains is mediated by the linker helix, and several
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phosphate ions bind in a positively charged groove. Furthermore, the position of the
phosphate ions in the groove helped in the placement of DNA in the topoisomerase domain
and this provides an initial model of how topoisomerase V interacts with DNA. Thus the
present study helps to establish the role of different domains more clearly, to illustrate a
mechanism to drive the conformational changes needed for activity, and to suggest a
possible manner of binding DNA. Additional work on structures of protein/DNA complexes
and intermediates in the swiveling reaction are needed to understand the way this new type
of topoisomerases interacts with DNA to perform a complex reaction.
Experimental Procedures
Protein purification
The N-terminal 31 kDa (Topo-31: residues 1-269), and 44 kDa (Topo-44: residues 1 to 380)
fragments of topoisomerase V protein were cloned into the pET15b plasmid and
transformed into Escherichia coli BL21 Rosetta (DE3) cells. The N-terminal 78 kDa
(Topo-78: residues 1 to 685) fragment of topoisomerase V protein was cloned into the
pET14b plasmid (Belova et al., 2002) and transformed into Escherichia coli BL21(DE3)
cells. For protein production, cells were grown at 37° C in LB medium containing 100 μg/ml
ampicillin and 100 μg/ml chloramphenicol for Rosetta cells and LB medium with 100 μg/ml
ampicillin for BL21(DE3) cells to an optical density (OD600) of 0.6. The cells were then
cooled down on ice, followed by the addition of isopropyl β-D-1-thiogalactopyranoside
(IPTG) to a final 0.5 mM concentration, and grown overnight at 16° C. Cells were harvested
and resuspended in 50 mM Tris pH 8, 500 mM NaCl, 0.5 mM EDTA, 1 mM DTT, flash
frozen in liquid nitrogen and stored at −80° C. After thawing the pellet, pepstatin (1μg/ml),
benzamidine (1mM), PMSF (1mM), and Brij 58 (0.1%) were added to the cells and the
protein was purified as described earlier (Taneja et al., 2006) The protein was further
purified by anion exchange and gel filtration chromatography. Pure protein was
concentrated and stored in 50 mM Tris pH 8, 250 mM NaCl, and 1 mM DTT. The seleno-
methionine substituted Topo-44 was prepared from cells grown in a minimal medium
supplemented with nutrients and salts (Doublie, 1997); protein purification followed the
same procedure as for the native protein except that 5mM DTT was used in all the
purification steps and for storage.
Relaxation assays
Relaxation assays with the different topoisomerase V fragments were carried out at pH
values ranging from 4 to 10. The pH of the buffers was adjusted at 65 °C to account for the
change in pH at higher temperature. The different buffers used were: sodium acetate for pH
4 and 5, MES for pH 6, HEPES for pH 7, TRIS for pH 8, CHES for pH 9, and CAPS for pH
10. Topoisomerase activity assays were performed by incubating varying amounts of protein
(Topo-31, Topo-44 or Topo-78) with 0.2 μg negatively supercoiled pUC19 DNA in 50 mM
of the required buffer, 30 mM NaCl, 0.2 mM or 5 mM EDTA or 1 mM MgCl2. The
reactions were carried out at 65 °C for 15 min and terminated by cooling and addition of
SDS to a final 1% concentration. The products were resolved on a 1% agarose gel and
visualized by ethidium bromide staining.
Electrophoretic Mobility Shift Assay
For Electrophoretic Mobility Shift Assay (EMSA), 4 μM of a 39mer double stranded DNA
oligonucleotide (5′ GCGACGCGAGGCTGGATGGCCTTCCCCATTATGATTCTT3′) was
incubated with different concentrations of topoisomerase V fragments in 50 mM sodium
acetate pH 5, 30 mM NaCl, 1 mM MgCl2 at 65 °C for 30 minutes. Glycerol was added to
the reaction mixture to a final concentration of 8% and the products were separated on a 4 %
acrylamide native gel. The gel was stained with ethidium bromide to detect the DNA. When
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a stable protein-DNA complex was formed, there was an upward shift in the band indicating
a higher molecular weight complex.
Crystallization
Topo-31 crystals were grown using the sitting drop vapor diffusion method equilibrated
against, 23% PEG 6000, 0.1 M Na citrate pH 5.5, at 22°C. For data collection, the Topo-31
crystals were cryo-protected by adding glycerol to the mother liquor to a final 20%
concentration. Topo-44 was crystallized by the hanging drop vapor diffusion method under
three different crystallization conditions (Forms I, II, and III). Crystal Form I grew under 0.1
M phosphate citrate pH 5, 0.2 M NaCl, 15% PEG 3350 and 8% dioxane. The crystals were
cryo-protected by increasing the PEG concentration to 30%. Form II crystals grew under 0.1
M phosphate citrate pH 5, 0.2 M NaCl, 16% PEG 8000 and 1M guanidium hydrochloride.
For cryo-protection, they were transferred to a solution with 1.5X reservoir solution and
20% 2,3 butanediol or 20% DMSO for 10 seconds and immediately flash frozen under
liquid nitrogen. Form III crystals grew under 0.1 M phosphate citrate pH 5.5, 0.15 M sodium
sulfate, 0.01 M MgCl2, 1 M guanidium hydrochloride, and 28 % PEG 3350. The crystals
were grown at 30°C and were cryo-protected by increasing the PEG concentration to 40%.
Further details of crystallization are presented in the Supplementary Information.
Data collection and structure determination
Diffraction data were collected at the Dupont Northwestern Dow and Life Science
Collaborative Access Team stations (DND and LS CAT) at the Advanced Photon Source in
Argonne National Laboratory. Data collection and refinement statistics are shown in Table I.
All data were processed and integrated using XDS (Kabsch, 1993) and scaled with SCALA
(Collaborative-Computational-Project-4, 1994). Data on the Topo-31 crystals were collected
to 2.4 Å resolution. The structure was solved by Molecular Replacement (McCoy et al.,
2007) using the topoisomerase domain from the Topo-61 structure (residues 1-266) (Taneja
et al., 2006) as the search model. It was refined with refmac5 (Murshudov et al., 1997) and
Phenix (Afonine et al., 2005) to a final Rwork of 20.0 % and Rfree of 24.8 %. Topo-44 Form I
crystals diffract to 1.8 Å. The structure of Form I crystals was solved by Molecular
Replacement (McCoy et al., 2007) using the topoisomerase domain from the Topo-61
structure as the search model. Model rebuilding was performed using coot (Emsley and
Cowtan, 2004), and refinement using refmac5 (Murshudov et al., 1997). The final Rwork and
Rfree are 17.5 % and 22.0 % respectively.
For Topo-44 Form II and Form III crystals, seleno-methionine derivatized crystals were
used for single-wavelength anomalous dispersion (SAD) phasing. AutoSharp (Vonrhein et
al., 2007) was used for locating the selenium atoms; model building was done using coot
(Emsley and Cowtan, 2004), and refinement was carried out using refmac5 (Murshudov et
al., 1997) Three phosphate ions were noticed in the Form II structure; two of which present
in the topoisomerase active site and are separated by a distance of ~7.5 Å. The structure was
refined to a final Rwork of 24.1 % and Rfree of 28.9 %. Topo-44 Form III crystals diffracted
to 1.4 Å. The final Rwork and Rfree are 16.5 % and 18.4%, respectively. An interesting
observation is the presence of both phosphate and guanidium ions in the Form III Topo-44
structure. The linker helix and part of the first HhH motif of the B monomer show alternate
conformations and were built as two separate chains with occupancy of 0.5 each. Further
details on data collection and structure determination are given in the Supplementary
Information.
All figures were made with Pymol (DeLano, 2002) and the electrostatic surfaces were
calculated with APBS (Baker et al., 2001).
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Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We acknowledge staff and instrumentation support from the Keck Biophysics Facility and the Center for Structural
Biology at Northwestern University, and DND and LS-CAT at the Advanced Photon Source (APS) at Argonne
National Laboratory. Support from the R.H. Lurie Comprehensive Cancer Center of Northwestern University to the
Structural Biology Facility is also acknowledged. DND-CAT is supported by Dupont, DOW and the NSF. LS-CAT
was supported by the Michigan Economic Development Corporation and the Michigan Technology Tri-Corridor.
Use of the APS is supported by the Department of Energy (DOE). Research was supported by NIH grant GM51350
(to AM).
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Figure 1. Organization of topoisomerase V
Topoisomerase V is a multi-domain protein consisting of 24 helix-hairpin-helix (HhH) DNA
binding motifs arranged as 12 (HhH)2 domains following the N-terminal topoisomerase
domain. A) Schematic diagram of various topoisomerase V fragments. The topoisomerase
domain is shown in red, the (HhH)2 domains are shown in alternating colors of cyan and
yellow. The (HhH)2 domains with repair activity are shown in green. All fragments shown
have topoisomerase activity, but only the full length protein and the Topo78 fragment have
repair activity. B) Crystal structure of Topo-61 fragment (Taneja et al., 2006). The coloring
scheme is the same as in Figure 1A, except that the linker helix is shown in grey.
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Figure 2. DNA relaxation activity and EMSA for Topo-31, Topo-44 and Topo-78 fragments of
topoisomerase V
A) pH profile of the DNA relaxation activity of Topo-78 and Topo-44 fragments. 0.2 μg of
pUC19 DNA were incubated with 0.1 μg of Topo-78 or 1.5 μg of Topo-44 proteins at 65°C
for 15 minutes for the relaxation reaction. The reaction buffer contained 50 mM of the
appropriate buffer, 30 mM NaCl and 0.2 mM EDTA. Topo-78 relaxes DNA at a wider pH
range (5 to 9) than Topo-44, which relaxes DNA efficiently only at pH 5. DNA relaxation
activity of Topo-31 (B) and Topo-44 (C) fragments in the absence and presence of MgCl2.
0.2 μg of pUC19 DNA were incubated with 9 μg of Topo-31 or 1.5 μg of Topo-44 proteins
at 65°C for 15 minutes for the relaxation reaction. The reaction buffer contained 50 mM of
the appropriate buffer, 30 mM NaCl and 5 mM EDTA or 1 mM MgCl2. Both Topo-31 and
Topo-44 fragments can relax DNA in the absence of MgCl2, but MgCl2 enhances the DNA
relaxation activity of the topoisomerase V fragments. The black triangle in panels A, B and
C represents increasing pH from 4 to 10 by one pH unit. D) EMSA of Topo-44 and Topo-78
fragments with a 39mer double stranded DNA. Both Topo-44 and Topo-78 form stable
complexes with DNA, although Topo-78 seems to saturate DNA binding while Topo-44
does not. In addition, Topo-44 shows some cleavage of the DNA (bottom free DNA band),
while the cleavage is not apparent in Topo-78. The numbers at the bottom represent the
molar ratio of protein to DNA.
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Figure 3. Structure of Topo-44 fragments
A) Overlay of Form I (green), Form II (B subunit: blue), and Form III (B subunit: magenta)
structures onto the Topo-61 structure (B subunit: orange). The topoisomerase domains
superimpose very well for all the structures, while the linker helix and (HhH)2 domains
show differences in orientation. B) Overlay of the linker helices of Form I, II, and III
structures with that of Topo-61. The color scheme is same for all the figures unless
mentioned otherwise. Note that the linker helices have the same orientation at the start and
they change as they move further down the helix. C) Superposition of Form I, II, and III
Topo-44 structures with that of Topo-61. Only the (HhH)2 domains are colored while the
remaining parts are shown in gray for clarity. The active site residues are shown as orange
sticks. Note that the (HhH)2 domains adopt different orientations in all the structures. D)
Orientation of the (HhH)2 domains of Form I, II and Topo-61 structures. In Form I and II
structures, the (HhH)2 domains are moved away from the topoisomerase domain. For clarity,
the (HhH)2 domains of Form III are not shown. In panels C and D, the topoisomerase
domains were superposed to emphasize the different orientation of the other domains.
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Figure 4. Phosphate ions present near the active site of the Topo-44 structure
A) Stereo view of a Form III difference electron density map calculated with a model not
including the phosphates. The electron density is contoured at 3.7σ and shows the
tetrahedral shape of the phosphate ions. The active site residues are shown in stick. B)
Stereo view of the interaction of the phosphate ions with the putative active site residues.
The B subunit of Form II structure was superimposed onto the B subunit of Form III
structure and the phosphates ions from both structures are shown together with the Form II
B subunit protein backbone. The interactions made by the phosphate ion with the active site
residues and the corresponding distances in Å are represented as black dotted lines.
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Figure 5. Representation of the unique phosphate ions from Form II and Form III Topo-44
structures
A) An overlay of the A (magenta) and B (brown) subunits of the Form III structure and B
(blue) subunit of Form II Topo-44 structures. The positions of eight unique phosphate ions
(orange spheres) are shown. Note that most phosphate ions are found along the DNA
binding groove of the topoisomerase domain. B) The phosphate ions in the DNA binding
groove are separated by distances of around 7 Å. The protein backbone is that of the B
subunit of Form III structure. The active site residues are represented as sticks and distances
in Å between adjacent phosphate ions are shown as black dotted lines.
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Figure 6. Model showing DNA bound to the topoisomerase domain
A) Model of a 17-mer double stranded DNA bound to the Topo-31 structure (teal). The
DNA is represented as green sticks, where as phosphate ions are represented as orange
sticks. DNA binds along the DNA binding groove and five of the eight phosphate ions noted
in the Topo-44 structures coincide with the DNA backbone. B) Model of Topo-44 (Form II,
B subunit: blue) binding to 17-mer double stranded DNA. Note that the linker helix and the
(HhH)2 domains interfere with DNA binding to the topoisomerase domain and are likely to
move away to allow binding. C) Electrostatic surface representation of the Topo-31
structure. The positively charged DNA binding groove is clearly visible and the phosphate
ions are bound in this groove. The orientation corresponds to a 90° rotation of the one shown
in Figure 6A in the direction of the arrow. Note that the DNA binding groove goes from one
end of the molecule to the other and it is narrower at one end (start of the linker helix) and
wider at the other end. The putative active site residues (green sticks) are located at the
wider end of the groove. Other residues lining the groove and interacting with the phosphate
ions are shown as cyan sticks. D) Electrostatic surface representation of Topo-31 with
phosphate ions (orange) and DNA (green). Three phosphate ions (P3, P4, and P5) coincide
with the phosphates of one of the DNA strands, where as P1 coincides with a phosphate of
the opposite DNA strand. The model shows that the DNA binding groove of topoisomerase
V is wide enough to bind DNA and that the movement of linker helix and (HhH)2 domains
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are required to accommodate the DNA. The electrostatic potential was calculated with a
dielectric constant of 80 for solvent and 2 for protein. The surface is colored with a blue to
red gradient from +10 to −10 KbT/ec.
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Table 1
Data collection and refinement statistics
Topo-31
Topo-44 Form I
Topo-44 Form II
Topo-44 Form III
Data Collection
Space group
C2221
C121
P41212
P212121
Cell dimensions
a=106.7 Å, b=119.4
Å, c=63.7 Å
a=104.2 Å, b=47.7 Å,
c=81.2 Å (β=112.48)
a=b=70.1 Å, c=349.6 Å
a=63.6 Å, b=80.1 Å,
c=137.2 Å
Resolution (Å)a
79.56 – 2.4 (2.53 –
2.4)
75.05 – 1.82 (1.91 –
1.82)
29.5- 2.6 (2.72-2.6)
28.9-1.4 (1.46-1.4)
Number of observed
reflections
78,729 (11,538
134,411 (13,220)
227,408 (19,917)
1,157,917 (126,319)
Number of unique reflections
16,259 (2,346)
32,998 (4,301)
28,151 (3,331)
136,662 (15,986)
Completeness (%)
99.8 (99.8)
98.3 (88.6)
99.9 (100.0)
98.8 (95.5)
Multiplicity
4.8 (4.9)
4.1 (3.1)
8.1 (6.0)
8.5 (7.9)
Rmerge (%)b
4.7 (71.1)
4.0 (16.3)
7.4 (52.2)
4.5 (37.9)
Rmeas (%)c
5.3 (79.6)
4.6 (19.4)
7.9 (57.2)
4.8 (40.5)
≪I>/σ(<I>)>d
20.5 (2.5)
23.0 (6.8)
19 (3.2)
27.5 (5.3)
Refinement
Resolution (Å)
79.56 - 2.4 (2.46 -
2.4)
28.06 -1.82 (1.87 –
1.82)
29.14 – 2.6 (2.67 – 2.6)
28.9 - 1.4 (1.44 - 1.4)
Number of reflections
working/test
15,419/821
31,317/1,673
26,710/1,438
129,802/6,859
Rwork (%)e
20.0(24.3)
17.5 (17.9)
24.1(36.6)
16.5 (19.3)
Rfree(%)f
24.8 (31.1)
22.0 (24.8)
28.9 (45.1)
18.4 (22.1)
Protein residues/atomsg
269/2,203
376/3212
727/5,970
738/7,511
Atoms in alternate
conformations
0
258 (20 protein
residues)
8 (1 protein residue)
2846 (157 protein
residues)
Water molecules
29
238
30
573
Other atoms
-
-
3 PO4
7 PO4, 3 Gmh, 3 Mg++, 2
Cl−
B-factor (Å2)
Protein atoms (chain)
68.4
22.8
A:53.8; B:58.2
A:13.4; B:14.9
Water molecules
59.1
29.3
40.0
23.7
r.m.s. deviations
bond lengths (Å)
0.015
0.006
0.01
0.009
bond angles (°)
1.42
0.920
1.2
1.2
Ramachandran ploti
Favored regions (%)
94.3
98.9
96.2
98.5
Outliers (%)
0.0
0.0
0.3
0
aNumbers in parenthesis correspond to highest resolution shell.
bRmerge= Σ|I − <I>|/ΣI, where I is the observed intensity and <I> the average intensity obtained from multiple measurements.
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cRmeas as described in Diederichs and Karplus (Diederichs and Karplus, 1997).
d≪I>/σ(<I>)> = Mean Ih over the standard deviation of the mean Ih averaged over all reflections in a resolution shell.
eRwork= Σ ||Fo| − |Fc||/Σ|Fo|, where |Fo| is the observed structure factor amplitude and |Fc| the calculated structure factor amplitude.
fRfree: Rfactor based on 5% of the data excluded from refinement.
gTotal number of protein atoms, including those in alternate conformations.
hGm: guanidinum ion.
iAs reported by Molprobity (Davis et al., 2004).
Structure. Author manuscript; available in PMC 2011 July 14.
|
3M71
|
Crystal Structure of Plant SLAC1 homolog TehA
|
Homolog Structure of the SLAC1 Anion Channel for Closing
Stomata in Leaves
Yuhang Chen1,6, Lei Hu2, Marco Punta6,7, Renato Bruni6, Brandan Hillerich6, Brian Kloss6,
Burkhard Rost1,6,1, James Love6, Steven A. Siegelbaum2,3,5, and Wayne A.
Hendrickson1,4,5,6
1Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY
10032, USA
2Department of Neuroscience, Columbia University, New York, NY 10032, USA
3Department of Pharmacology, Columbia University, New York, NY 10032, USA
4Department of Physiology and Cellular Biophysics, Columbia University, New York, NY 10032,
USA
5Howard Hughes Medical Institute, Columbia University, New York, NY 10032, USA
6NYCOMPS, New York Structural Biology Center 89 Convent Avenue, New York, NY 10027 USA
7Department of Computer Science and Institute for Advanced Study Technical University of
Munich D-85748 Munich, Germany
Summary
The plant SLAC1 anion channel controls turgor pressure in the aperture-defining guard cells of
plant stomata, thereby regulating exchange of water vapor and photosynthetic gases in response to
environmental signals such as drought or high levels of carbon dioxide. We determined the crystal
structure of a bacterial homolog of SLAC1 at 1.20Å resolution, and we have used structure-
inspired mutagenesis to analyze the conductance properties of SLAC1 channels. SLAC1 is a
symmetric trimer composed from quasi-symmetric subunits, each having ten transmembrane
helices arranged from helical hairpin pairs to form a central five-helix transmembrane pore that is
gated by an extremely conserved phenylalanine residue. Conformational features suggest a
mechanism for control of gating by kinase activation, and electrostatic features of the pore coupled
with electrophysiological characteristics suggest that selectivity among different anions is largely
a function of the energetic cost of ion dehydration.
Users may view, print, copy, download and text and data- mine the content in such documents, for the purposes of academic research,
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Correspondence and request for materials should be addressed to WAH (wayne@convex.hhmi.columbia.edu)..
Supplementary Information is linked to the online version of the paper at www.nature.com/nature.
Author Contributions YC, LH, MP, RB, BH, BK and JL performed experiments; YC, LH, BR, SAS, and WAH analyzed data; YC,
LH, SAS, and WAH prepared the manuscript.
Author Information Atomic coordinates and diffraction data are deposited in the Protein Data Bank with accession codes 3M71,
3M73, 3M74, 3M75, 3M76, and 3M7L. See Table S4 for identifications. Reprints and permissions information is available at
www.Nature.com/reprints.
HHS Public Access
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Published in final edited form as:
Nature. 2010 October 28; 467(7319): 1074–1080. doi:10.1038/nature09487.
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Stomatal pores in the leaves of plants permit the influx of atmospheric carbon dioxide in
exchange for transpirational evaporation of water1,2. A pair of kidney-shaped guard cells
define each pore aperture, and turgor pressure variation in these cells determines the degree
of stomatal pore openness. Depending on diverse environmental factors, the stomata close to
prevent H2O losses and open to admit CO2 for photosynthesis. Environmental stimuli that
lead to stomatal closure include darkness, high CO2 levels, ozone, low air humidity and
drought. The plant hormone abscisic acid (ABA) is critical for signal transduction from
these stimuli. Mutational screens in Arabidopsis thaliana for CO2 and ozone sensitivity
identified a protein with ten predicted transmembrane (TM) helices, now called slow anion
channel 1 (SLAC1), as having a central role in the control of stomatal closure3-5. Recent
studies proved that SLAC1 is indeed an anion channel6-7, with characteristics like those of
slow anion channels found in guard cells8, and that it is activated by phosphorylation from
the OST1 kinase9. OST1 activity is negatively regulated by the ABI1 phosphatase10,11,
which is in turn inhibited by the stomatal ABA receptors PYR/RCAR12,13 when in the
ternary hormone-receptor-phosphatase complex14-18. Thereby, ABA stimulates SLAC1
channel activity. Resulting Cl− efflux through SLAC1 causes membrane depolarization,
which activates outward-rectifying K+ channels, leading to KCl and water efflux to further
reduce turgor and cause stomatal closure.
SLAC1 expression in Arabidopsis is confined to the guard cells of leaves, but other
Arabidopsis tissues do have SLAC1 homologs3, named SLAH1-SLAH4. The identifying
mutations slac1-14 and slac1-23 are, respectively, in predicted transmembrane segments 9
(S456F) and 1 (G194D) of a protein that includes substantial N- and C-terminal extensions
outside a 10-helix transmembrane (TM) domain. SLAH1, which is absent from leaves and
lacks the terminal extensions of SLAC1, fully complements the mutant phenotype in slac1-2
guard cell protoplasts3. SLAC1 and homologs are also present in other plant genomes,
including nine in rice (Oryza sativa) and five in grape vines (Vitis vinifera). SLAC1
relatives, some quite remote, also occur in bacteria, archaea and fungi. Known prokaryotic
homologs contain only the predicted transmembrane domain of SLAC1, but some fungal
homologs do have N- and C-terminal extensions. One homolog, Mae1 from the yeast
Schizosaccharomyces pombe, functions as a malate uptake transporter19; another, Ssu1 from
Saccharomyces cerevisiae and other fungi including Aspergillus fumigatus, is characterized
as a sulfite efflux pump20,21; and TehA from Escherichia coli is identified as a tellurite
resistance protein by virtue of its association in the tehA/tehB operon22,23. Despite a lack of
further biochemical characterization, many homologs are annotated as tellurite resistance/
dicarboxylate transporter (TDT) proteins.
We have undertaken structural and functional characterizations of the SLAC1 anion
channel. We first solved an atomic-resolution crystal structure of the TehA homolog from
Haemophilus influenzae, and we then developed a homology model for Arabidopsis SLAC1.
This model allowed us to conduct mutagenesis for functional testing of structure-inspired
hypotheses on gating and selectivity. We expressed Haemophilus TehA and Arabidopsis
SLAC1 in Xenopus oocytes to characterize channel properties of these proteins and mutant
variants. We also determined crystal structures for several mutant variants, including the
homolog of slac1-2.
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Structure of SLAC1 bacterial homolog TehA
We performed a bioinformatic analysis of SLAC1-related proteins, first clustering nearly
900 non-redundant sequences into a superfamily at the PSI-BLAST level E≤10−3, then into
three distinct families at an initial threshold of E≤10−30, and finally into subfamilies at a
typical initial threshold of E≤10−55. Since previous annotation is not well founded in
experiment and SLAC1 is now the best characterized member, we adopt a nomenclature
defining a SLAC superfamily divided into families identified as SF1-SF3 and subfamilies
SF1A, SF1B etc. Family SF1 comprises the plant SLAC proteins and close bacterial
homologs, family SF2 comprises a distinct set of bacterial proteins often annotated as
exfoliative toxins, and family SF3 comprises the fungal Mae1 and Ssu1 proteins and their
archaeal or bacterial homologs, respectively. SLAC family SF1 has three large subfamilies:
the plant SLAC and SLAH proteins are in subfamily SF1A, closest bacterial homologs are
in SF1B, and the TehA homologs are in SF1C (Fig. 1a). The other families also divide into
subfamilies as detailed in Table S1, and family SF1 is divided into sub-subfamilies (Table
S2). Two pertinent SF1 sequences are aligned in Fig. 1b.
We used a structural genomics approach to obtain structural information, testing expression
and purification for 43 bacterial and archaeal likely homologs, assaying for detergent choice
and stability on 8 of these, finding two with appropriate profiles by size exclusion
chromatography, and obtaining suitable crystals for one. This protein, TehA from H.
influenzae (HiTehA), was found to be trimeric both by size-exclusion multi-angle light
scattering (SEC-MALS) measurements and by chemical cross-linking. When solubilized in
β-octylglucoside, HiTehA crystallized in space group R3 with a=b=96.01Å and c=136.27Å.
Each asymmetric unit contains one subunit and 65% solvent. The structure was solved by
selenomethionyl (SeMet) SAD phasing, ultimately at 1.50Å resolution (Table S3, Fig. S1),
and then refined at 1.20Å resolution (Fig. 2a) to R/Rfree values of 14.1%/15.7% for a model
that includes ordered residues 6-313, 213 water molecules and four detergent molecules
(Table S4).
The crystal structure has TehA trimers aligned with three-fold axes of the lattice (Fig. 2b).
Subunits are tightly associated, burying 8947 Å2 of total surface area within trimer
interfaces. The electrostatic potential surface is largely negative on the extracellular surface
(Fig. 2c) and largely positive on the cytoplasmic surface (Fig. 2d). The membrane
orientation is specified experimentally from GFP tagging of E. coli TehA24. Each TehA
protomer has ten TM helices, as predicted; however, the fold is novel. Tandemly repeated
helical hairpins are arranged with quasi-five-fold symmetry (Figs. 2e,S2). Extracellular
inter-helix loops are short (2-5 residues) whereas intracellular inter-helix connections are
longer, including a nine-residue helix H2,3 between TM2 and TM3 (Fig. 2f). An inner pentad
of outwardly directed, TModd, helices creates an apparent pore through each protomer
perpendicular to the putative membrane plane. TMeven helices from the five hairpins
surround the inner pore and make an outer layer.
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Homology model for plant SLAC1
Arabidopsis SLAC1 (AtSLAC1) is substantially similar to bacterial homologs, notably
HiTehA (Fig. 1b). All HiTehA TM helices are fully aligned to predicted SLAC1 TM
helices, but there are short inter-helical gaps (1-5 residues) in all five extracellular loops and
in two of the intracellular loops. The TM domain of AtSLAC1 (residues 188-504) aligns to
HiTehA with 19% sequence identity and with a PSI-BLAST E-value of 3×10−22. For
comparison, and in keeping with the family tree (Fig. 1a), the TM domain of AtSLAC1
shares sequence identities of 76% with rice SLAC1, 41% with Arabidopsis SLAH1, 25%
with an SL1B homolog from Halorhodospira halophila, 11% with S. cerevisiae Ssu1, and
9% with S. pombe Mae1. A conceptual model with the AtSLAC1 sequence transposed onto
the HiTehA helices sufficed to guide most of our mutational tests, but a detailed AtSLAC1
homology model helped to refine our ideas. Surface variability and electrostatic potential are
plotted onto the surface of this model (Fig. 2g,2h).
The most remarkable feature of the TehA structure and corresponding SLAC1 model is the
central pore through each protomer. As for acetylcholine receptors25, the SLAC1 pore is
formed by five helices; but the SLAC1 helices come from one protein molecule rather than
five. The SLAC1 pore has a relatively uniform diameter of approximately 5Å across nearly
five helical turns (Fig. S3), except for a pronounced constriction in the middle of the
membrane (Fig. 3a) where the pore is occluded by the side chain of Phe 450 (Phe 262 in
HiTehA). This residue is the only absolutely conserved amino acid residue of the SLAC1
family. The pore is lined with highly conserved (86% identity among five SLAC1 orthologs;
32% identity between AtSLAC1 and HiTehA) and generally hydrophobic residues (Figs. 1b,
3b,3c,S4). Despite this hydrophobicity, the electrostatic potential on the pore surface is
polarized (Fig. 3a), presumably due to an invaginated shape adjacent to charged residues
outside the membrane. The generally electropositive character of the cytoplasmic surface
likely contributes to anion efflux.
Kinks in the pore helices contribute to formation of a relatively constant pore diameter
across the membrane. Four of the five HiTehA inner helices have centrally located proline
residues, which necessarily generate kinks, and TM9 is kinked at a backbone-coordinated
water molecule (Fig. 3d). Proline replaces Gly263 in all SF1A and SF1B relatives, including
Pro451 AtSLAC1. This water-displacing change is isostructural26. Centrally located prolines
also prevail in TM3, TM5 and TM7 across the SF1 proteins. The outer helices are longer and
straighter, but more inclined. The only outer-helix proline kink of HiTehA is in TM6 at the
trimer three-fold axis.
Two of the gene-identifying mutations in Arabadopsis SLAC1 are selected point mutations,
others are disruptive transfer DNA (T-DNA) insertions4. In the AtSLAC1 homology model,
the slac1-2 (G194D) mutation3 points into the pore from TM1 and can be accommodated
structurally whereas the slac1-1 (S456F) mutation4 points away from the pore six residues
after pore-blocking Phe450 on TM9 and would be expected to disruptive. Ser456 interacts
with outer helix TM10 in the homology model, and the phenyl bulk from S456F would not
fit. Position 456 has alanine in HiTehA, 66% of all 204 SF1 homologs and another 27%
have threonine or serine (as in all SLAC1 channels); phenylalanine never occurs among all
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814 SLAC superfamily members. Position 194 has glycine in 58% of SF1 homologs and
alanine in another 22%. Asp194, which never occurs naturally, would block the pore and be
expected to repel anions.
Mutational tests of channel function
Mutational studies corroborate the hypothesis that a TehA-based SLAC1 model is
appropriate. First, as discussed above, the slac1-1 mutant (S456F) is expected to be
structurally disruptive, and indeed it is inactive in guard cells4, and the slac1-2 mutation
(G194D) is expected to block the pore, and we show below that this variant is also inactive.
We have also shown that the introduction of SLAC1-conserved proline residues into
HiTehA (A208P/G263P) is accommodated isomorphously26. Moreover, as shown below,
channel conductance properties of several mutants are similar for AtSLAC1 and HiTehA.
To examine characteristics of the SLAC1 channel in light of the structural model, we
performed electrophysiological tests of membrane currents from voltage-clamped Xenopus
oocytes following injection of wild-type or mutant AtSLAC1 or HiTehA cRNAs. We
observed modest-sized chloride currents with wild-type AtSLAC1 cRNA, as found
previously6,7, but did not detect any chloride current following injection of wild-type
HiTehA cRNA. We found that SLAC1 Cl− conductance was enhanced when the OST1
kinase cRNA was co-injected with SLAC1 cRNA, but only to the levels found by Lee et al.6
and not to the much higher levels found by Geiger et al.7 with OST1 physically connected to
SLAC1 by split YFP linkage. In keeping with structural evidence that Phe262 blocks the
HiTehA pore, removal of the phenyl group in HiTehA F262A or in the homologous
AtSLAC1 F450A resulted in very large Cl− currents relative to wild-type levels, and the
SLAC1 currents were now less enhanced by presence of OST1 (Fig. 4a). The tempting
interpretation of an opened gate will require validation with appropriately analyzed single-
channel recordings27. In keeping with the slac1-2 phenotype3, neither the functionally
impaired AtSLAC1 G194D nor its HiTehA G15D homolog showed any substantial
conductance; moreover, further consistent with pore blockage by Asp194 in slac1-2, the
large conductances of HiTehA F262A and AtSLAC1 F450A were stopped in the double
mutants AtSLAC1 G194D/F450A and HiTehA G15D/F262A (Fig. 4a). Here again, the
effects in SLAC1 were independent of OST1.
We also tested conductance characteristics for a series of AtSLAC1 F450X substitution
mutants – F450A,G,T,V,L– and for the corresponding HiTehA F262X series –
F262A,G,T,V,L (Figs. 4b,S5, Table S5). Findings from the two series are roughly parallel;
in particular, the alanine and glycine substitutions lead to large currents for both and in
comparison to the others. There are distinctions, of course, including generally higher
conductances for AtSLAC1 over HiTehA and less conductance of F262T TehA compared to
F450T SLAC1. It is also noteworthy that OST1 activation is very muted for the F450A,G,L
mutants, which is consistent with SLAC1 gating at Phe450.
Crystal structures were also determined for several of the HiTehA mutant variants (Table
S4). The structures of F262A (1.15Å), F262V (1.60Å), F262L (1.65Å) and G15D (1.50Å)
are all essentially isomorphous with the wild-type TehA structure with changes localized
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primarily at the sites of mutation; the same is true for the double mutations of F262A/G15D,
F262G/G15D and A208P/G263P26. The F262A structure has a wide open pore (Fig. 5a)
with a relatively uniform pore diameter of ~5Å through ~30Å across the membrane (Fig.
S3) whereas G15D has a doubly occluded pore (Fig. 5b). The pores of other mutant variants
are consistent with the sizes of constrictive residues and with the observed conductances.
Gating and activation
The crystal structures of TehA and its mutant variants when taken together with the
functional studies in Xenopus oocytes point to a crucial role for Phe450 in gating of the
SLAC1 anion channel. Conservation of this residue across the SLAC1 family implies
functional importance. The occlusion of the pore by the presence of F262 in the structure of
wild-type TehA and the openness of the pore upon its substitution by alanine in the structure
of the F262A mutant provides physical evidence for a gating role of this residue. This
interpretation is supported by the correlated conductance characteristics from variants of the
AtSLAC1 and HiTehA channels (Figs. 4b,S5). While these observations may suffice for
placing the gate within the channel pore, they do not by themselves suggest a mechanism for
gating in response to physiological stimuli. Some insight does come from conformational
details defined at high resolution.
One important structural clue is that the side chain of Phe262 is in a high energy
conformation in the HiTehA structure, with χ1/χ2 at −160°/-4°. Although χ1 is in a preferred
trans conformation, the phenyl ring is restricted by contacts with Val210 and Leu18 to a χ2
value near 0° rather than near to the preferred 90° orientation (Fig. 5c). Further evidence that
Phe262 is restrained from local equilibrium comes from shifts observed in crystal structures
of the F262A, F262V, F262L and F262G/G15D variants, which all show consistent
backbone movements that displace Cβ(262) by 0.47-0.78Å (Fig. 5d). By contrast, Leu262 in
F262L is in a preferred trans/gauche+ conformation at χ1/χ2 = 177°/63° as is Val262 in
F262V at χ1 = −176°. What might control activation of HiTehA is unclear, but for AtSLAC1
activation is by OST1 phosphorylation6,7. The molecular consequences of OST1
phosphorylation of SLAC1 remain unknown, but it is plausible that associated shifts in pore-
helix orientations would unlatch Phe450 in SLAC1 from a TehA-like restrained orientation.
By analogy with Leu262 in F262L, we expect a preferred rotameric state for Leu450 in
AtSLAC1 F450L. Thus, the lack of appreciable OST1 activation of conductance in
AtSLAC1 F450L (Fig. 5e) might be explained by the lack of a restraining latch, whereby the
channel remains closed despite OST1 activation. Puzzles certainly remain since OST1 does
substantially activate AtSLAC1 F450T or F450V, which like HiTehA F262V should also be
unrestrained; presumably activating adjustments widen the pore enough for ion permeation
past threonine and valine but not leucine.
Phosphorylation sites have been discovered in the N- and C-terminal tails of AtSLAC16,7,28
(179 and 51 residues long, respectively), but these alone cannot explain OST1 activation of
SLAC1. First SLAH1, which fully complements the slac1-1 mutation, does not have these
cytoplasmic tails. Secondly, although OST1 phosphorylation of Ser120 in the N-terminal tail
is necessary for SLAC1 activation, it is not sufficient7. Thus, we surmise that direct
phosphorylation of the SLAC1 TM domain must be critical, and SLAC1 has four conserved
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Ser/Thr candidates in its cytoplasmic loops. Moreover, SLAC1 proteins have proline-
mediated kinks at the putative Phe450 gate in helix TM9, and also in adjacent helix TM7;
these features may play a role in phosphorylation-driven unlatching of the Phe450 gate in
SLAC1.
Ion selectivity and discrimination
Our studies of SLAC1 channel relative ion permeabilities, based on measurements of current
reversal potential, are consistent with earlier work demonstrating that AtSLAC1 conducts
anions but not cations and is selective among anions, with greater permeability for nitrate
than for chloride (as in Vicia faba guard cell protoplasts27) and much reduced permeability
for malate, bicarbonate or sulfate6,7. We also find that SLAC1 has little permeability for
sulfite. Additionally, we find that wild-type SLAC1, F450A and F450T all have similar
relative permeabilities to chloride, sulfite and malate, despite having widely different
conductance levels, but the gating mutants do show small but significant decreases in nitrate
permeability (Fig. 4c, Table S6).
The relative insensitivity of anion permeability to gating residue changes suggests that
selectivity for these anions may occur away from the central constriction at the channel gate.
To some extent, ionic discrimination must depend on pore geometry; thus, an organic anion
such as malate may be simply too large to pass through the 5-Å wide pore. Although the
SLAC1 pore is lined largely with hydrophobic sidechains (Fig. 3c), it is sprinkled with
hydroxyl groups from serine and threonine residues (16%); their electropositive hydrogen
atoms may facilitate conductance. Most strikingly, the electrostatic potential within the
AtSLAC1 pore is electropositive throughout (Fig. 3a). This polarization, promoted by
charges on extra-membranous loops, no doubt contributes significantly in discrimination
against cations.
The relative anion permeability sequence of SLAC1 determined by us and others, I− > NO3−
> Br− > Cl− 6,7, corresponds to selectivity sequence 1 compiled by Wright and Diamond29
for a range of anion-selective proteins. This sequence correlates inversely with the hydration
energies of monovalent anions – anions with a lower hydration energy have a greater
channel permeability. It is thought to be generated in proteins with weak, low field-strength,
anion binding sites, where selectivity is largely determined by the energetic cost of anion
dehydration. These selectivity results are thus consistent with the SLAC1 structure, where
the pore lacks any obvious anion binding site.
Distinctiveness of the SLAC1 channel
SLAC1 anion channels are entirely novel in structure and, apparently, in the mechanism for
ion conductance. The best characterized of anion channels belong to the CLC family of Cl−
channels and transporters30-32. CLC channels have an altogether different architecture from
the SLAC1 channel, and the mechanism for selectivity is also very different. Bacterial CLC
transporters bind halide ions at three sites in a highly constricted pore30. By contrast, the
SLAC1 pore has a relatively uniform diameter across the membrane, except where closed by
the gating phenyl group, and we do not find discrete ion binding sites. CLC selectivity is
governed by specific residues surrounding these binding sites30,32. The anion selectivity
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sequence for the CLC channels of Cl−>Br−>NO −3>I−, opposite of that in SLAC1, is
consistent with the high field-strength anion binding sites in CLC channels29. Interestingly,
as for AtSLAC1, an Arabidopsis CLCa channel also preferentially transports nitrate ions33,
and an E. coli CLC channel is converted to preference of nitrate when a generally conserved
serine at the central site is substituted with proline as in AtCLCa32.
SLAC1 also differs radically from other structurally characterized anion channels and
transporters. These include the VDAC1 voltage-gated anion channel from mitochondrial
outer membranes, which has a porin-like beta-barrel structure34-36, and a light-driven
halorhodopsin chloride pump, which has a transmembrane conductance pathway similar to
that of the proton-pumping pore of bacteriorhodopsin37. Although its channel structure is
still only known by homology to other ABC transporters, CFTR is another obviously
distinct chloride channel38. Cys-loop receptors also include anion channels39, and these are
similar to SLAC1 in having five-helix pores25, but here selectivity is governed by charged
groups at the entrance to the pore, which distinguish the anion-selective GABAA and
glycine receptors from the cation-selective acetylcholine and serotonin 5HT3 receptors39.
Finally, the recently identified TMEM16A gene for calcium-activated chloride channels40-42
appears to encode an 8-TM protein that is again distinct from SLAC1.
Stomatal guard cells show both rapidly activated (R-type) and slow (S-type) anion channel
activity43. Although slac1 guard cells have very defective S-type activity, their R-type
currents are normal4. Guard cell protoplasts from the slac1-2 mutant abnormally accumulate
Cl−, K+, malate and fumarate3, whereas SLAC1 shows negligible malate conductance7. As
for SLAC1-associated K+ movements, other channels or transporters must be responsible for
SLAC1-associated malate movements. Recent studies indicate that AtALMT12, an
aluminium-activated malate transporter (ALMT) family member, is a malate-dependent R-
type anion channel44 needed for stomatal closure45.
Conclusions
We find that many functional properties of the plant SLAC1 anion channel are explained
well by the structure of an uncharacterized bacterial TehA protein that has been associated
with tellurite resistance. SLAC1 and TehA belong to distinct subfamilies within one branch
of a larger SLAC1 superfamily, but AtSLAC1 and HiTehA are sufficiently similar (19%
sequence identity) that the SLAC1 homology model is predictive for function, including a
verified placement of the identifying slac1-2 mutation G194D and a phenylalanine gate. One
remaining puzzle concerns the structural change that activating phosphorylation elicits in
SLAC1, and another puzzle is the biochemical role of the TehA homologs in bacteria. In a
companion paper26, we examine functional and structural properties of TehA in bacteria,
showing that it is anion channel, although actually not conferring tellurite resistance, and
identifying a mutant variant with properties suggestive of an activated state. Thus, SLAC1
and TehA likely represent a large family of selective anion channels controlled by
environmental stimuli.
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METHODS
Selection of target sequences
TehA from E. coli (EcTehA) was a centrally selected sequence for expansion as a
NYCOMPS targeted family. Target selection criteria utilized at NYCOMPS are described in
details elsewhere46. Briefly, the EcTehA sequence was run against a dataset of about 40,000
predicted alpha helical integral membrane protein sequences from prokaryotic genomes
(NYCOMPS98 dataset46) using PSI-BLAST50. Sequences that matched EcTehA with an E-
value lower than 10−3 in an alignment extending over at least 50% of both predicted TM
regions and passing our post-seed-expansion filtering criteria46 were passed to the protein
production pipeline.
Protein expression screening
Full-length homologs from the following 38 species, including 2 sequences each from 5 of
these, were amplified from genomic DNA by PCR: Thermoplasma acidophilum,
Lactococcus lactis subsp. lactis Il1403, Streptococcus pyogenes M1 GAS, Streptococcus
pneumoniae TIGR4, Haemophilus influenzae Rd KW20, Methanocaldococcus jannaschii
DSM 2661 (2), Escherichia coli K12, Salmonella typhimurium LT2, Clostridium
perfringens ATCC 13124, Xanthomonas campestris pv. campestris str. ATCC 33913,
Streptococcus agalactiae 2603V/R, Shewanella oneidensis MR-1, Streptococcus mutans
UA159, Archaeoglobus fulgidus DSM 4304, Vibrio parahaemolyticus RIMD 2210633 (2),
Pseudomonas syringae pv. tomato str. DC3000 (2), Enterococcus faecalis V583,
Pyrococcus horikoshii OT3, Bordetella bronchiseptica RB50, Streptomyces coelicolor A3,
Corynebacterium glutamicum ATCC 13032, Picrophilus torridus DSM 9790, Acinetobacter
sp. ADP1, Vibrio fischeri ES114, Pseudomonas fluorescens Pf-5, Sulfolobus acidocaldarius
DSM 639, Colwellia psychrerythraea 34H, Neisseria meningitidis MC58, Rhodobacter
sphaeroides 2.4.1, Vibrio cholerae O1 biovar eltor str. N16961 (2), Marinobacter aquaeolei
VT8, Acinetobacter baumannii ATCC 17978, Klebsiella pneumoniae subsp. pneumoniae
MGH 78578, Streptomyces avermitili MA-4680, Bordetella parapertussis 12822 (2),
Streptococcus thermophilus LMG 18311, Salmonella enterica subsp. enterica serovar
Paratyphi A str. ATCC 9150, and Anaeromyxobacter dehalogenans 2CP-C.
Selected cDNAs were cloned into a modified pET vector (Novagen, Inc) that fuses a FLAG
and deca-histidine tag at the C-terminus, which are cleavable by TEV protease. Proteins
were expressed in E. coli BL21(DE3) plysS by a high throughput format (0.6ml in a deep
well block) and purified after lysis by sonication using metal affinity purification in a buffer
containing N-Dodecyl-β-D-Maltopyranoside. Samples were passed over an analytical size
exclusion column in 12 different detergent-containing mobile phases, which included N-
dodecyl-β-D-maltopyranoside (DDM), N-decyl-β-D-altopyranoside (DM), N-nonyl-β-D-
altopyranoside (NM), N-octyl-β-D-altopyranoside (OM), N-octyl-β-D-Glucopyranoside
(OG), N-nonyl-β-D-Glucopyranoside (NG) and lauryl dimethyl amine oxide (LDAO).
Multi-angle light scattering with refractive index detection was used to analyze the
oligomeric state51. The E. coli and H. influenzae proteins were judged to be monodisperse
and stable and were passed to scale up.
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Scaled-up production and purification
For scale up, in brief, transformed BL21 plysS cells were grown at 37°C in 2× YT media to
OD 0.6-0.8 after being inoculated with 1% of the overnight culture. The culture was induced
with 0.4mM IPTG and continued to grow at 37 °C for another 4 hours. The cells were
harvested by centrifugation and stored at −80°C before use. Selenomethionyl (SeMet) TehA
was expressed in a similar way, but using containing SeMet in place of methionine in
defined minimal media. Cells were resuspended in a buffer containing 50 mM Tris-HCl (pH
8.0) and 200 mM NaCl and lysed using a French Press with two passes at 15-20,000 psi.
Cell debris were removed by centrifugation at 10,000g for 20min, and the membrane
fraction was isolated from that supernatant by ultra-centrifugation at 150,000g for 1 hr.
The membrane fraction was homogenized in a solubilization buffer containing 50 mM Tris
(pH 8.0) and 200 mM NaCl, and incubated with final concentration of 1% (w/v) dodecyl-β-
D-maltopyranoside (DDM, Anatrace, Inc) for 1 hr at 4°C. The non-dissolved matter was
remove by ultracentrifugation at 150,000g for 30 min, and the supernatant was loaded to a
5ml Hitrap Ni2+-NTA affinity column (GE Healthcare), pre-equilibrated with the same
solubilization buffer supplemented with 0.05% DDM. After 20 column volume buffer wash,
the protein was eluted with 250mM imidazole in the solublization buffer. The Flag and 10-
His tags were removed by adding super TEV at 1:100 mass ratio and incubating at 4°C
overnight. Tag removal was confirmed by SDS-PAGE, and the resulting sample was
concentrated to around 10mg/ml. Preparative size exclusion chromatography was carried out
on a Superdex-200 column for further purification, removal of TEV protease and the
cleaved tag, and for buffer and detergent exchange. The gel-filtration buffer contained 10
mM Tris (pH 8.0), 200 mM NaCl, 1mM EDTA, 0.5 mM Tris [2-carboxyethyl] phosphine
(TCEP), and 2×CMC of detergent. In the case of HiTehA, the protein was well behaved and
stable in nearly all tested detergents, and we have purified it from DDM, DM, NM, OM, OG
and LDAO.
Protein characterization
We performed N-terminal amino-acid sequencing of purified HiTehA and EcTehA prior to
TEV protease treatment. Results from these analyses proved that true initiating methionine
residue is located 14 residues after one annotated as N-terminal. The intervening nucleotide
sequence contains a Shine-Delgarno sequence.
For cross-linking experiments, purified HiTehA was incubated with 10 mM disuccinimidyl
glutarate at room temperate for 30 min, and 100mM Tris-HCl pH 7.5 was added to stop the
reaction. The incubated sample when run on a 8-25% gradient SDS-PAGE gel showed a
ladder consistent with a trimeric structure.
Crystallization and data collection
Purified protein was concentrated to ~10mg/ml for initial crystal trials in a Mosquito robot
with commercial screens from Hampton research, Emerald Biosystems and Molecular
Dimension. We obtained crystals of HiTehA from protein in detergent DDM, DM, NM,
OM, OG and LDAO, but only those from LDAO and OG gave diffraction to beyond 4Å
spacings. Crystals that proved useful were all grown at 4°C using the sitting-drop vapor
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diffusion method. After extensive optimization we reached conditions supporting very high
resolution. Wild-type HiTehA and most of its variants crystallized from 1mM ZnSO4,
50mM Hepes-Na pH 7.8 and 28% PEG600. HiTehA F262A crystallized from 200mM
Li2SO4, 100mM glycine pH9.3 and 33% PEG400). Addition of 10mM spermidine as an
additive helped to obtain slightly better diffracting crystals. Cryoprotection was achieved by
adding 5% ethylene glycol or PEG400 to the crystallization solution.
Structure determination and refinement
Native and SeMet SAD data set were collected at NSLS beamline X4A and processed using
the software HKL200052. Crystals of HiTehA grew in space group R3 with a=b=c = 85.0Å
and α=β=γ = 93.5°. All subsequent manipulations were done in the hexagonal setting of this
space group with a=b= 96.0Å and c = 136.7Å. The asymmetric unit contains one TehA
protomer and 65% solvent volume. The structure was determined at 2.0Å resolution from
single-wavelength anomalous diffraction (SAD) using selenomethionine-substituted protein
crystals. Assessment of data quality for phasing, location of heavy atom sites and initial
phases were calculated using the HKL2MAP interface to SHELX programs53.
All the secondary structure elements were clearly visible in the experimental electron
density map. Automatic model building was done in Arp/wArp54 and completed manually
in the program COOT55. The model was refined against native data at 1.20Å resolution
using the program Refmac5.5 in CCP456, with anisotropic B factor restrained refinement
applied. Subsequent structural analyses of mutant variants were refined as isomorphous
structures.
Site-directed mutagenesis
Site-directed mutants were constructed using the QuikChange Site-Directed Mutagenesis Kit
(Stratagene, La Jolla, CA 92037) and expressed from pET vectors in E. coli BL21(DE3)
plysS cells as for the wild-type protein.
Electrophysiology
All constructs were cloned into plasmid pGHME2, linearized, and transcribed into cRNA
using T7 polymerase (mMessage mMachine, Ambion). Oocytes were injected with 50 nl of
cRNA solution each, at a constant concentration of 0.5 mg/ml for HiTehA, or 0.5 mg/ml for
AtSLAC1 constructs, with or without 0.5 mg/ml of AtOST1 constructs and recorded in
voltage-clamp experiments 2 days after injection. For mixed expression experiments, 25nl of
cRNA solution was injected for each AtSLAC1 component. Two-electrode voltage clamp
recordings were performed to measure HiTehA or AtSLAC1 currents as described6,7. The
pipette solutions contained 3 M KCl. For voltage-clamp current recordings, the bath solution
contained 1 mM MgCl2, 1 mM CaCl2, 10 mM Mes/Tris (pH 5.6) and 30 mM CsCl; For
anion selectivity measurements, the bath solutions contained 50 mM Cl−, NO −3, malate, or
30 mM SO 2-3 sodium salts, 45 mM Na-gluconate, 1 mM Ca-gluconate2, 1 mM Mg-
gluconate2, 1 mM K-gluconate, as well as 10 mM Tris/Mes (pH 5.6). Osmolality was
adjusted with D-mannitol to 220 mOsmol/kg. The bath electrode was a 3M KCl agar bridge.
The testing potentials were +50 to −110 or −130 mV in 20 mV decrements with 7.5 s
duration. The holding and return potentials were 0 mV with a 1.45 s or 2 s, respectively. I-V
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relations of HiTehA or AtSLAC1 were generated from the current reached after 0.5 s in
testing potentials. The Goldman-Hodgkin-Katz equation was applied to estimate
permeability ratios for monovalent ions as described6. For divalent anions, the permeability
ratios were derived according to Fatt and Ginsborg57.
Bioinformatic analysis of SLAC-related proteins
Sequences related to SLAC1 were analyzed comprehensively by PSI-BLAST50. Searches at
E<10−3 starting from five disparate homologs each identified a common pool of over 900
proteins, which when pooled were used for sub-classification into families and subfamilies.
Details of these analyses are reported in footnotes to Table S1.
Molecular figures were produced in PyMOL58.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We thank Filippo Mancia, Larry Shapiro and Ming Zhou for helpful discussions; Patricia Rodriguez and Arianne
Morrison for help with cloning and expression; Jonah Cheung, Mark Collins, Chris Min, Sheng Ye and Zhen Zhang
for advice in protein chemistry and crystallography; John Schwanof and Randy Abramowitz for help with
synchrotron experiments; John Riley for help with Xenopus oocyte injections; and Min Su for advice on the PSI-
BLAST analysis of SLAC relatives. This project was supported in part by an award to the New York Consortium
on Membrane Protein Structure (NYCOMPS) from the NIGMS Protein Structure Initiative. Beamline X4A at the
National Synchrotron Light Source (NSLS), a DOE facility at Brookhaven National Laboratory, is supported by the
New York Structural Biology Center.
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Figure 1. Sequence analysis for the SLAC1 superfamily
a, Family tree. The presentation was computed by the program COBALT47 from
representative subfamily sequences (Table S1), including Arabidopsis thaliana SLAC1 for
SF1Ai, Haemophilus influenzae TehA for SF1Ci, Escherichia coli TehA for SF1Cii, Vibrio
parahaemolyticus for SF2A, Staphylococcus aureus for SF2B, Aspergillus fumigatus Ssu1
for SF3A, and Schizosaccharomyces pombe Mae1 for SF3B. b, Structure-based sequence
alignment of TehA from Haemophilus influenzae (HiTehA) and SLAC1 from Arabidopsis
thaliana (AtSLAC1). The TehA structure has been used to restrict sequence gaps to inter-
helical segments. Superior coils define extents of the HiTehA helical segments; red letters
mark residue identities; red boxes are drawn for residues that are >95% identical within the
plant subfamily SF1A for AtSLAC1 or within the TehA subfamily for HiTehA; red
diamonds mark HiTehA residues that line the central pore; and the colored inferior bar
encodes ConSurf sequence variability48 for the SF1 family of 204 non-redundant proteins.
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Figure 2. Crystal structure of HiTehA and homology model of AtSLAC1
a, Electron density distribution from the HiTehA crystal structure at 1.2Å resolution. The
map has (2Fo-Fc) coefficients based on the superimposed model. Contours are at 2.5σ. b,
Ribbon diagram of the HiTehA trimer. Each protomer is colored spectrally from blue at its
N-terminus to red at its C-terminus. c, DelPhi49 electrostatic potential at the extracellular
surface. Electronegative and electropositive potential are colored in degrees of red and blue
saturation, respectively. d, Electrostatic potential49 at the intracellular surface. e, Ribbon
diagram of an HiTehA protomer viewed from outside the membrane. The ribbon is colored
spectrally as in 2b. f, Ribbon diagram of an HiTehA protomer viewed from within the
membrane from below the view of e. g, Surface of a homology model of AtSLAC1, viewed
as in f, and colored by sequence variability48. h, Surface of AtSLAC1 as in g, but colored by
electrostatic potential49.
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Figure 3. Putative structure of the SLAC1 conductance pore
a, Cross-section through the homology model of AtSLAC1. The model is viewed as in 2i,
with the electrostatic potential49 shown on the external surface of the molecular envelope.
The side chain of Phe450 is shown as a stick model (red) on the backbone ribbon colored
yellow. b, Cross-section as in a, but colored by surface conservation48 as in 2h. c, Pore-
lining residues in the SLAC1 homology model. A cylinder model (left), spectrally colored as
in 2b, provides a key for viewing the rolled-open structure (left) with pore-lining residues of
AtSLAC1 shown on the TModd helices. d, Ribbon diagrams of HiTehA TM9 (left) and TM7
(right) viewed from within the conductance pore. The side chains of Pro207 and Phe262 are
shown as well as a kink-stabilizing water HOH25 that is coordinated by the NH of Gly263
and by C=O groups of Gly202 and Ala259. Density contours are shown for the water
molecule.
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Figure 4. Ionic conductance measurements
a, Typical microelectrode voltage-clamp current traces from oocytes injected with various
channel cRNAs. Left column, oocytes injected with cRNAs encoding wild-type HiTehA
channels (WT), or F262A, G15D/F262A or G15D mutants. Right two columns, oocytes
injected with cRNAs for WT AtSLAC1, or F450A, G194D/F450A or G194D mutants, with
or without co-injection of AtOST1. Dotted lines represent zero current levels. Extracellular
solution contained 30mM CsCl. Schematic keys at the left place phenyl and/or aspartyl
gates. b, Effects of gating residue mutations. Mean chloride currents, measured at −90 mV,
are shown comparing WT HiTehA and its mutant series F262A,G,T,V,L with WT AtSLAC1
and its corresponding series F450A,G,T,V,L, both alone and co-expressed with AtOST1.
Full I-V relations are shown in Fig. S5. c, Effect of gating residues on relative AtSLAC1
anion permeabilities. Relative permeabilities (P[X]/P[Cl]) for chloride, nitrate, sulfite and
malate of WT, F450A and F450T SLAC1 channels were measured from the change in
current reversal potential with Cl− or anion X− as the sole permeant anion in the bath
solution (Methods, Table S6).
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Figure 5. Structural features at the SLAC1 homolog gate
a,b, Cross-sections through the conductance pores of HiTehA F262A and HiTehA G15D.
The view and presentations are as in 3a, except that helices are colored purple. c, Molecular
basis for conformational strain in gating residue Phe262 of HiTehA. Helices TM1 (left),
TM9 (center) and TM7 (right) are viewed from within the pore and presented as ribbon
diagrams with selected side chains drawn in stick representation. The local low-energy
conformation for the phenyl ring (χ2 = 90°) is shown in thin lines with short contacts
indicated in dashed lines: Leu18 Cδ2 – Phe262 Cδ1, d = 2.4 Å; Val210 Cγ2 – Phe262 Cε2, d
= 2.8 Å. d, Conformational shifts consequent to release of strain in gating residue Phe262.
Cα backbone structures of F262A, F262V and F262L HiTehA were first superimposed onto
WT HiTehA, and residues 258-266 were drawn in stereo as superimposed. All backbone
atoms are shown for peptides 262±1, but only Cα atoms are shown otherwise. The WT
backbone and phenyl group are green; other backbone are all magenta; side chains of
Ala262, Val262 and Leu262 are cyan, blue and red, respectively; oxygen-directed bonds are
red and nitrogen-directed bonds are blue. e, Typical microelectrode voltage-clamp current
traces from oocytes injected with wild-type (WT) or F450L AtSLAC1 cRNA. Experimental
conditions and displays are as in 4a.
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|
3M73
|
Crystal Structure of Plant SLAC1 homolog TehA
|
Homolog Structure of the SLAC1 Anion Channel for Closing
Stomata in Leaves
Yuhang Chen1,6, Lei Hu2, Marco Punta6,7, Renato Bruni6, Brandan Hillerich6, Brian Kloss6,
Burkhard Rost1,6,1, James Love6, Steven A. Siegelbaum2,3,5, and Wayne A.
Hendrickson1,4,5,6
1Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY
10032, USA
2Department of Neuroscience, Columbia University, New York, NY 10032, USA
3Department of Pharmacology, Columbia University, New York, NY 10032, USA
4Department of Physiology and Cellular Biophysics, Columbia University, New York, NY 10032,
USA
5Howard Hughes Medical Institute, Columbia University, New York, NY 10032, USA
6NYCOMPS, New York Structural Biology Center 89 Convent Avenue, New York, NY 10027 USA
7Department of Computer Science and Institute for Advanced Study Technical University of
Munich D-85748 Munich, Germany
Summary
The plant SLAC1 anion channel controls turgor pressure in the aperture-defining guard cells of
plant stomata, thereby regulating exchange of water vapor and photosynthetic gases in response to
environmental signals such as drought or high levels of carbon dioxide. We determined the crystal
structure of a bacterial homolog of SLAC1 at 1.20Å resolution, and we have used structure-
inspired mutagenesis to analyze the conductance properties of SLAC1 channels. SLAC1 is a
symmetric trimer composed from quasi-symmetric subunits, each having ten transmembrane
helices arranged from helical hairpin pairs to form a central five-helix transmembrane pore that is
gated by an extremely conserved phenylalanine residue. Conformational features suggest a
mechanism for control of gating by kinase activation, and electrostatic features of the pore coupled
with electrophysiological characteristics suggest that selectivity among different anions is largely
a function of the energetic cost of ion dehydration.
Users may view, print, copy, download and text and data- mine the content in such documents, for the purposes of academic research,
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Correspondence and request for materials should be addressed to WAH (wayne@convex.hhmi.columbia.edu)..
Supplementary Information is linked to the online version of the paper at www.nature.com/nature.
Author Contributions YC, LH, MP, RB, BH, BK and JL performed experiments; YC, LH, BR, SAS, and WAH analyzed data; YC,
LH, SAS, and WAH prepared the manuscript.
Author Information Atomic coordinates and diffraction data are deposited in the Protein Data Bank with accession codes 3M71,
3M73, 3M74, 3M75, 3M76, and 3M7L. See Table S4 for identifications. Reprints and permissions information is available at
www.Nature.com/reprints.
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Published in final edited form as:
Nature. 2010 October 28; 467(7319): 1074–1080. doi:10.1038/nature09487.
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Stomatal pores in the leaves of plants permit the influx of atmospheric carbon dioxide in
exchange for transpirational evaporation of water1,2. A pair of kidney-shaped guard cells
define each pore aperture, and turgor pressure variation in these cells determines the degree
of stomatal pore openness. Depending on diverse environmental factors, the stomata close to
prevent H2O losses and open to admit CO2 for photosynthesis. Environmental stimuli that
lead to stomatal closure include darkness, high CO2 levels, ozone, low air humidity and
drought. The plant hormone abscisic acid (ABA) is critical for signal transduction from
these stimuli. Mutational screens in Arabidopsis thaliana for CO2 and ozone sensitivity
identified a protein with ten predicted transmembrane (TM) helices, now called slow anion
channel 1 (SLAC1), as having a central role in the control of stomatal closure3-5. Recent
studies proved that SLAC1 is indeed an anion channel6-7, with characteristics like those of
slow anion channels found in guard cells8, and that it is activated by phosphorylation from
the OST1 kinase9. OST1 activity is negatively regulated by the ABI1 phosphatase10,11,
which is in turn inhibited by the stomatal ABA receptors PYR/RCAR12,13 when in the
ternary hormone-receptor-phosphatase complex14-18. Thereby, ABA stimulates SLAC1
channel activity. Resulting Cl− efflux through SLAC1 causes membrane depolarization,
which activates outward-rectifying K+ channels, leading to KCl and water efflux to further
reduce turgor and cause stomatal closure.
SLAC1 expression in Arabidopsis is confined to the guard cells of leaves, but other
Arabidopsis tissues do have SLAC1 homologs3, named SLAH1-SLAH4. The identifying
mutations slac1-14 and slac1-23 are, respectively, in predicted transmembrane segments 9
(S456F) and 1 (G194D) of a protein that includes substantial N- and C-terminal extensions
outside a 10-helix transmembrane (TM) domain. SLAH1, which is absent from leaves and
lacks the terminal extensions of SLAC1, fully complements the mutant phenotype in slac1-2
guard cell protoplasts3. SLAC1 and homologs are also present in other plant genomes,
including nine in rice (Oryza sativa) and five in grape vines (Vitis vinifera). SLAC1
relatives, some quite remote, also occur in bacteria, archaea and fungi. Known prokaryotic
homologs contain only the predicted transmembrane domain of SLAC1, but some fungal
homologs do have N- and C-terminal extensions. One homolog, Mae1 from the yeast
Schizosaccharomyces pombe, functions as a malate uptake transporter19; another, Ssu1 from
Saccharomyces cerevisiae and other fungi including Aspergillus fumigatus, is characterized
as a sulfite efflux pump20,21; and TehA from Escherichia coli is identified as a tellurite
resistance protein by virtue of its association in the tehA/tehB operon22,23. Despite a lack of
further biochemical characterization, many homologs are annotated as tellurite resistance/
dicarboxylate transporter (TDT) proteins.
We have undertaken structural and functional characterizations of the SLAC1 anion
channel. We first solved an atomic-resolution crystal structure of the TehA homolog from
Haemophilus influenzae, and we then developed a homology model for Arabidopsis SLAC1.
This model allowed us to conduct mutagenesis for functional testing of structure-inspired
hypotheses on gating and selectivity. We expressed Haemophilus TehA and Arabidopsis
SLAC1 in Xenopus oocytes to characterize channel properties of these proteins and mutant
variants. We also determined crystal structures for several mutant variants, including the
homolog of slac1-2.
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Structure of SLAC1 bacterial homolog TehA
We performed a bioinformatic analysis of SLAC1-related proteins, first clustering nearly
900 non-redundant sequences into a superfamily at the PSI-BLAST level E≤10−3, then into
three distinct families at an initial threshold of E≤10−30, and finally into subfamilies at a
typical initial threshold of E≤10−55. Since previous annotation is not well founded in
experiment and SLAC1 is now the best characterized member, we adopt a nomenclature
defining a SLAC superfamily divided into families identified as SF1-SF3 and subfamilies
SF1A, SF1B etc. Family SF1 comprises the plant SLAC proteins and close bacterial
homologs, family SF2 comprises a distinct set of bacterial proteins often annotated as
exfoliative toxins, and family SF3 comprises the fungal Mae1 and Ssu1 proteins and their
archaeal or bacterial homologs, respectively. SLAC family SF1 has three large subfamilies:
the plant SLAC and SLAH proteins are in subfamily SF1A, closest bacterial homologs are
in SF1B, and the TehA homologs are in SF1C (Fig. 1a). The other families also divide into
subfamilies as detailed in Table S1, and family SF1 is divided into sub-subfamilies (Table
S2). Two pertinent SF1 sequences are aligned in Fig. 1b.
We used a structural genomics approach to obtain structural information, testing expression
and purification for 43 bacterial and archaeal likely homologs, assaying for detergent choice
and stability on 8 of these, finding two with appropriate profiles by size exclusion
chromatography, and obtaining suitable crystals for one. This protein, TehA from H.
influenzae (HiTehA), was found to be trimeric both by size-exclusion multi-angle light
scattering (SEC-MALS) measurements and by chemical cross-linking. When solubilized in
β-octylglucoside, HiTehA crystallized in space group R3 with a=b=96.01Å and c=136.27Å.
Each asymmetric unit contains one subunit and 65% solvent. The structure was solved by
selenomethionyl (SeMet) SAD phasing, ultimately at 1.50Å resolution (Table S3, Fig. S1),
and then refined at 1.20Å resolution (Fig. 2a) to R/Rfree values of 14.1%/15.7% for a model
that includes ordered residues 6-313, 213 water molecules and four detergent molecules
(Table S4).
The crystal structure has TehA trimers aligned with three-fold axes of the lattice (Fig. 2b).
Subunits are tightly associated, burying 8947 Å2 of total surface area within trimer
interfaces. The electrostatic potential surface is largely negative on the extracellular surface
(Fig. 2c) and largely positive on the cytoplasmic surface (Fig. 2d). The membrane
orientation is specified experimentally from GFP tagging of E. coli TehA24. Each TehA
protomer has ten TM helices, as predicted; however, the fold is novel. Tandemly repeated
helical hairpins are arranged with quasi-five-fold symmetry (Figs. 2e,S2). Extracellular
inter-helix loops are short (2-5 residues) whereas intracellular inter-helix connections are
longer, including a nine-residue helix H2,3 between TM2 and TM3 (Fig. 2f). An inner pentad
of outwardly directed, TModd, helices creates an apparent pore through each protomer
perpendicular to the putative membrane plane. TMeven helices from the five hairpins
surround the inner pore and make an outer layer.
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Homology model for plant SLAC1
Arabidopsis SLAC1 (AtSLAC1) is substantially similar to bacterial homologs, notably
HiTehA (Fig. 1b). All HiTehA TM helices are fully aligned to predicted SLAC1 TM
helices, but there are short inter-helical gaps (1-5 residues) in all five extracellular loops and
in two of the intracellular loops. The TM domain of AtSLAC1 (residues 188-504) aligns to
HiTehA with 19% sequence identity and with a PSI-BLAST E-value of 3×10−22. For
comparison, and in keeping with the family tree (Fig. 1a), the TM domain of AtSLAC1
shares sequence identities of 76% with rice SLAC1, 41% with Arabidopsis SLAH1, 25%
with an SL1B homolog from Halorhodospira halophila, 11% with S. cerevisiae Ssu1, and
9% with S. pombe Mae1. A conceptual model with the AtSLAC1 sequence transposed onto
the HiTehA helices sufficed to guide most of our mutational tests, but a detailed AtSLAC1
homology model helped to refine our ideas. Surface variability and electrostatic potential are
plotted onto the surface of this model (Fig. 2g,2h).
The most remarkable feature of the TehA structure and corresponding SLAC1 model is the
central pore through each protomer. As for acetylcholine receptors25, the SLAC1 pore is
formed by five helices; but the SLAC1 helices come from one protein molecule rather than
five. The SLAC1 pore has a relatively uniform diameter of approximately 5Å across nearly
five helical turns (Fig. S3), except for a pronounced constriction in the middle of the
membrane (Fig. 3a) where the pore is occluded by the side chain of Phe 450 (Phe 262 in
HiTehA). This residue is the only absolutely conserved amino acid residue of the SLAC1
family. The pore is lined with highly conserved (86% identity among five SLAC1 orthologs;
32% identity between AtSLAC1 and HiTehA) and generally hydrophobic residues (Figs. 1b,
3b,3c,S4). Despite this hydrophobicity, the electrostatic potential on the pore surface is
polarized (Fig. 3a), presumably due to an invaginated shape adjacent to charged residues
outside the membrane. The generally electropositive character of the cytoplasmic surface
likely contributes to anion efflux.
Kinks in the pore helices contribute to formation of a relatively constant pore diameter
across the membrane. Four of the five HiTehA inner helices have centrally located proline
residues, which necessarily generate kinks, and TM9 is kinked at a backbone-coordinated
water molecule (Fig. 3d). Proline replaces Gly263 in all SF1A and SF1B relatives, including
Pro451 AtSLAC1. This water-displacing change is isostructural26. Centrally located prolines
also prevail in TM3, TM5 and TM7 across the SF1 proteins. The outer helices are longer and
straighter, but more inclined. The only outer-helix proline kink of HiTehA is in TM6 at the
trimer three-fold axis.
Two of the gene-identifying mutations in Arabadopsis SLAC1 are selected point mutations,
others are disruptive transfer DNA (T-DNA) insertions4. In the AtSLAC1 homology model,
the slac1-2 (G194D) mutation3 points into the pore from TM1 and can be accommodated
structurally whereas the slac1-1 (S456F) mutation4 points away from the pore six residues
after pore-blocking Phe450 on TM9 and would be expected to disruptive. Ser456 interacts
with outer helix TM10 in the homology model, and the phenyl bulk from S456F would not
fit. Position 456 has alanine in HiTehA, 66% of all 204 SF1 homologs and another 27%
have threonine or serine (as in all SLAC1 channels); phenylalanine never occurs among all
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814 SLAC superfamily members. Position 194 has glycine in 58% of SF1 homologs and
alanine in another 22%. Asp194, which never occurs naturally, would block the pore and be
expected to repel anions.
Mutational tests of channel function
Mutational studies corroborate the hypothesis that a TehA-based SLAC1 model is
appropriate. First, as discussed above, the slac1-1 mutant (S456F) is expected to be
structurally disruptive, and indeed it is inactive in guard cells4, and the slac1-2 mutation
(G194D) is expected to block the pore, and we show below that this variant is also inactive.
We have also shown that the introduction of SLAC1-conserved proline residues into
HiTehA (A208P/G263P) is accommodated isomorphously26. Moreover, as shown below,
channel conductance properties of several mutants are similar for AtSLAC1 and HiTehA.
To examine characteristics of the SLAC1 channel in light of the structural model, we
performed electrophysiological tests of membrane currents from voltage-clamped Xenopus
oocytes following injection of wild-type or mutant AtSLAC1 or HiTehA cRNAs. We
observed modest-sized chloride currents with wild-type AtSLAC1 cRNA, as found
previously6,7, but did not detect any chloride current following injection of wild-type
HiTehA cRNA. We found that SLAC1 Cl− conductance was enhanced when the OST1
kinase cRNA was co-injected with SLAC1 cRNA, but only to the levels found by Lee et al.6
and not to the much higher levels found by Geiger et al.7 with OST1 physically connected to
SLAC1 by split YFP linkage. In keeping with structural evidence that Phe262 blocks the
HiTehA pore, removal of the phenyl group in HiTehA F262A or in the homologous
AtSLAC1 F450A resulted in very large Cl− currents relative to wild-type levels, and the
SLAC1 currents were now less enhanced by presence of OST1 (Fig. 4a). The tempting
interpretation of an opened gate will require validation with appropriately analyzed single-
channel recordings27. In keeping with the slac1-2 phenotype3, neither the functionally
impaired AtSLAC1 G194D nor its HiTehA G15D homolog showed any substantial
conductance; moreover, further consistent with pore blockage by Asp194 in slac1-2, the
large conductances of HiTehA F262A and AtSLAC1 F450A were stopped in the double
mutants AtSLAC1 G194D/F450A and HiTehA G15D/F262A (Fig. 4a). Here again, the
effects in SLAC1 were independent of OST1.
We also tested conductance characteristics for a series of AtSLAC1 F450X substitution
mutants – F450A,G,T,V,L– and for the corresponding HiTehA F262X series –
F262A,G,T,V,L (Figs. 4b,S5, Table S5). Findings from the two series are roughly parallel;
in particular, the alanine and glycine substitutions lead to large currents for both and in
comparison to the others. There are distinctions, of course, including generally higher
conductances for AtSLAC1 over HiTehA and less conductance of F262T TehA compared to
F450T SLAC1. It is also noteworthy that OST1 activation is very muted for the F450A,G,L
mutants, which is consistent with SLAC1 gating at Phe450.
Crystal structures were also determined for several of the HiTehA mutant variants (Table
S4). The structures of F262A (1.15Å), F262V (1.60Å), F262L (1.65Å) and G15D (1.50Å)
are all essentially isomorphous with the wild-type TehA structure with changes localized
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primarily at the sites of mutation; the same is true for the double mutations of F262A/G15D,
F262G/G15D and A208P/G263P26. The F262A structure has a wide open pore (Fig. 5a)
with a relatively uniform pore diameter of ~5Å through ~30Å across the membrane (Fig.
S3) whereas G15D has a doubly occluded pore (Fig. 5b). The pores of other mutant variants
are consistent with the sizes of constrictive residues and with the observed conductances.
Gating and activation
The crystal structures of TehA and its mutant variants when taken together with the
functional studies in Xenopus oocytes point to a crucial role for Phe450 in gating of the
SLAC1 anion channel. Conservation of this residue across the SLAC1 family implies
functional importance. The occlusion of the pore by the presence of F262 in the structure of
wild-type TehA and the openness of the pore upon its substitution by alanine in the structure
of the F262A mutant provides physical evidence for a gating role of this residue. This
interpretation is supported by the correlated conductance characteristics from variants of the
AtSLAC1 and HiTehA channels (Figs. 4b,S5). While these observations may suffice for
placing the gate within the channel pore, they do not by themselves suggest a mechanism for
gating in response to physiological stimuli. Some insight does come from conformational
details defined at high resolution.
One important structural clue is that the side chain of Phe262 is in a high energy
conformation in the HiTehA structure, with χ1/χ2 at −160°/-4°. Although χ1 is in a preferred
trans conformation, the phenyl ring is restricted by contacts with Val210 and Leu18 to a χ2
value near 0° rather than near to the preferred 90° orientation (Fig. 5c). Further evidence that
Phe262 is restrained from local equilibrium comes from shifts observed in crystal structures
of the F262A, F262V, F262L and F262G/G15D variants, which all show consistent
backbone movements that displace Cβ(262) by 0.47-0.78Å (Fig. 5d). By contrast, Leu262 in
F262L is in a preferred trans/gauche+ conformation at χ1/χ2 = 177°/63° as is Val262 in
F262V at χ1 = −176°. What might control activation of HiTehA is unclear, but for AtSLAC1
activation is by OST1 phosphorylation6,7. The molecular consequences of OST1
phosphorylation of SLAC1 remain unknown, but it is plausible that associated shifts in pore-
helix orientations would unlatch Phe450 in SLAC1 from a TehA-like restrained orientation.
By analogy with Leu262 in F262L, we expect a preferred rotameric state for Leu450 in
AtSLAC1 F450L. Thus, the lack of appreciable OST1 activation of conductance in
AtSLAC1 F450L (Fig. 5e) might be explained by the lack of a restraining latch, whereby the
channel remains closed despite OST1 activation. Puzzles certainly remain since OST1 does
substantially activate AtSLAC1 F450T or F450V, which like HiTehA F262V should also be
unrestrained; presumably activating adjustments widen the pore enough for ion permeation
past threonine and valine but not leucine.
Phosphorylation sites have been discovered in the N- and C-terminal tails of AtSLAC16,7,28
(179 and 51 residues long, respectively), but these alone cannot explain OST1 activation of
SLAC1. First SLAH1, which fully complements the slac1-1 mutation, does not have these
cytoplasmic tails. Secondly, although OST1 phosphorylation of Ser120 in the N-terminal tail
is necessary for SLAC1 activation, it is not sufficient7. Thus, we surmise that direct
phosphorylation of the SLAC1 TM domain must be critical, and SLAC1 has four conserved
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Ser/Thr candidates in its cytoplasmic loops. Moreover, SLAC1 proteins have proline-
mediated kinks at the putative Phe450 gate in helix TM9, and also in adjacent helix TM7;
these features may play a role in phosphorylation-driven unlatching of the Phe450 gate in
SLAC1.
Ion selectivity and discrimination
Our studies of SLAC1 channel relative ion permeabilities, based on measurements of current
reversal potential, are consistent with earlier work demonstrating that AtSLAC1 conducts
anions but not cations and is selective among anions, with greater permeability for nitrate
than for chloride (as in Vicia faba guard cell protoplasts27) and much reduced permeability
for malate, bicarbonate or sulfate6,7. We also find that SLAC1 has little permeability for
sulfite. Additionally, we find that wild-type SLAC1, F450A and F450T all have similar
relative permeabilities to chloride, sulfite and malate, despite having widely different
conductance levels, but the gating mutants do show small but significant decreases in nitrate
permeability (Fig. 4c, Table S6).
The relative insensitivity of anion permeability to gating residue changes suggests that
selectivity for these anions may occur away from the central constriction at the channel gate.
To some extent, ionic discrimination must depend on pore geometry; thus, an organic anion
such as malate may be simply too large to pass through the 5-Å wide pore. Although the
SLAC1 pore is lined largely with hydrophobic sidechains (Fig. 3c), it is sprinkled with
hydroxyl groups from serine and threonine residues (16%); their electropositive hydrogen
atoms may facilitate conductance. Most strikingly, the electrostatic potential within the
AtSLAC1 pore is electropositive throughout (Fig. 3a). This polarization, promoted by
charges on extra-membranous loops, no doubt contributes significantly in discrimination
against cations.
The relative anion permeability sequence of SLAC1 determined by us and others, I− > NO3−
> Br− > Cl− 6,7, corresponds to selectivity sequence 1 compiled by Wright and Diamond29
for a range of anion-selective proteins. This sequence correlates inversely with the hydration
energies of monovalent anions – anions with a lower hydration energy have a greater
channel permeability. It is thought to be generated in proteins with weak, low field-strength,
anion binding sites, where selectivity is largely determined by the energetic cost of anion
dehydration. These selectivity results are thus consistent with the SLAC1 structure, where
the pore lacks any obvious anion binding site.
Distinctiveness of the SLAC1 channel
SLAC1 anion channels are entirely novel in structure and, apparently, in the mechanism for
ion conductance. The best characterized of anion channels belong to the CLC family of Cl−
channels and transporters30-32. CLC channels have an altogether different architecture from
the SLAC1 channel, and the mechanism for selectivity is also very different. Bacterial CLC
transporters bind halide ions at three sites in a highly constricted pore30. By contrast, the
SLAC1 pore has a relatively uniform diameter across the membrane, except where closed by
the gating phenyl group, and we do not find discrete ion binding sites. CLC selectivity is
governed by specific residues surrounding these binding sites30,32. The anion selectivity
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sequence for the CLC channels of Cl−>Br−>NO −3>I−, opposite of that in SLAC1, is
consistent with the high field-strength anion binding sites in CLC channels29. Interestingly,
as for AtSLAC1, an Arabidopsis CLCa channel also preferentially transports nitrate ions33,
and an E. coli CLC channel is converted to preference of nitrate when a generally conserved
serine at the central site is substituted with proline as in AtCLCa32.
SLAC1 also differs radically from other structurally characterized anion channels and
transporters. These include the VDAC1 voltage-gated anion channel from mitochondrial
outer membranes, which has a porin-like beta-barrel structure34-36, and a light-driven
halorhodopsin chloride pump, which has a transmembrane conductance pathway similar to
that of the proton-pumping pore of bacteriorhodopsin37. Although its channel structure is
still only known by homology to other ABC transporters, CFTR is another obviously
distinct chloride channel38. Cys-loop receptors also include anion channels39, and these are
similar to SLAC1 in having five-helix pores25, but here selectivity is governed by charged
groups at the entrance to the pore, which distinguish the anion-selective GABAA and
glycine receptors from the cation-selective acetylcholine and serotonin 5HT3 receptors39.
Finally, the recently identified TMEM16A gene for calcium-activated chloride channels40-42
appears to encode an 8-TM protein that is again distinct from SLAC1.
Stomatal guard cells show both rapidly activated (R-type) and slow (S-type) anion channel
activity43. Although slac1 guard cells have very defective S-type activity, their R-type
currents are normal4. Guard cell protoplasts from the slac1-2 mutant abnormally accumulate
Cl−, K+, malate and fumarate3, whereas SLAC1 shows negligible malate conductance7. As
for SLAC1-associated K+ movements, other channels or transporters must be responsible for
SLAC1-associated malate movements. Recent studies indicate that AtALMT12, an
aluminium-activated malate transporter (ALMT) family member, is a malate-dependent R-
type anion channel44 needed for stomatal closure45.
Conclusions
We find that many functional properties of the plant SLAC1 anion channel are explained
well by the structure of an uncharacterized bacterial TehA protein that has been associated
with tellurite resistance. SLAC1 and TehA belong to distinct subfamilies within one branch
of a larger SLAC1 superfamily, but AtSLAC1 and HiTehA are sufficiently similar (19%
sequence identity) that the SLAC1 homology model is predictive for function, including a
verified placement of the identifying slac1-2 mutation G194D and a phenylalanine gate. One
remaining puzzle concerns the structural change that activating phosphorylation elicits in
SLAC1, and another puzzle is the biochemical role of the TehA homologs in bacteria. In a
companion paper26, we examine functional and structural properties of TehA in bacteria,
showing that it is anion channel, although actually not conferring tellurite resistance, and
identifying a mutant variant with properties suggestive of an activated state. Thus, SLAC1
and TehA likely represent a large family of selective anion channels controlled by
environmental stimuli.
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METHODS
Selection of target sequences
TehA from E. coli (EcTehA) was a centrally selected sequence for expansion as a
NYCOMPS targeted family. Target selection criteria utilized at NYCOMPS are described in
details elsewhere46. Briefly, the EcTehA sequence was run against a dataset of about 40,000
predicted alpha helical integral membrane protein sequences from prokaryotic genomes
(NYCOMPS98 dataset46) using PSI-BLAST50. Sequences that matched EcTehA with an E-
value lower than 10−3 in an alignment extending over at least 50% of both predicted TM
regions and passing our post-seed-expansion filtering criteria46 were passed to the protein
production pipeline.
Protein expression screening
Full-length homologs from the following 38 species, including 2 sequences each from 5 of
these, were amplified from genomic DNA by PCR: Thermoplasma acidophilum,
Lactococcus lactis subsp. lactis Il1403, Streptococcus pyogenes M1 GAS, Streptococcus
pneumoniae TIGR4, Haemophilus influenzae Rd KW20, Methanocaldococcus jannaschii
DSM 2661 (2), Escherichia coli K12, Salmonella typhimurium LT2, Clostridium
perfringens ATCC 13124, Xanthomonas campestris pv. campestris str. ATCC 33913,
Streptococcus agalactiae 2603V/R, Shewanella oneidensis MR-1, Streptococcus mutans
UA159, Archaeoglobus fulgidus DSM 4304, Vibrio parahaemolyticus RIMD 2210633 (2),
Pseudomonas syringae pv. tomato str. DC3000 (2), Enterococcus faecalis V583,
Pyrococcus horikoshii OT3, Bordetella bronchiseptica RB50, Streptomyces coelicolor A3,
Corynebacterium glutamicum ATCC 13032, Picrophilus torridus DSM 9790, Acinetobacter
sp. ADP1, Vibrio fischeri ES114, Pseudomonas fluorescens Pf-5, Sulfolobus acidocaldarius
DSM 639, Colwellia psychrerythraea 34H, Neisseria meningitidis MC58, Rhodobacter
sphaeroides 2.4.1, Vibrio cholerae O1 biovar eltor str. N16961 (2), Marinobacter aquaeolei
VT8, Acinetobacter baumannii ATCC 17978, Klebsiella pneumoniae subsp. pneumoniae
MGH 78578, Streptomyces avermitili MA-4680, Bordetella parapertussis 12822 (2),
Streptococcus thermophilus LMG 18311, Salmonella enterica subsp. enterica serovar
Paratyphi A str. ATCC 9150, and Anaeromyxobacter dehalogenans 2CP-C.
Selected cDNAs were cloned into a modified pET vector (Novagen, Inc) that fuses a FLAG
and deca-histidine tag at the C-terminus, which are cleavable by TEV protease. Proteins
were expressed in E. coli BL21(DE3) plysS by a high throughput format (0.6ml in a deep
well block) and purified after lysis by sonication using metal affinity purification in a buffer
containing N-Dodecyl-β-D-Maltopyranoside. Samples were passed over an analytical size
exclusion column in 12 different detergent-containing mobile phases, which included N-
dodecyl-β-D-maltopyranoside (DDM), N-decyl-β-D-altopyranoside (DM), N-nonyl-β-D-
altopyranoside (NM), N-octyl-β-D-altopyranoside (OM), N-octyl-β-D-Glucopyranoside
(OG), N-nonyl-β-D-Glucopyranoside (NG) and lauryl dimethyl amine oxide (LDAO).
Multi-angle light scattering with refractive index detection was used to analyze the
oligomeric state51. The E. coli and H. influenzae proteins were judged to be monodisperse
and stable and were passed to scale up.
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Scaled-up production and purification
For scale up, in brief, transformed BL21 plysS cells were grown at 37°C in 2× YT media to
OD 0.6-0.8 after being inoculated with 1% of the overnight culture. The culture was induced
with 0.4mM IPTG and continued to grow at 37 °C for another 4 hours. The cells were
harvested by centrifugation and stored at −80°C before use. Selenomethionyl (SeMet) TehA
was expressed in a similar way, but using containing SeMet in place of methionine in
defined minimal media. Cells were resuspended in a buffer containing 50 mM Tris-HCl (pH
8.0) and 200 mM NaCl and lysed using a French Press with two passes at 15-20,000 psi.
Cell debris were removed by centrifugation at 10,000g for 20min, and the membrane
fraction was isolated from that supernatant by ultra-centrifugation at 150,000g for 1 hr.
The membrane fraction was homogenized in a solubilization buffer containing 50 mM Tris
(pH 8.0) and 200 mM NaCl, and incubated with final concentration of 1% (w/v) dodecyl-β-
D-maltopyranoside (DDM, Anatrace, Inc) for 1 hr at 4°C. The non-dissolved matter was
remove by ultracentrifugation at 150,000g for 30 min, and the supernatant was loaded to a
5ml Hitrap Ni2+-NTA affinity column (GE Healthcare), pre-equilibrated with the same
solubilization buffer supplemented with 0.05% DDM. After 20 column volume buffer wash,
the protein was eluted with 250mM imidazole in the solublization buffer. The Flag and 10-
His tags were removed by adding super TEV at 1:100 mass ratio and incubating at 4°C
overnight. Tag removal was confirmed by SDS-PAGE, and the resulting sample was
concentrated to around 10mg/ml. Preparative size exclusion chromatography was carried out
on a Superdex-200 column for further purification, removal of TEV protease and the
cleaved tag, and for buffer and detergent exchange. The gel-filtration buffer contained 10
mM Tris (pH 8.0), 200 mM NaCl, 1mM EDTA, 0.5 mM Tris [2-carboxyethyl] phosphine
(TCEP), and 2×CMC of detergent. In the case of HiTehA, the protein was well behaved and
stable in nearly all tested detergents, and we have purified it from DDM, DM, NM, OM, OG
and LDAO.
Protein characterization
We performed N-terminal amino-acid sequencing of purified HiTehA and EcTehA prior to
TEV protease treatment. Results from these analyses proved that true initiating methionine
residue is located 14 residues after one annotated as N-terminal. The intervening nucleotide
sequence contains a Shine-Delgarno sequence.
For cross-linking experiments, purified HiTehA was incubated with 10 mM disuccinimidyl
glutarate at room temperate for 30 min, and 100mM Tris-HCl pH 7.5 was added to stop the
reaction. The incubated sample when run on a 8-25% gradient SDS-PAGE gel showed a
ladder consistent with a trimeric structure.
Crystallization and data collection
Purified protein was concentrated to ~10mg/ml for initial crystal trials in a Mosquito robot
with commercial screens from Hampton research, Emerald Biosystems and Molecular
Dimension. We obtained crystals of HiTehA from protein in detergent DDM, DM, NM,
OM, OG and LDAO, but only those from LDAO and OG gave diffraction to beyond 4Å
spacings. Crystals that proved useful were all grown at 4°C using the sitting-drop vapor
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diffusion method. After extensive optimization we reached conditions supporting very high
resolution. Wild-type HiTehA and most of its variants crystallized from 1mM ZnSO4,
50mM Hepes-Na pH 7.8 and 28% PEG600. HiTehA F262A crystallized from 200mM
Li2SO4, 100mM glycine pH9.3 and 33% PEG400). Addition of 10mM spermidine as an
additive helped to obtain slightly better diffracting crystals. Cryoprotection was achieved by
adding 5% ethylene glycol or PEG400 to the crystallization solution.
Structure determination and refinement
Native and SeMet SAD data set were collected at NSLS beamline X4A and processed using
the software HKL200052. Crystals of HiTehA grew in space group R3 with a=b=c = 85.0Å
and α=β=γ = 93.5°. All subsequent manipulations were done in the hexagonal setting of this
space group with a=b= 96.0Å and c = 136.7Å. The asymmetric unit contains one TehA
protomer and 65% solvent volume. The structure was determined at 2.0Å resolution from
single-wavelength anomalous diffraction (SAD) using selenomethionine-substituted protein
crystals. Assessment of data quality for phasing, location of heavy atom sites and initial
phases were calculated using the HKL2MAP interface to SHELX programs53.
All the secondary structure elements were clearly visible in the experimental electron
density map. Automatic model building was done in Arp/wArp54 and completed manually
in the program COOT55. The model was refined against native data at 1.20Å resolution
using the program Refmac5.5 in CCP456, with anisotropic B factor restrained refinement
applied. Subsequent structural analyses of mutant variants were refined as isomorphous
structures.
Site-directed mutagenesis
Site-directed mutants were constructed using the QuikChange Site-Directed Mutagenesis Kit
(Stratagene, La Jolla, CA 92037) and expressed from pET vectors in E. coli BL21(DE3)
plysS cells as for the wild-type protein.
Electrophysiology
All constructs were cloned into plasmid pGHME2, linearized, and transcribed into cRNA
using T7 polymerase (mMessage mMachine, Ambion). Oocytes were injected with 50 nl of
cRNA solution each, at a constant concentration of 0.5 mg/ml for HiTehA, or 0.5 mg/ml for
AtSLAC1 constructs, with or without 0.5 mg/ml of AtOST1 constructs and recorded in
voltage-clamp experiments 2 days after injection. For mixed expression experiments, 25nl of
cRNA solution was injected for each AtSLAC1 component. Two-electrode voltage clamp
recordings were performed to measure HiTehA or AtSLAC1 currents as described6,7. The
pipette solutions contained 3 M KCl. For voltage-clamp current recordings, the bath solution
contained 1 mM MgCl2, 1 mM CaCl2, 10 mM Mes/Tris (pH 5.6) and 30 mM CsCl; For
anion selectivity measurements, the bath solutions contained 50 mM Cl−, NO −3, malate, or
30 mM SO 2-3 sodium salts, 45 mM Na-gluconate, 1 mM Ca-gluconate2, 1 mM Mg-
gluconate2, 1 mM K-gluconate, as well as 10 mM Tris/Mes (pH 5.6). Osmolality was
adjusted with D-mannitol to 220 mOsmol/kg. The bath electrode was a 3M KCl agar bridge.
The testing potentials were +50 to −110 or −130 mV in 20 mV decrements with 7.5 s
duration. The holding and return potentials were 0 mV with a 1.45 s or 2 s, respectively. I-V
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relations of HiTehA or AtSLAC1 were generated from the current reached after 0.5 s in
testing potentials. The Goldman-Hodgkin-Katz equation was applied to estimate
permeability ratios for monovalent ions as described6. For divalent anions, the permeability
ratios were derived according to Fatt and Ginsborg57.
Bioinformatic analysis of SLAC-related proteins
Sequences related to SLAC1 were analyzed comprehensively by PSI-BLAST50. Searches at
E<10−3 starting from five disparate homologs each identified a common pool of over 900
proteins, which when pooled were used for sub-classification into families and subfamilies.
Details of these analyses are reported in footnotes to Table S1.
Molecular figures were produced in PyMOL58.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We thank Filippo Mancia, Larry Shapiro and Ming Zhou for helpful discussions; Patricia Rodriguez and Arianne
Morrison for help with cloning and expression; Jonah Cheung, Mark Collins, Chris Min, Sheng Ye and Zhen Zhang
for advice in protein chemistry and crystallography; John Schwanof and Randy Abramowitz for help with
synchrotron experiments; John Riley for help with Xenopus oocyte injections; and Min Su for advice on the PSI-
BLAST analysis of SLAC relatives. This project was supported in part by an award to the New York Consortium
on Membrane Protein Structure (NYCOMPS) from the NIGMS Protein Structure Initiative. Beamline X4A at the
National Synchrotron Light Source (NSLS), a DOE facility at Brookhaven National Laboratory, is supported by the
New York Structural Biology Center.
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Figure 1. Sequence analysis for the SLAC1 superfamily
a, Family tree. The presentation was computed by the program COBALT47 from
representative subfamily sequences (Table S1), including Arabidopsis thaliana SLAC1 for
SF1Ai, Haemophilus influenzae TehA for SF1Ci, Escherichia coli TehA for SF1Cii, Vibrio
parahaemolyticus for SF2A, Staphylococcus aureus for SF2B, Aspergillus fumigatus Ssu1
for SF3A, and Schizosaccharomyces pombe Mae1 for SF3B. b, Structure-based sequence
alignment of TehA from Haemophilus influenzae (HiTehA) and SLAC1 from Arabidopsis
thaliana (AtSLAC1). The TehA structure has been used to restrict sequence gaps to inter-
helical segments. Superior coils define extents of the HiTehA helical segments; red letters
mark residue identities; red boxes are drawn for residues that are >95% identical within the
plant subfamily SF1A for AtSLAC1 or within the TehA subfamily for HiTehA; red
diamonds mark HiTehA residues that line the central pore; and the colored inferior bar
encodes ConSurf sequence variability48 for the SF1 family of 204 non-redundant proteins.
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Figure 2. Crystal structure of HiTehA and homology model of AtSLAC1
a, Electron density distribution from the HiTehA crystal structure at 1.2Å resolution. The
map has (2Fo-Fc) coefficients based on the superimposed model. Contours are at 2.5σ. b,
Ribbon diagram of the HiTehA trimer. Each protomer is colored spectrally from blue at its
N-terminus to red at its C-terminus. c, DelPhi49 electrostatic potential at the extracellular
surface. Electronegative and electropositive potential are colored in degrees of red and blue
saturation, respectively. d, Electrostatic potential49 at the intracellular surface. e, Ribbon
diagram of an HiTehA protomer viewed from outside the membrane. The ribbon is colored
spectrally as in 2b. f, Ribbon diagram of an HiTehA protomer viewed from within the
membrane from below the view of e. g, Surface of a homology model of AtSLAC1, viewed
as in f, and colored by sequence variability48. h, Surface of AtSLAC1 as in g, but colored by
electrostatic potential49.
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Figure 3. Putative structure of the SLAC1 conductance pore
a, Cross-section through the homology model of AtSLAC1. The model is viewed as in 2i,
with the electrostatic potential49 shown on the external surface of the molecular envelope.
The side chain of Phe450 is shown as a stick model (red) on the backbone ribbon colored
yellow. b, Cross-section as in a, but colored by surface conservation48 as in 2h. c, Pore-
lining residues in the SLAC1 homology model. A cylinder model (left), spectrally colored as
in 2b, provides a key for viewing the rolled-open structure (left) with pore-lining residues of
AtSLAC1 shown on the TModd helices. d, Ribbon diagrams of HiTehA TM9 (left) and TM7
(right) viewed from within the conductance pore. The side chains of Pro207 and Phe262 are
shown as well as a kink-stabilizing water HOH25 that is coordinated by the NH of Gly263
and by C=O groups of Gly202 and Ala259. Density contours are shown for the water
molecule.
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Figure 4. Ionic conductance measurements
a, Typical microelectrode voltage-clamp current traces from oocytes injected with various
channel cRNAs. Left column, oocytes injected with cRNAs encoding wild-type HiTehA
channels (WT), or F262A, G15D/F262A or G15D mutants. Right two columns, oocytes
injected with cRNAs for WT AtSLAC1, or F450A, G194D/F450A or G194D mutants, with
or without co-injection of AtOST1. Dotted lines represent zero current levels. Extracellular
solution contained 30mM CsCl. Schematic keys at the left place phenyl and/or aspartyl
gates. b, Effects of gating residue mutations. Mean chloride currents, measured at −90 mV,
are shown comparing WT HiTehA and its mutant series F262A,G,T,V,L with WT AtSLAC1
and its corresponding series F450A,G,T,V,L, both alone and co-expressed with AtOST1.
Full I-V relations are shown in Fig. S5. c, Effect of gating residues on relative AtSLAC1
anion permeabilities. Relative permeabilities (P[X]/P[Cl]) for chloride, nitrate, sulfite and
malate of WT, F450A and F450T SLAC1 channels were measured from the change in
current reversal potential with Cl− or anion X− as the sole permeant anion in the bath
solution (Methods, Table S6).
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Figure 5. Structural features at the SLAC1 homolog gate
a,b, Cross-sections through the conductance pores of HiTehA F262A and HiTehA G15D.
The view and presentations are as in 3a, except that helices are colored purple. c, Molecular
basis for conformational strain in gating residue Phe262 of HiTehA. Helices TM1 (left),
TM9 (center) and TM7 (right) are viewed from within the pore and presented as ribbon
diagrams with selected side chains drawn in stick representation. The local low-energy
conformation for the phenyl ring (χ2 = 90°) is shown in thin lines with short contacts
indicated in dashed lines: Leu18 Cδ2 – Phe262 Cδ1, d = 2.4 Å; Val210 Cγ2 – Phe262 Cε2, d
= 2.8 Å. d, Conformational shifts consequent to release of strain in gating residue Phe262.
Cα backbone structures of F262A, F262V and F262L HiTehA were first superimposed onto
WT HiTehA, and residues 258-266 were drawn in stereo as superimposed. All backbone
atoms are shown for peptides 262±1, but only Cα atoms are shown otherwise. The WT
backbone and phenyl group are green; other backbone are all magenta; side chains of
Ala262, Val262 and Leu262 are cyan, blue and red, respectively; oxygen-directed bonds are
red and nitrogen-directed bonds are blue. e, Typical microelectrode voltage-clamp current
traces from oocytes injected with wild-type (WT) or F450L AtSLAC1 cRNA. Experimental
conditions and displays are as in 4a.
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|
3M74
|
Crystal Structure of Plant SLAC1 homolog TehA
|
Homolog Structure of the SLAC1 Anion Channel for Closing
Stomata in Leaves
Yuhang Chen1,6, Lei Hu2, Marco Punta6,7, Renato Bruni6, Brandan Hillerich6, Brian Kloss6,
Burkhard Rost1,6,1, James Love6, Steven A. Siegelbaum2,3,5, and Wayne A.
Hendrickson1,4,5,6
1Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY
10032, USA
2Department of Neuroscience, Columbia University, New York, NY 10032, USA
3Department of Pharmacology, Columbia University, New York, NY 10032, USA
4Department of Physiology and Cellular Biophysics, Columbia University, New York, NY 10032,
USA
5Howard Hughes Medical Institute, Columbia University, New York, NY 10032, USA
6NYCOMPS, New York Structural Biology Center 89 Convent Avenue, New York, NY 10027 USA
7Department of Computer Science and Institute for Advanced Study Technical University of
Munich D-85748 Munich, Germany
Summary
The plant SLAC1 anion channel controls turgor pressure in the aperture-defining guard cells of
plant stomata, thereby regulating exchange of water vapor and photosynthetic gases in response to
environmental signals such as drought or high levels of carbon dioxide. We determined the crystal
structure of a bacterial homolog of SLAC1 at 1.20Å resolution, and we have used structure-
inspired mutagenesis to analyze the conductance properties of SLAC1 channels. SLAC1 is a
symmetric trimer composed from quasi-symmetric subunits, each having ten transmembrane
helices arranged from helical hairpin pairs to form a central five-helix transmembrane pore that is
gated by an extremely conserved phenylalanine residue. Conformational features suggest a
mechanism for control of gating by kinase activation, and electrostatic features of the pore coupled
with electrophysiological characteristics suggest that selectivity among different anions is largely
a function of the energetic cost of ion dehydration.
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Correspondence and request for materials should be addressed to WAH (wayne@convex.hhmi.columbia.edu)..
Supplementary Information is linked to the online version of the paper at www.nature.com/nature.
Author Contributions YC, LH, MP, RB, BH, BK and JL performed experiments; YC, LH, BR, SAS, and WAH analyzed data; YC,
LH, SAS, and WAH prepared the manuscript.
Author Information Atomic coordinates and diffraction data are deposited in the Protein Data Bank with accession codes 3M71,
3M73, 3M74, 3M75, 3M76, and 3M7L. See Table S4 for identifications. Reprints and permissions information is available at
www.Nature.com/reprints.
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Published in final edited form as:
Nature. 2010 October 28; 467(7319): 1074–1080. doi:10.1038/nature09487.
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Stomatal pores in the leaves of plants permit the influx of atmospheric carbon dioxide in
exchange for transpirational evaporation of water1,2. A pair of kidney-shaped guard cells
define each pore aperture, and turgor pressure variation in these cells determines the degree
of stomatal pore openness. Depending on diverse environmental factors, the stomata close to
prevent H2O losses and open to admit CO2 for photosynthesis. Environmental stimuli that
lead to stomatal closure include darkness, high CO2 levels, ozone, low air humidity and
drought. The plant hormone abscisic acid (ABA) is critical for signal transduction from
these stimuli. Mutational screens in Arabidopsis thaliana for CO2 and ozone sensitivity
identified a protein with ten predicted transmembrane (TM) helices, now called slow anion
channel 1 (SLAC1), as having a central role in the control of stomatal closure3-5. Recent
studies proved that SLAC1 is indeed an anion channel6-7, with characteristics like those of
slow anion channels found in guard cells8, and that it is activated by phosphorylation from
the OST1 kinase9. OST1 activity is negatively regulated by the ABI1 phosphatase10,11,
which is in turn inhibited by the stomatal ABA receptors PYR/RCAR12,13 when in the
ternary hormone-receptor-phosphatase complex14-18. Thereby, ABA stimulates SLAC1
channel activity. Resulting Cl− efflux through SLAC1 causes membrane depolarization,
which activates outward-rectifying K+ channels, leading to KCl and water efflux to further
reduce turgor and cause stomatal closure.
SLAC1 expression in Arabidopsis is confined to the guard cells of leaves, but other
Arabidopsis tissues do have SLAC1 homologs3, named SLAH1-SLAH4. The identifying
mutations slac1-14 and slac1-23 are, respectively, in predicted transmembrane segments 9
(S456F) and 1 (G194D) of a protein that includes substantial N- and C-terminal extensions
outside a 10-helix transmembrane (TM) domain. SLAH1, which is absent from leaves and
lacks the terminal extensions of SLAC1, fully complements the mutant phenotype in slac1-2
guard cell protoplasts3. SLAC1 and homologs are also present in other plant genomes,
including nine in rice (Oryza sativa) and five in grape vines (Vitis vinifera). SLAC1
relatives, some quite remote, also occur in bacteria, archaea and fungi. Known prokaryotic
homologs contain only the predicted transmembrane domain of SLAC1, but some fungal
homologs do have N- and C-terminal extensions. One homolog, Mae1 from the yeast
Schizosaccharomyces pombe, functions as a malate uptake transporter19; another, Ssu1 from
Saccharomyces cerevisiae and other fungi including Aspergillus fumigatus, is characterized
as a sulfite efflux pump20,21; and TehA from Escherichia coli is identified as a tellurite
resistance protein by virtue of its association in the tehA/tehB operon22,23. Despite a lack of
further biochemical characterization, many homologs are annotated as tellurite resistance/
dicarboxylate transporter (TDT) proteins.
We have undertaken structural and functional characterizations of the SLAC1 anion
channel. We first solved an atomic-resolution crystal structure of the TehA homolog from
Haemophilus influenzae, and we then developed a homology model for Arabidopsis SLAC1.
This model allowed us to conduct mutagenesis for functional testing of structure-inspired
hypotheses on gating and selectivity. We expressed Haemophilus TehA and Arabidopsis
SLAC1 in Xenopus oocytes to characterize channel properties of these proteins and mutant
variants. We also determined crystal structures for several mutant variants, including the
homolog of slac1-2.
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Structure of SLAC1 bacterial homolog TehA
We performed a bioinformatic analysis of SLAC1-related proteins, first clustering nearly
900 non-redundant sequences into a superfamily at the PSI-BLAST level E≤10−3, then into
three distinct families at an initial threshold of E≤10−30, and finally into subfamilies at a
typical initial threshold of E≤10−55. Since previous annotation is not well founded in
experiment and SLAC1 is now the best characterized member, we adopt a nomenclature
defining a SLAC superfamily divided into families identified as SF1-SF3 and subfamilies
SF1A, SF1B etc. Family SF1 comprises the plant SLAC proteins and close bacterial
homologs, family SF2 comprises a distinct set of bacterial proteins often annotated as
exfoliative toxins, and family SF3 comprises the fungal Mae1 and Ssu1 proteins and their
archaeal or bacterial homologs, respectively. SLAC family SF1 has three large subfamilies:
the plant SLAC and SLAH proteins are in subfamily SF1A, closest bacterial homologs are
in SF1B, and the TehA homologs are in SF1C (Fig. 1a). The other families also divide into
subfamilies as detailed in Table S1, and family SF1 is divided into sub-subfamilies (Table
S2). Two pertinent SF1 sequences are aligned in Fig. 1b.
We used a structural genomics approach to obtain structural information, testing expression
and purification for 43 bacterial and archaeal likely homologs, assaying for detergent choice
and stability on 8 of these, finding two with appropriate profiles by size exclusion
chromatography, and obtaining suitable crystals for one. This protein, TehA from H.
influenzae (HiTehA), was found to be trimeric both by size-exclusion multi-angle light
scattering (SEC-MALS) measurements and by chemical cross-linking. When solubilized in
β-octylglucoside, HiTehA crystallized in space group R3 with a=b=96.01Å and c=136.27Å.
Each asymmetric unit contains one subunit and 65% solvent. The structure was solved by
selenomethionyl (SeMet) SAD phasing, ultimately at 1.50Å resolution (Table S3, Fig. S1),
and then refined at 1.20Å resolution (Fig. 2a) to R/Rfree values of 14.1%/15.7% for a model
that includes ordered residues 6-313, 213 water molecules and four detergent molecules
(Table S4).
The crystal structure has TehA trimers aligned with three-fold axes of the lattice (Fig. 2b).
Subunits are tightly associated, burying 8947 Å2 of total surface area within trimer
interfaces. The electrostatic potential surface is largely negative on the extracellular surface
(Fig. 2c) and largely positive on the cytoplasmic surface (Fig. 2d). The membrane
orientation is specified experimentally from GFP tagging of E. coli TehA24. Each TehA
protomer has ten TM helices, as predicted; however, the fold is novel. Tandemly repeated
helical hairpins are arranged with quasi-five-fold symmetry (Figs. 2e,S2). Extracellular
inter-helix loops are short (2-5 residues) whereas intracellular inter-helix connections are
longer, including a nine-residue helix H2,3 between TM2 and TM3 (Fig. 2f). An inner pentad
of outwardly directed, TModd, helices creates an apparent pore through each protomer
perpendicular to the putative membrane plane. TMeven helices from the five hairpins
surround the inner pore and make an outer layer.
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Homology model for plant SLAC1
Arabidopsis SLAC1 (AtSLAC1) is substantially similar to bacterial homologs, notably
HiTehA (Fig. 1b). All HiTehA TM helices are fully aligned to predicted SLAC1 TM
helices, but there are short inter-helical gaps (1-5 residues) in all five extracellular loops and
in two of the intracellular loops. The TM domain of AtSLAC1 (residues 188-504) aligns to
HiTehA with 19% sequence identity and with a PSI-BLAST E-value of 3×10−22. For
comparison, and in keeping with the family tree (Fig. 1a), the TM domain of AtSLAC1
shares sequence identities of 76% with rice SLAC1, 41% with Arabidopsis SLAH1, 25%
with an SL1B homolog from Halorhodospira halophila, 11% with S. cerevisiae Ssu1, and
9% with S. pombe Mae1. A conceptual model with the AtSLAC1 sequence transposed onto
the HiTehA helices sufficed to guide most of our mutational tests, but a detailed AtSLAC1
homology model helped to refine our ideas. Surface variability and electrostatic potential are
plotted onto the surface of this model (Fig. 2g,2h).
The most remarkable feature of the TehA structure and corresponding SLAC1 model is the
central pore through each protomer. As for acetylcholine receptors25, the SLAC1 pore is
formed by five helices; but the SLAC1 helices come from one protein molecule rather than
five. The SLAC1 pore has a relatively uniform diameter of approximately 5Å across nearly
five helical turns (Fig. S3), except for a pronounced constriction in the middle of the
membrane (Fig. 3a) where the pore is occluded by the side chain of Phe 450 (Phe 262 in
HiTehA). This residue is the only absolutely conserved amino acid residue of the SLAC1
family. The pore is lined with highly conserved (86% identity among five SLAC1 orthologs;
32% identity between AtSLAC1 and HiTehA) and generally hydrophobic residues (Figs. 1b,
3b,3c,S4). Despite this hydrophobicity, the electrostatic potential on the pore surface is
polarized (Fig. 3a), presumably due to an invaginated shape adjacent to charged residues
outside the membrane. The generally electropositive character of the cytoplasmic surface
likely contributes to anion efflux.
Kinks in the pore helices contribute to formation of a relatively constant pore diameter
across the membrane. Four of the five HiTehA inner helices have centrally located proline
residues, which necessarily generate kinks, and TM9 is kinked at a backbone-coordinated
water molecule (Fig. 3d). Proline replaces Gly263 in all SF1A and SF1B relatives, including
Pro451 AtSLAC1. This water-displacing change is isostructural26. Centrally located prolines
also prevail in TM3, TM5 and TM7 across the SF1 proteins. The outer helices are longer and
straighter, but more inclined. The only outer-helix proline kink of HiTehA is in TM6 at the
trimer three-fold axis.
Two of the gene-identifying mutations in Arabadopsis SLAC1 are selected point mutations,
others are disruptive transfer DNA (T-DNA) insertions4. In the AtSLAC1 homology model,
the slac1-2 (G194D) mutation3 points into the pore from TM1 and can be accommodated
structurally whereas the slac1-1 (S456F) mutation4 points away from the pore six residues
after pore-blocking Phe450 on TM9 and would be expected to disruptive. Ser456 interacts
with outer helix TM10 in the homology model, and the phenyl bulk from S456F would not
fit. Position 456 has alanine in HiTehA, 66% of all 204 SF1 homologs and another 27%
have threonine or serine (as in all SLAC1 channels); phenylalanine never occurs among all
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814 SLAC superfamily members. Position 194 has glycine in 58% of SF1 homologs and
alanine in another 22%. Asp194, which never occurs naturally, would block the pore and be
expected to repel anions.
Mutational tests of channel function
Mutational studies corroborate the hypothesis that a TehA-based SLAC1 model is
appropriate. First, as discussed above, the slac1-1 mutant (S456F) is expected to be
structurally disruptive, and indeed it is inactive in guard cells4, and the slac1-2 mutation
(G194D) is expected to block the pore, and we show below that this variant is also inactive.
We have also shown that the introduction of SLAC1-conserved proline residues into
HiTehA (A208P/G263P) is accommodated isomorphously26. Moreover, as shown below,
channel conductance properties of several mutants are similar for AtSLAC1 and HiTehA.
To examine characteristics of the SLAC1 channel in light of the structural model, we
performed electrophysiological tests of membrane currents from voltage-clamped Xenopus
oocytes following injection of wild-type or mutant AtSLAC1 or HiTehA cRNAs. We
observed modest-sized chloride currents with wild-type AtSLAC1 cRNA, as found
previously6,7, but did not detect any chloride current following injection of wild-type
HiTehA cRNA. We found that SLAC1 Cl− conductance was enhanced when the OST1
kinase cRNA was co-injected with SLAC1 cRNA, but only to the levels found by Lee et al.6
and not to the much higher levels found by Geiger et al.7 with OST1 physically connected to
SLAC1 by split YFP linkage. In keeping with structural evidence that Phe262 blocks the
HiTehA pore, removal of the phenyl group in HiTehA F262A or in the homologous
AtSLAC1 F450A resulted in very large Cl− currents relative to wild-type levels, and the
SLAC1 currents were now less enhanced by presence of OST1 (Fig. 4a). The tempting
interpretation of an opened gate will require validation with appropriately analyzed single-
channel recordings27. In keeping with the slac1-2 phenotype3, neither the functionally
impaired AtSLAC1 G194D nor its HiTehA G15D homolog showed any substantial
conductance; moreover, further consistent with pore blockage by Asp194 in slac1-2, the
large conductances of HiTehA F262A and AtSLAC1 F450A were stopped in the double
mutants AtSLAC1 G194D/F450A and HiTehA G15D/F262A (Fig. 4a). Here again, the
effects in SLAC1 were independent of OST1.
We also tested conductance characteristics for a series of AtSLAC1 F450X substitution
mutants – F450A,G,T,V,L– and for the corresponding HiTehA F262X series –
F262A,G,T,V,L (Figs. 4b,S5, Table S5). Findings from the two series are roughly parallel;
in particular, the alanine and glycine substitutions lead to large currents for both and in
comparison to the others. There are distinctions, of course, including generally higher
conductances for AtSLAC1 over HiTehA and less conductance of F262T TehA compared to
F450T SLAC1. It is also noteworthy that OST1 activation is very muted for the F450A,G,L
mutants, which is consistent with SLAC1 gating at Phe450.
Crystal structures were also determined for several of the HiTehA mutant variants (Table
S4). The structures of F262A (1.15Å), F262V (1.60Å), F262L (1.65Å) and G15D (1.50Å)
are all essentially isomorphous with the wild-type TehA structure with changes localized
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primarily at the sites of mutation; the same is true for the double mutations of F262A/G15D,
F262G/G15D and A208P/G263P26. The F262A structure has a wide open pore (Fig. 5a)
with a relatively uniform pore diameter of ~5Å through ~30Å across the membrane (Fig.
S3) whereas G15D has a doubly occluded pore (Fig. 5b). The pores of other mutant variants
are consistent with the sizes of constrictive residues and with the observed conductances.
Gating and activation
The crystal structures of TehA and its mutant variants when taken together with the
functional studies in Xenopus oocytes point to a crucial role for Phe450 in gating of the
SLAC1 anion channel. Conservation of this residue across the SLAC1 family implies
functional importance. The occlusion of the pore by the presence of F262 in the structure of
wild-type TehA and the openness of the pore upon its substitution by alanine in the structure
of the F262A mutant provides physical evidence for a gating role of this residue. This
interpretation is supported by the correlated conductance characteristics from variants of the
AtSLAC1 and HiTehA channels (Figs. 4b,S5). While these observations may suffice for
placing the gate within the channel pore, they do not by themselves suggest a mechanism for
gating in response to physiological stimuli. Some insight does come from conformational
details defined at high resolution.
One important structural clue is that the side chain of Phe262 is in a high energy
conformation in the HiTehA structure, with χ1/χ2 at −160°/-4°. Although χ1 is in a preferred
trans conformation, the phenyl ring is restricted by contacts with Val210 and Leu18 to a χ2
value near 0° rather than near to the preferred 90° orientation (Fig. 5c). Further evidence that
Phe262 is restrained from local equilibrium comes from shifts observed in crystal structures
of the F262A, F262V, F262L and F262G/G15D variants, which all show consistent
backbone movements that displace Cβ(262) by 0.47-0.78Å (Fig. 5d). By contrast, Leu262 in
F262L is in a preferred trans/gauche+ conformation at χ1/χ2 = 177°/63° as is Val262 in
F262V at χ1 = −176°. What might control activation of HiTehA is unclear, but for AtSLAC1
activation is by OST1 phosphorylation6,7. The molecular consequences of OST1
phosphorylation of SLAC1 remain unknown, but it is plausible that associated shifts in pore-
helix orientations would unlatch Phe450 in SLAC1 from a TehA-like restrained orientation.
By analogy with Leu262 in F262L, we expect a preferred rotameric state for Leu450 in
AtSLAC1 F450L. Thus, the lack of appreciable OST1 activation of conductance in
AtSLAC1 F450L (Fig. 5e) might be explained by the lack of a restraining latch, whereby the
channel remains closed despite OST1 activation. Puzzles certainly remain since OST1 does
substantially activate AtSLAC1 F450T or F450V, which like HiTehA F262V should also be
unrestrained; presumably activating adjustments widen the pore enough for ion permeation
past threonine and valine but not leucine.
Phosphorylation sites have been discovered in the N- and C-terminal tails of AtSLAC16,7,28
(179 and 51 residues long, respectively), but these alone cannot explain OST1 activation of
SLAC1. First SLAH1, which fully complements the slac1-1 mutation, does not have these
cytoplasmic tails. Secondly, although OST1 phosphorylation of Ser120 in the N-terminal tail
is necessary for SLAC1 activation, it is not sufficient7. Thus, we surmise that direct
phosphorylation of the SLAC1 TM domain must be critical, and SLAC1 has four conserved
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Ser/Thr candidates in its cytoplasmic loops. Moreover, SLAC1 proteins have proline-
mediated kinks at the putative Phe450 gate in helix TM9, and also in adjacent helix TM7;
these features may play a role in phosphorylation-driven unlatching of the Phe450 gate in
SLAC1.
Ion selectivity and discrimination
Our studies of SLAC1 channel relative ion permeabilities, based on measurements of current
reversal potential, are consistent with earlier work demonstrating that AtSLAC1 conducts
anions but not cations and is selective among anions, with greater permeability for nitrate
than for chloride (as in Vicia faba guard cell protoplasts27) and much reduced permeability
for malate, bicarbonate or sulfate6,7. We also find that SLAC1 has little permeability for
sulfite. Additionally, we find that wild-type SLAC1, F450A and F450T all have similar
relative permeabilities to chloride, sulfite and malate, despite having widely different
conductance levels, but the gating mutants do show small but significant decreases in nitrate
permeability (Fig. 4c, Table S6).
The relative insensitivity of anion permeability to gating residue changes suggests that
selectivity for these anions may occur away from the central constriction at the channel gate.
To some extent, ionic discrimination must depend on pore geometry; thus, an organic anion
such as malate may be simply too large to pass through the 5-Å wide pore. Although the
SLAC1 pore is lined largely with hydrophobic sidechains (Fig. 3c), it is sprinkled with
hydroxyl groups from serine and threonine residues (16%); their electropositive hydrogen
atoms may facilitate conductance. Most strikingly, the electrostatic potential within the
AtSLAC1 pore is electropositive throughout (Fig. 3a). This polarization, promoted by
charges on extra-membranous loops, no doubt contributes significantly in discrimination
against cations.
The relative anion permeability sequence of SLAC1 determined by us and others, I− > NO3−
> Br− > Cl− 6,7, corresponds to selectivity sequence 1 compiled by Wright and Diamond29
for a range of anion-selective proteins. This sequence correlates inversely with the hydration
energies of monovalent anions – anions with a lower hydration energy have a greater
channel permeability. It is thought to be generated in proteins with weak, low field-strength,
anion binding sites, where selectivity is largely determined by the energetic cost of anion
dehydration. These selectivity results are thus consistent with the SLAC1 structure, where
the pore lacks any obvious anion binding site.
Distinctiveness of the SLAC1 channel
SLAC1 anion channels are entirely novel in structure and, apparently, in the mechanism for
ion conductance. The best characterized of anion channels belong to the CLC family of Cl−
channels and transporters30-32. CLC channels have an altogether different architecture from
the SLAC1 channel, and the mechanism for selectivity is also very different. Bacterial CLC
transporters bind halide ions at three sites in a highly constricted pore30. By contrast, the
SLAC1 pore has a relatively uniform diameter across the membrane, except where closed by
the gating phenyl group, and we do not find discrete ion binding sites. CLC selectivity is
governed by specific residues surrounding these binding sites30,32. The anion selectivity
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sequence for the CLC channels of Cl−>Br−>NO −3>I−, opposite of that in SLAC1, is
consistent with the high field-strength anion binding sites in CLC channels29. Interestingly,
as for AtSLAC1, an Arabidopsis CLCa channel also preferentially transports nitrate ions33,
and an E. coli CLC channel is converted to preference of nitrate when a generally conserved
serine at the central site is substituted with proline as in AtCLCa32.
SLAC1 also differs radically from other structurally characterized anion channels and
transporters. These include the VDAC1 voltage-gated anion channel from mitochondrial
outer membranes, which has a porin-like beta-barrel structure34-36, and a light-driven
halorhodopsin chloride pump, which has a transmembrane conductance pathway similar to
that of the proton-pumping pore of bacteriorhodopsin37. Although its channel structure is
still only known by homology to other ABC transporters, CFTR is another obviously
distinct chloride channel38. Cys-loop receptors also include anion channels39, and these are
similar to SLAC1 in having five-helix pores25, but here selectivity is governed by charged
groups at the entrance to the pore, which distinguish the anion-selective GABAA and
glycine receptors from the cation-selective acetylcholine and serotonin 5HT3 receptors39.
Finally, the recently identified TMEM16A gene for calcium-activated chloride channels40-42
appears to encode an 8-TM protein that is again distinct from SLAC1.
Stomatal guard cells show both rapidly activated (R-type) and slow (S-type) anion channel
activity43. Although slac1 guard cells have very defective S-type activity, their R-type
currents are normal4. Guard cell protoplasts from the slac1-2 mutant abnormally accumulate
Cl−, K+, malate and fumarate3, whereas SLAC1 shows negligible malate conductance7. As
for SLAC1-associated K+ movements, other channels or transporters must be responsible for
SLAC1-associated malate movements. Recent studies indicate that AtALMT12, an
aluminium-activated malate transporter (ALMT) family member, is a malate-dependent R-
type anion channel44 needed for stomatal closure45.
Conclusions
We find that many functional properties of the plant SLAC1 anion channel are explained
well by the structure of an uncharacterized bacterial TehA protein that has been associated
with tellurite resistance. SLAC1 and TehA belong to distinct subfamilies within one branch
of a larger SLAC1 superfamily, but AtSLAC1 and HiTehA are sufficiently similar (19%
sequence identity) that the SLAC1 homology model is predictive for function, including a
verified placement of the identifying slac1-2 mutation G194D and a phenylalanine gate. One
remaining puzzle concerns the structural change that activating phosphorylation elicits in
SLAC1, and another puzzle is the biochemical role of the TehA homologs in bacteria. In a
companion paper26, we examine functional and structural properties of TehA in bacteria,
showing that it is anion channel, although actually not conferring tellurite resistance, and
identifying a mutant variant with properties suggestive of an activated state. Thus, SLAC1
and TehA likely represent a large family of selective anion channels controlled by
environmental stimuli.
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METHODS
Selection of target sequences
TehA from E. coli (EcTehA) was a centrally selected sequence for expansion as a
NYCOMPS targeted family. Target selection criteria utilized at NYCOMPS are described in
details elsewhere46. Briefly, the EcTehA sequence was run against a dataset of about 40,000
predicted alpha helical integral membrane protein sequences from prokaryotic genomes
(NYCOMPS98 dataset46) using PSI-BLAST50. Sequences that matched EcTehA with an E-
value lower than 10−3 in an alignment extending over at least 50% of both predicted TM
regions and passing our post-seed-expansion filtering criteria46 were passed to the protein
production pipeline.
Protein expression screening
Full-length homologs from the following 38 species, including 2 sequences each from 5 of
these, were amplified from genomic DNA by PCR: Thermoplasma acidophilum,
Lactococcus lactis subsp. lactis Il1403, Streptococcus pyogenes M1 GAS, Streptococcus
pneumoniae TIGR4, Haemophilus influenzae Rd KW20, Methanocaldococcus jannaschii
DSM 2661 (2), Escherichia coli K12, Salmonella typhimurium LT2, Clostridium
perfringens ATCC 13124, Xanthomonas campestris pv. campestris str. ATCC 33913,
Streptococcus agalactiae 2603V/R, Shewanella oneidensis MR-1, Streptococcus mutans
UA159, Archaeoglobus fulgidus DSM 4304, Vibrio parahaemolyticus RIMD 2210633 (2),
Pseudomonas syringae pv. tomato str. DC3000 (2), Enterococcus faecalis V583,
Pyrococcus horikoshii OT3, Bordetella bronchiseptica RB50, Streptomyces coelicolor A3,
Corynebacterium glutamicum ATCC 13032, Picrophilus torridus DSM 9790, Acinetobacter
sp. ADP1, Vibrio fischeri ES114, Pseudomonas fluorescens Pf-5, Sulfolobus acidocaldarius
DSM 639, Colwellia psychrerythraea 34H, Neisseria meningitidis MC58, Rhodobacter
sphaeroides 2.4.1, Vibrio cholerae O1 biovar eltor str. N16961 (2), Marinobacter aquaeolei
VT8, Acinetobacter baumannii ATCC 17978, Klebsiella pneumoniae subsp. pneumoniae
MGH 78578, Streptomyces avermitili MA-4680, Bordetella parapertussis 12822 (2),
Streptococcus thermophilus LMG 18311, Salmonella enterica subsp. enterica serovar
Paratyphi A str. ATCC 9150, and Anaeromyxobacter dehalogenans 2CP-C.
Selected cDNAs were cloned into a modified pET vector (Novagen, Inc) that fuses a FLAG
and deca-histidine tag at the C-terminus, which are cleavable by TEV protease. Proteins
were expressed in E. coli BL21(DE3) plysS by a high throughput format (0.6ml in a deep
well block) and purified after lysis by sonication using metal affinity purification in a buffer
containing N-Dodecyl-β-D-Maltopyranoside. Samples were passed over an analytical size
exclusion column in 12 different detergent-containing mobile phases, which included N-
dodecyl-β-D-maltopyranoside (DDM), N-decyl-β-D-altopyranoside (DM), N-nonyl-β-D-
altopyranoside (NM), N-octyl-β-D-altopyranoside (OM), N-octyl-β-D-Glucopyranoside
(OG), N-nonyl-β-D-Glucopyranoside (NG) and lauryl dimethyl amine oxide (LDAO).
Multi-angle light scattering with refractive index detection was used to analyze the
oligomeric state51. The E. coli and H. influenzae proteins were judged to be monodisperse
and stable and were passed to scale up.
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Scaled-up production and purification
For scale up, in brief, transformed BL21 plysS cells were grown at 37°C in 2× YT media to
OD 0.6-0.8 after being inoculated with 1% of the overnight culture. The culture was induced
with 0.4mM IPTG and continued to grow at 37 °C for another 4 hours. The cells were
harvested by centrifugation and stored at −80°C before use. Selenomethionyl (SeMet) TehA
was expressed in a similar way, but using containing SeMet in place of methionine in
defined minimal media. Cells were resuspended in a buffer containing 50 mM Tris-HCl (pH
8.0) and 200 mM NaCl and lysed using a French Press with two passes at 15-20,000 psi.
Cell debris were removed by centrifugation at 10,000g for 20min, and the membrane
fraction was isolated from that supernatant by ultra-centrifugation at 150,000g for 1 hr.
The membrane fraction was homogenized in a solubilization buffer containing 50 mM Tris
(pH 8.0) and 200 mM NaCl, and incubated with final concentration of 1% (w/v) dodecyl-β-
D-maltopyranoside (DDM, Anatrace, Inc) for 1 hr at 4°C. The non-dissolved matter was
remove by ultracentrifugation at 150,000g for 30 min, and the supernatant was loaded to a
5ml Hitrap Ni2+-NTA affinity column (GE Healthcare), pre-equilibrated with the same
solubilization buffer supplemented with 0.05% DDM. After 20 column volume buffer wash,
the protein was eluted with 250mM imidazole in the solublization buffer. The Flag and 10-
His tags were removed by adding super TEV at 1:100 mass ratio and incubating at 4°C
overnight. Tag removal was confirmed by SDS-PAGE, and the resulting sample was
concentrated to around 10mg/ml. Preparative size exclusion chromatography was carried out
on a Superdex-200 column for further purification, removal of TEV protease and the
cleaved tag, and for buffer and detergent exchange. The gel-filtration buffer contained 10
mM Tris (pH 8.0), 200 mM NaCl, 1mM EDTA, 0.5 mM Tris [2-carboxyethyl] phosphine
(TCEP), and 2×CMC of detergent. In the case of HiTehA, the protein was well behaved and
stable in nearly all tested detergents, and we have purified it from DDM, DM, NM, OM, OG
and LDAO.
Protein characterization
We performed N-terminal amino-acid sequencing of purified HiTehA and EcTehA prior to
TEV protease treatment. Results from these analyses proved that true initiating methionine
residue is located 14 residues after one annotated as N-terminal. The intervening nucleotide
sequence contains a Shine-Delgarno sequence.
For cross-linking experiments, purified HiTehA was incubated with 10 mM disuccinimidyl
glutarate at room temperate for 30 min, and 100mM Tris-HCl pH 7.5 was added to stop the
reaction. The incubated sample when run on a 8-25% gradient SDS-PAGE gel showed a
ladder consistent with a trimeric structure.
Crystallization and data collection
Purified protein was concentrated to ~10mg/ml for initial crystal trials in a Mosquito robot
with commercial screens from Hampton research, Emerald Biosystems and Molecular
Dimension. We obtained crystals of HiTehA from protein in detergent DDM, DM, NM,
OM, OG and LDAO, but only those from LDAO and OG gave diffraction to beyond 4Å
spacings. Crystals that proved useful were all grown at 4°C using the sitting-drop vapor
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diffusion method. After extensive optimization we reached conditions supporting very high
resolution. Wild-type HiTehA and most of its variants crystallized from 1mM ZnSO4,
50mM Hepes-Na pH 7.8 and 28% PEG600. HiTehA F262A crystallized from 200mM
Li2SO4, 100mM glycine pH9.3 and 33% PEG400). Addition of 10mM spermidine as an
additive helped to obtain slightly better diffracting crystals. Cryoprotection was achieved by
adding 5% ethylene glycol or PEG400 to the crystallization solution.
Structure determination and refinement
Native and SeMet SAD data set were collected at NSLS beamline X4A and processed using
the software HKL200052. Crystals of HiTehA grew in space group R3 with a=b=c = 85.0Å
and α=β=γ = 93.5°. All subsequent manipulations were done in the hexagonal setting of this
space group with a=b= 96.0Å and c = 136.7Å. The asymmetric unit contains one TehA
protomer and 65% solvent volume. The structure was determined at 2.0Å resolution from
single-wavelength anomalous diffraction (SAD) using selenomethionine-substituted protein
crystals. Assessment of data quality for phasing, location of heavy atom sites and initial
phases were calculated using the HKL2MAP interface to SHELX programs53.
All the secondary structure elements were clearly visible in the experimental electron
density map. Automatic model building was done in Arp/wArp54 and completed manually
in the program COOT55. The model was refined against native data at 1.20Å resolution
using the program Refmac5.5 in CCP456, with anisotropic B factor restrained refinement
applied. Subsequent structural analyses of mutant variants were refined as isomorphous
structures.
Site-directed mutagenesis
Site-directed mutants were constructed using the QuikChange Site-Directed Mutagenesis Kit
(Stratagene, La Jolla, CA 92037) and expressed from pET vectors in E. coli BL21(DE3)
plysS cells as for the wild-type protein.
Electrophysiology
All constructs were cloned into plasmid pGHME2, linearized, and transcribed into cRNA
using T7 polymerase (mMessage mMachine, Ambion). Oocytes were injected with 50 nl of
cRNA solution each, at a constant concentration of 0.5 mg/ml for HiTehA, or 0.5 mg/ml for
AtSLAC1 constructs, with or without 0.5 mg/ml of AtOST1 constructs and recorded in
voltage-clamp experiments 2 days after injection. For mixed expression experiments, 25nl of
cRNA solution was injected for each AtSLAC1 component. Two-electrode voltage clamp
recordings were performed to measure HiTehA or AtSLAC1 currents as described6,7. The
pipette solutions contained 3 M KCl. For voltage-clamp current recordings, the bath solution
contained 1 mM MgCl2, 1 mM CaCl2, 10 mM Mes/Tris (pH 5.6) and 30 mM CsCl; For
anion selectivity measurements, the bath solutions contained 50 mM Cl−, NO −3, malate, or
30 mM SO 2-3 sodium salts, 45 mM Na-gluconate, 1 mM Ca-gluconate2, 1 mM Mg-
gluconate2, 1 mM K-gluconate, as well as 10 mM Tris/Mes (pH 5.6). Osmolality was
adjusted with D-mannitol to 220 mOsmol/kg. The bath electrode was a 3M KCl agar bridge.
The testing potentials were +50 to −110 or −130 mV in 20 mV decrements with 7.5 s
duration. The holding and return potentials were 0 mV with a 1.45 s or 2 s, respectively. I-V
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relations of HiTehA or AtSLAC1 were generated from the current reached after 0.5 s in
testing potentials. The Goldman-Hodgkin-Katz equation was applied to estimate
permeability ratios for monovalent ions as described6. For divalent anions, the permeability
ratios were derived according to Fatt and Ginsborg57.
Bioinformatic analysis of SLAC-related proteins
Sequences related to SLAC1 were analyzed comprehensively by PSI-BLAST50. Searches at
E<10−3 starting from five disparate homologs each identified a common pool of over 900
proteins, which when pooled were used for sub-classification into families and subfamilies.
Details of these analyses are reported in footnotes to Table S1.
Molecular figures were produced in PyMOL58.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We thank Filippo Mancia, Larry Shapiro and Ming Zhou for helpful discussions; Patricia Rodriguez and Arianne
Morrison for help with cloning and expression; Jonah Cheung, Mark Collins, Chris Min, Sheng Ye and Zhen Zhang
for advice in protein chemistry and crystallography; John Schwanof and Randy Abramowitz for help with
synchrotron experiments; John Riley for help with Xenopus oocyte injections; and Min Su for advice on the PSI-
BLAST analysis of SLAC relatives. This project was supported in part by an award to the New York Consortium
on Membrane Protein Structure (NYCOMPS) from the NIGMS Protein Structure Initiative. Beamline X4A at the
National Synchrotron Light Source (NSLS), a DOE facility at Brookhaven National Laboratory, is supported by the
New York Structural Biology Center.
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Figure 1. Sequence analysis for the SLAC1 superfamily
a, Family tree. The presentation was computed by the program COBALT47 from
representative subfamily sequences (Table S1), including Arabidopsis thaliana SLAC1 for
SF1Ai, Haemophilus influenzae TehA for SF1Ci, Escherichia coli TehA for SF1Cii, Vibrio
parahaemolyticus for SF2A, Staphylococcus aureus for SF2B, Aspergillus fumigatus Ssu1
for SF3A, and Schizosaccharomyces pombe Mae1 for SF3B. b, Structure-based sequence
alignment of TehA from Haemophilus influenzae (HiTehA) and SLAC1 from Arabidopsis
thaliana (AtSLAC1). The TehA structure has been used to restrict sequence gaps to inter-
helical segments. Superior coils define extents of the HiTehA helical segments; red letters
mark residue identities; red boxes are drawn for residues that are >95% identical within the
plant subfamily SF1A for AtSLAC1 or within the TehA subfamily for HiTehA; red
diamonds mark HiTehA residues that line the central pore; and the colored inferior bar
encodes ConSurf sequence variability48 for the SF1 family of 204 non-redundant proteins.
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Figure 2. Crystal structure of HiTehA and homology model of AtSLAC1
a, Electron density distribution from the HiTehA crystal structure at 1.2Å resolution. The
map has (2Fo-Fc) coefficients based on the superimposed model. Contours are at 2.5σ. b,
Ribbon diagram of the HiTehA trimer. Each protomer is colored spectrally from blue at its
N-terminus to red at its C-terminus. c, DelPhi49 electrostatic potential at the extracellular
surface. Electronegative and electropositive potential are colored in degrees of red and blue
saturation, respectively. d, Electrostatic potential49 at the intracellular surface. e, Ribbon
diagram of an HiTehA protomer viewed from outside the membrane. The ribbon is colored
spectrally as in 2b. f, Ribbon diagram of an HiTehA protomer viewed from within the
membrane from below the view of e. g, Surface of a homology model of AtSLAC1, viewed
as in f, and colored by sequence variability48. h, Surface of AtSLAC1 as in g, but colored by
electrostatic potential49.
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Figure 3. Putative structure of the SLAC1 conductance pore
a, Cross-section through the homology model of AtSLAC1. The model is viewed as in 2i,
with the electrostatic potential49 shown on the external surface of the molecular envelope.
The side chain of Phe450 is shown as a stick model (red) on the backbone ribbon colored
yellow. b, Cross-section as in a, but colored by surface conservation48 as in 2h. c, Pore-
lining residues in the SLAC1 homology model. A cylinder model (left), spectrally colored as
in 2b, provides a key for viewing the rolled-open structure (left) with pore-lining residues of
AtSLAC1 shown on the TModd helices. d, Ribbon diagrams of HiTehA TM9 (left) and TM7
(right) viewed from within the conductance pore. The side chains of Pro207 and Phe262 are
shown as well as a kink-stabilizing water HOH25 that is coordinated by the NH of Gly263
and by C=O groups of Gly202 and Ala259. Density contours are shown for the water
molecule.
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Figure 4. Ionic conductance measurements
a, Typical microelectrode voltage-clamp current traces from oocytes injected with various
channel cRNAs. Left column, oocytes injected with cRNAs encoding wild-type HiTehA
channels (WT), or F262A, G15D/F262A or G15D mutants. Right two columns, oocytes
injected with cRNAs for WT AtSLAC1, or F450A, G194D/F450A or G194D mutants, with
or without co-injection of AtOST1. Dotted lines represent zero current levels. Extracellular
solution contained 30mM CsCl. Schematic keys at the left place phenyl and/or aspartyl
gates. b, Effects of gating residue mutations. Mean chloride currents, measured at −90 mV,
are shown comparing WT HiTehA and its mutant series F262A,G,T,V,L with WT AtSLAC1
and its corresponding series F450A,G,T,V,L, both alone and co-expressed with AtOST1.
Full I-V relations are shown in Fig. S5. c, Effect of gating residues on relative AtSLAC1
anion permeabilities. Relative permeabilities (P[X]/P[Cl]) for chloride, nitrate, sulfite and
malate of WT, F450A and F450T SLAC1 channels were measured from the change in
current reversal potential with Cl− or anion X− as the sole permeant anion in the bath
solution (Methods, Table S6).
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Figure 5. Structural features at the SLAC1 homolog gate
a,b, Cross-sections through the conductance pores of HiTehA F262A and HiTehA G15D.
The view and presentations are as in 3a, except that helices are colored purple. c, Molecular
basis for conformational strain in gating residue Phe262 of HiTehA. Helices TM1 (left),
TM9 (center) and TM7 (right) are viewed from within the pore and presented as ribbon
diagrams with selected side chains drawn in stick representation. The local low-energy
conformation for the phenyl ring (χ2 = 90°) is shown in thin lines with short contacts
indicated in dashed lines: Leu18 Cδ2 – Phe262 Cδ1, d = 2.4 Å; Val210 Cγ2 – Phe262 Cε2, d
= 2.8 Å. d, Conformational shifts consequent to release of strain in gating residue Phe262.
Cα backbone structures of F262A, F262V and F262L HiTehA were first superimposed onto
WT HiTehA, and residues 258-266 were drawn in stereo as superimposed. All backbone
atoms are shown for peptides 262±1, but only Cα atoms are shown otherwise. The WT
backbone and phenyl group are green; other backbone are all magenta; side chains of
Ala262, Val262 and Leu262 are cyan, blue and red, respectively; oxygen-directed bonds are
red and nitrogen-directed bonds are blue. e, Typical microelectrode voltage-clamp current
traces from oocytes injected with wild-type (WT) or F450L AtSLAC1 cRNA. Experimental
conditions and displays are as in 4a.
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|
3M75
|
Crystal Structure of Plant SLAC1 homolog TehA
|
Homolog Structure of the SLAC1 Anion Channel for Closing
Stomata in Leaves
Yuhang Chen1,6, Lei Hu2, Marco Punta6,7, Renato Bruni6, Brandan Hillerich6, Brian Kloss6,
Burkhard Rost1,6,1, James Love6, Steven A. Siegelbaum2,3,5, and Wayne A.
Hendrickson1,4,5,6
1Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY
10032, USA
2Department of Neuroscience, Columbia University, New York, NY 10032, USA
3Department of Pharmacology, Columbia University, New York, NY 10032, USA
4Department of Physiology and Cellular Biophysics, Columbia University, New York, NY 10032,
USA
5Howard Hughes Medical Institute, Columbia University, New York, NY 10032, USA
6NYCOMPS, New York Structural Biology Center 89 Convent Avenue, New York, NY 10027 USA
7Department of Computer Science and Institute for Advanced Study Technical University of
Munich D-85748 Munich, Germany
Summary
The plant SLAC1 anion channel controls turgor pressure in the aperture-defining guard cells of
plant stomata, thereby regulating exchange of water vapor and photosynthetic gases in response to
environmental signals such as drought or high levels of carbon dioxide. We determined the crystal
structure of a bacterial homolog of SLAC1 at 1.20Å resolution, and we have used structure-
inspired mutagenesis to analyze the conductance properties of SLAC1 channels. SLAC1 is a
symmetric trimer composed from quasi-symmetric subunits, each having ten transmembrane
helices arranged from helical hairpin pairs to form a central five-helix transmembrane pore that is
gated by an extremely conserved phenylalanine residue. Conformational features suggest a
mechanism for control of gating by kinase activation, and electrostatic features of the pore coupled
with electrophysiological characteristics suggest that selectivity among different anions is largely
a function of the energetic cost of ion dehydration.
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Correspondence and request for materials should be addressed to WAH (wayne@convex.hhmi.columbia.edu)..
Supplementary Information is linked to the online version of the paper at www.nature.com/nature.
Author Contributions YC, LH, MP, RB, BH, BK and JL performed experiments; YC, LH, BR, SAS, and WAH analyzed data; YC,
LH, SAS, and WAH prepared the manuscript.
Author Information Atomic coordinates and diffraction data are deposited in the Protein Data Bank with accession codes 3M71,
3M73, 3M74, 3M75, 3M76, and 3M7L. See Table S4 for identifications. Reprints and permissions information is available at
www.Nature.com/reprints.
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Nature. 2010 October 28; 467(7319): 1074–1080. doi:10.1038/nature09487.
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Stomatal pores in the leaves of plants permit the influx of atmospheric carbon dioxide in
exchange for transpirational evaporation of water1,2. A pair of kidney-shaped guard cells
define each pore aperture, and turgor pressure variation in these cells determines the degree
of stomatal pore openness. Depending on diverse environmental factors, the stomata close to
prevent H2O losses and open to admit CO2 for photosynthesis. Environmental stimuli that
lead to stomatal closure include darkness, high CO2 levels, ozone, low air humidity and
drought. The plant hormone abscisic acid (ABA) is critical for signal transduction from
these stimuli. Mutational screens in Arabidopsis thaliana for CO2 and ozone sensitivity
identified a protein with ten predicted transmembrane (TM) helices, now called slow anion
channel 1 (SLAC1), as having a central role in the control of stomatal closure3-5. Recent
studies proved that SLAC1 is indeed an anion channel6-7, with characteristics like those of
slow anion channels found in guard cells8, and that it is activated by phosphorylation from
the OST1 kinase9. OST1 activity is negatively regulated by the ABI1 phosphatase10,11,
which is in turn inhibited by the stomatal ABA receptors PYR/RCAR12,13 when in the
ternary hormone-receptor-phosphatase complex14-18. Thereby, ABA stimulates SLAC1
channel activity. Resulting Cl− efflux through SLAC1 causes membrane depolarization,
which activates outward-rectifying K+ channels, leading to KCl and water efflux to further
reduce turgor and cause stomatal closure.
SLAC1 expression in Arabidopsis is confined to the guard cells of leaves, but other
Arabidopsis tissues do have SLAC1 homologs3, named SLAH1-SLAH4. The identifying
mutations slac1-14 and slac1-23 are, respectively, in predicted transmembrane segments 9
(S456F) and 1 (G194D) of a protein that includes substantial N- and C-terminal extensions
outside a 10-helix transmembrane (TM) domain. SLAH1, which is absent from leaves and
lacks the terminal extensions of SLAC1, fully complements the mutant phenotype in slac1-2
guard cell protoplasts3. SLAC1 and homologs are also present in other plant genomes,
including nine in rice (Oryza sativa) and five in grape vines (Vitis vinifera). SLAC1
relatives, some quite remote, also occur in bacteria, archaea and fungi. Known prokaryotic
homologs contain only the predicted transmembrane domain of SLAC1, but some fungal
homologs do have N- and C-terminal extensions. One homolog, Mae1 from the yeast
Schizosaccharomyces pombe, functions as a malate uptake transporter19; another, Ssu1 from
Saccharomyces cerevisiae and other fungi including Aspergillus fumigatus, is characterized
as a sulfite efflux pump20,21; and TehA from Escherichia coli is identified as a tellurite
resistance protein by virtue of its association in the tehA/tehB operon22,23. Despite a lack of
further biochemical characterization, many homologs are annotated as tellurite resistance/
dicarboxylate transporter (TDT) proteins.
We have undertaken structural and functional characterizations of the SLAC1 anion
channel. We first solved an atomic-resolution crystal structure of the TehA homolog from
Haemophilus influenzae, and we then developed a homology model for Arabidopsis SLAC1.
This model allowed us to conduct mutagenesis for functional testing of structure-inspired
hypotheses on gating and selectivity. We expressed Haemophilus TehA and Arabidopsis
SLAC1 in Xenopus oocytes to characterize channel properties of these proteins and mutant
variants. We also determined crystal structures for several mutant variants, including the
homolog of slac1-2.
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Structure of SLAC1 bacterial homolog TehA
We performed a bioinformatic analysis of SLAC1-related proteins, first clustering nearly
900 non-redundant sequences into a superfamily at the PSI-BLAST level E≤10−3, then into
three distinct families at an initial threshold of E≤10−30, and finally into subfamilies at a
typical initial threshold of E≤10−55. Since previous annotation is not well founded in
experiment and SLAC1 is now the best characterized member, we adopt a nomenclature
defining a SLAC superfamily divided into families identified as SF1-SF3 and subfamilies
SF1A, SF1B etc. Family SF1 comprises the plant SLAC proteins and close bacterial
homologs, family SF2 comprises a distinct set of bacterial proteins often annotated as
exfoliative toxins, and family SF3 comprises the fungal Mae1 and Ssu1 proteins and their
archaeal or bacterial homologs, respectively. SLAC family SF1 has three large subfamilies:
the plant SLAC and SLAH proteins are in subfamily SF1A, closest bacterial homologs are
in SF1B, and the TehA homologs are in SF1C (Fig. 1a). The other families also divide into
subfamilies as detailed in Table S1, and family SF1 is divided into sub-subfamilies (Table
S2). Two pertinent SF1 sequences are aligned in Fig. 1b.
We used a structural genomics approach to obtain structural information, testing expression
and purification for 43 bacterial and archaeal likely homologs, assaying for detergent choice
and stability on 8 of these, finding two with appropriate profiles by size exclusion
chromatography, and obtaining suitable crystals for one. This protein, TehA from H.
influenzae (HiTehA), was found to be trimeric both by size-exclusion multi-angle light
scattering (SEC-MALS) measurements and by chemical cross-linking. When solubilized in
β-octylglucoside, HiTehA crystallized in space group R3 with a=b=96.01Å and c=136.27Å.
Each asymmetric unit contains one subunit and 65% solvent. The structure was solved by
selenomethionyl (SeMet) SAD phasing, ultimately at 1.50Å resolution (Table S3, Fig. S1),
and then refined at 1.20Å resolution (Fig. 2a) to R/Rfree values of 14.1%/15.7% for a model
that includes ordered residues 6-313, 213 water molecules and four detergent molecules
(Table S4).
The crystal structure has TehA trimers aligned with three-fold axes of the lattice (Fig. 2b).
Subunits are tightly associated, burying 8947 Å2 of total surface area within trimer
interfaces. The electrostatic potential surface is largely negative on the extracellular surface
(Fig. 2c) and largely positive on the cytoplasmic surface (Fig. 2d). The membrane
orientation is specified experimentally from GFP tagging of E. coli TehA24. Each TehA
protomer has ten TM helices, as predicted; however, the fold is novel. Tandemly repeated
helical hairpins are arranged with quasi-five-fold symmetry (Figs. 2e,S2). Extracellular
inter-helix loops are short (2-5 residues) whereas intracellular inter-helix connections are
longer, including a nine-residue helix H2,3 between TM2 and TM3 (Fig. 2f). An inner pentad
of outwardly directed, TModd, helices creates an apparent pore through each protomer
perpendicular to the putative membrane plane. TMeven helices from the five hairpins
surround the inner pore and make an outer layer.
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Homology model for plant SLAC1
Arabidopsis SLAC1 (AtSLAC1) is substantially similar to bacterial homologs, notably
HiTehA (Fig. 1b). All HiTehA TM helices are fully aligned to predicted SLAC1 TM
helices, but there are short inter-helical gaps (1-5 residues) in all five extracellular loops and
in two of the intracellular loops. The TM domain of AtSLAC1 (residues 188-504) aligns to
HiTehA with 19% sequence identity and with a PSI-BLAST E-value of 3×10−22. For
comparison, and in keeping with the family tree (Fig. 1a), the TM domain of AtSLAC1
shares sequence identities of 76% with rice SLAC1, 41% with Arabidopsis SLAH1, 25%
with an SL1B homolog from Halorhodospira halophila, 11% with S. cerevisiae Ssu1, and
9% with S. pombe Mae1. A conceptual model with the AtSLAC1 sequence transposed onto
the HiTehA helices sufficed to guide most of our mutational tests, but a detailed AtSLAC1
homology model helped to refine our ideas. Surface variability and electrostatic potential are
plotted onto the surface of this model (Fig. 2g,2h).
The most remarkable feature of the TehA structure and corresponding SLAC1 model is the
central pore through each protomer. As for acetylcholine receptors25, the SLAC1 pore is
formed by five helices; but the SLAC1 helices come from one protein molecule rather than
five. The SLAC1 pore has a relatively uniform diameter of approximately 5Å across nearly
five helical turns (Fig. S3), except for a pronounced constriction in the middle of the
membrane (Fig. 3a) where the pore is occluded by the side chain of Phe 450 (Phe 262 in
HiTehA). This residue is the only absolutely conserved amino acid residue of the SLAC1
family. The pore is lined with highly conserved (86% identity among five SLAC1 orthologs;
32% identity between AtSLAC1 and HiTehA) and generally hydrophobic residues (Figs. 1b,
3b,3c,S4). Despite this hydrophobicity, the electrostatic potential on the pore surface is
polarized (Fig. 3a), presumably due to an invaginated shape adjacent to charged residues
outside the membrane. The generally electropositive character of the cytoplasmic surface
likely contributes to anion efflux.
Kinks in the pore helices contribute to formation of a relatively constant pore diameter
across the membrane. Four of the five HiTehA inner helices have centrally located proline
residues, which necessarily generate kinks, and TM9 is kinked at a backbone-coordinated
water molecule (Fig. 3d). Proline replaces Gly263 in all SF1A and SF1B relatives, including
Pro451 AtSLAC1. This water-displacing change is isostructural26. Centrally located prolines
also prevail in TM3, TM5 and TM7 across the SF1 proteins. The outer helices are longer and
straighter, but more inclined. The only outer-helix proline kink of HiTehA is in TM6 at the
trimer three-fold axis.
Two of the gene-identifying mutations in Arabadopsis SLAC1 are selected point mutations,
others are disruptive transfer DNA (T-DNA) insertions4. In the AtSLAC1 homology model,
the slac1-2 (G194D) mutation3 points into the pore from TM1 and can be accommodated
structurally whereas the slac1-1 (S456F) mutation4 points away from the pore six residues
after pore-blocking Phe450 on TM9 and would be expected to disruptive. Ser456 interacts
with outer helix TM10 in the homology model, and the phenyl bulk from S456F would not
fit. Position 456 has alanine in HiTehA, 66% of all 204 SF1 homologs and another 27%
have threonine or serine (as in all SLAC1 channels); phenylalanine never occurs among all
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814 SLAC superfamily members. Position 194 has glycine in 58% of SF1 homologs and
alanine in another 22%. Asp194, which never occurs naturally, would block the pore and be
expected to repel anions.
Mutational tests of channel function
Mutational studies corroborate the hypothesis that a TehA-based SLAC1 model is
appropriate. First, as discussed above, the slac1-1 mutant (S456F) is expected to be
structurally disruptive, and indeed it is inactive in guard cells4, and the slac1-2 mutation
(G194D) is expected to block the pore, and we show below that this variant is also inactive.
We have also shown that the introduction of SLAC1-conserved proline residues into
HiTehA (A208P/G263P) is accommodated isomorphously26. Moreover, as shown below,
channel conductance properties of several mutants are similar for AtSLAC1 and HiTehA.
To examine characteristics of the SLAC1 channel in light of the structural model, we
performed electrophysiological tests of membrane currents from voltage-clamped Xenopus
oocytes following injection of wild-type or mutant AtSLAC1 or HiTehA cRNAs. We
observed modest-sized chloride currents with wild-type AtSLAC1 cRNA, as found
previously6,7, but did not detect any chloride current following injection of wild-type
HiTehA cRNA. We found that SLAC1 Cl− conductance was enhanced when the OST1
kinase cRNA was co-injected with SLAC1 cRNA, but only to the levels found by Lee et al.6
and not to the much higher levels found by Geiger et al.7 with OST1 physically connected to
SLAC1 by split YFP linkage. In keeping with structural evidence that Phe262 blocks the
HiTehA pore, removal of the phenyl group in HiTehA F262A or in the homologous
AtSLAC1 F450A resulted in very large Cl− currents relative to wild-type levels, and the
SLAC1 currents were now less enhanced by presence of OST1 (Fig. 4a). The tempting
interpretation of an opened gate will require validation with appropriately analyzed single-
channel recordings27. In keeping with the slac1-2 phenotype3, neither the functionally
impaired AtSLAC1 G194D nor its HiTehA G15D homolog showed any substantial
conductance; moreover, further consistent with pore blockage by Asp194 in slac1-2, the
large conductances of HiTehA F262A and AtSLAC1 F450A were stopped in the double
mutants AtSLAC1 G194D/F450A and HiTehA G15D/F262A (Fig. 4a). Here again, the
effects in SLAC1 were independent of OST1.
We also tested conductance characteristics for a series of AtSLAC1 F450X substitution
mutants – F450A,G,T,V,L– and for the corresponding HiTehA F262X series –
F262A,G,T,V,L (Figs. 4b,S5, Table S5). Findings from the two series are roughly parallel;
in particular, the alanine and glycine substitutions lead to large currents for both and in
comparison to the others. There are distinctions, of course, including generally higher
conductances for AtSLAC1 over HiTehA and less conductance of F262T TehA compared to
F450T SLAC1. It is also noteworthy that OST1 activation is very muted for the F450A,G,L
mutants, which is consistent with SLAC1 gating at Phe450.
Crystal structures were also determined for several of the HiTehA mutant variants (Table
S4). The structures of F262A (1.15Å), F262V (1.60Å), F262L (1.65Å) and G15D (1.50Å)
are all essentially isomorphous with the wild-type TehA structure with changes localized
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primarily at the sites of mutation; the same is true for the double mutations of F262A/G15D,
F262G/G15D and A208P/G263P26. The F262A structure has a wide open pore (Fig. 5a)
with a relatively uniform pore diameter of ~5Å through ~30Å across the membrane (Fig.
S3) whereas G15D has a doubly occluded pore (Fig. 5b). The pores of other mutant variants
are consistent with the sizes of constrictive residues and with the observed conductances.
Gating and activation
The crystal structures of TehA and its mutant variants when taken together with the
functional studies in Xenopus oocytes point to a crucial role for Phe450 in gating of the
SLAC1 anion channel. Conservation of this residue across the SLAC1 family implies
functional importance. The occlusion of the pore by the presence of F262 in the structure of
wild-type TehA and the openness of the pore upon its substitution by alanine in the structure
of the F262A mutant provides physical evidence for a gating role of this residue. This
interpretation is supported by the correlated conductance characteristics from variants of the
AtSLAC1 and HiTehA channels (Figs. 4b,S5). While these observations may suffice for
placing the gate within the channel pore, they do not by themselves suggest a mechanism for
gating in response to physiological stimuli. Some insight does come from conformational
details defined at high resolution.
One important structural clue is that the side chain of Phe262 is in a high energy
conformation in the HiTehA structure, with χ1/χ2 at −160°/-4°. Although χ1 is in a preferred
trans conformation, the phenyl ring is restricted by contacts with Val210 and Leu18 to a χ2
value near 0° rather than near to the preferred 90° orientation (Fig. 5c). Further evidence that
Phe262 is restrained from local equilibrium comes from shifts observed in crystal structures
of the F262A, F262V, F262L and F262G/G15D variants, which all show consistent
backbone movements that displace Cβ(262) by 0.47-0.78Å (Fig. 5d). By contrast, Leu262 in
F262L is in a preferred trans/gauche+ conformation at χ1/χ2 = 177°/63° as is Val262 in
F262V at χ1 = −176°. What might control activation of HiTehA is unclear, but for AtSLAC1
activation is by OST1 phosphorylation6,7. The molecular consequences of OST1
phosphorylation of SLAC1 remain unknown, but it is plausible that associated shifts in pore-
helix orientations would unlatch Phe450 in SLAC1 from a TehA-like restrained orientation.
By analogy with Leu262 in F262L, we expect a preferred rotameric state for Leu450 in
AtSLAC1 F450L. Thus, the lack of appreciable OST1 activation of conductance in
AtSLAC1 F450L (Fig. 5e) might be explained by the lack of a restraining latch, whereby the
channel remains closed despite OST1 activation. Puzzles certainly remain since OST1 does
substantially activate AtSLAC1 F450T or F450V, which like HiTehA F262V should also be
unrestrained; presumably activating adjustments widen the pore enough for ion permeation
past threonine and valine but not leucine.
Phosphorylation sites have been discovered in the N- and C-terminal tails of AtSLAC16,7,28
(179 and 51 residues long, respectively), but these alone cannot explain OST1 activation of
SLAC1. First SLAH1, which fully complements the slac1-1 mutation, does not have these
cytoplasmic tails. Secondly, although OST1 phosphorylation of Ser120 in the N-terminal tail
is necessary for SLAC1 activation, it is not sufficient7. Thus, we surmise that direct
phosphorylation of the SLAC1 TM domain must be critical, and SLAC1 has four conserved
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Ser/Thr candidates in its cytoplasmic loops. Moreover, SLAC1 proteins have proline-
mediated kinks at the putative Phe450 gate in helix TM9, and also in adjacent helix TM7;
these features may play a role in phosphorylation-driven unlatching of the Phe450 gate in
SLAC1.
Ion selectivity and discrimination
Our studies of SLAC1 channel relative ion permeabilities, based on measurements of current
reversal potential, are consistent with earlier work demonstrating that AtSLAC1 conducts
anions but not cations and is selective among anions, with greater permeability for nitrate
than for chloride (as in Vicia faba guard cell protoplasts27) and much reduced permeability
for malate, bicarbonate or sulfate6,7. We also find that SLAC1 has little permeability for
sulfite. Additionally, we find that wild-type SLAC1, F450A and F450T all have similar
relative permeabilities to chloride, sulfite and malate, despite having widely different
conductance levels, but the gating mutants do show small but significant decreases in nitrate
permeability (Fig. 4c, Table S6).
The relative insensitivity of anion permeability to gating residue changes suggests that
selectivity for these anions may occur away from the central constriction at the channel gate.
To some extent, ionic discrimination must depend on pore geometry; thus, an organic anion
such as malate may be simply too large to pass through the 5-Å wide pore. Although the
SLAC1 pore is lined largely with hydrophobic sidechains (Fig. 3c), it is sprinkled with
hydroxyl groups from serine and threonine residues (16%); their electropositive hydrogen
atoms may facilitate conductance. Most strikingly, the electrostatic potential within the
AtSLAC1 pore is electropositive throughout (Fig. 3a). This polarization, promoted by
charges on extra-membranous loops, no doubt contributes significantly in discrimination
against cations.
The relative anion permeability sequence of SLAC1 determined by us and others, I− > NO3−
> Br− > Cl− 6,7, corresponds to selectivity sequence 1 compiled by Wright and Diamond29
for a range of anion-selective proteins. This sequence correlates inversely with the hydration
energies of monovalent anions – anions with a lower hydration energy have a greater
channel permeability. It is thought to be generated in proteins with weak, low field-strength,
anion binding sites, where selectivity is largely determined by the energetic cost of anion
dehydration. These selectivity results are thus consistent with the SLAC1 structure, where
the pore lacks any obvious anion binding site.
Distinctiveness of the SLAC1 channel
SLAC1 anion channels are entirely novel in structure and, apparently, in the mechanism for
ion conductance. The best characterized of anion channels belong to the CLC family of Cl−
channels and transporters30-32. CLC channels have an altogether different architecture from
the SLAC1 channel, and the mechanism for selectivity is also very different. Bacterial CLC
transporters bind halide ions at three sites in a highly constricted pore30. By contrast, the
SLAC1 pore has a relatively uniform diameter across the membrane, except where closed by
the gating phenyl group, and we do not find discrete ion binding sites. CLC selectivity is
governed by specific residues surrounding these binding sites30,32. The anion selectivity
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sequence for the CLC channels of Cl−>Br−>NO −3>I−, opposite of that in SLAC1, is
consistent with the high field-strength anion binding sites in CLC channels29. Interestingly,
as for AtSLAC1, an Arabidopsis CLCa channel also preferentially transports nitrate ions33,
and an E. coli CLC channel is converted to preference of nitrate when a generally conserved
serine at the central site is substituted with proline as in AtCLCa32.
SLAC1 also differs radically from other structurally characterized anion channels and
transporters. These include the VDAC1 voltage-gated anion channel from mitochondrial
outer membranes, which has a porin-like beta-barrel structure34-36, and a light-driven
halorhodopsin chloride pump, which has a transmembrane conductance pathway similar to
that of the proton-pumping pore of bacteriorhodopsin37. Although its channel structure is
still only known by homology to other ABC transporters, CFTR is another obviously
distinct chloride channel38. Cys-loop receptors also include anion channels39, and these are
similar to SLAC1 in having five-helix pores25, but here selectivity is governed by charged
groups at the entrance to the pore, which distinguish the anion-selective GABAA and
glycine receptors from the cation-selective acetylcholine and serotonin 5HT3 receptors39.
Finally, the recently identified TMEM16A gene for calcium-activated chloride channels40-42
appears to encode an 8-TM protein that is again distinct from SLAC1.
Stomatal guard cells show both rapidly activated (R-type) and slow (S-type) anion channel
activity43. Although slac1 guard cells have very defective S-type activity, their R-type
currents are normal4. Guard cell protoplasts from the slac1-2 mutant abnormally accumulate
Cl−, K+, malate and fumarate3, whereas SLAC1 shows negligible malate conductance7. As
for SLAC1-associated K+ movements, other channels or transporters must be responsible for
SLAC1-associated malate movements. Recent studies indicate that AtALMT12, an
aluminium-activated malate transporter (ALMT) family member, is a malate-dependent R-
type anion channel44 needed for stomatal closure45.
Conclusions
We find that many functional properties of the plant SLAC1 anion channel are explained
well by the structure of an uncharacterized bacterial TehA protein that has been associated
with tellurite resistance. SLAC1 and TehA belong to distinct subfamilies within one branch
of a larger SLAC1 superfamily, but AtSLAC1 and HiTehA are sufficiently similar (19%
sequence identity) that the SLAC1 homology model is predictive for function, including a
verified placement of the identifying slac1-2 mutation G194D and a phenylalanine gate. One
remaining puzzle concerns the structural change that activating phosphorylation elicits in
SLAC1, and another puzzle is the biochemical role of the TehA homologs in bacteria. In a
companion paper26, we examine functional and structural properties of TehA in bacteria,
showing that it is anion channel, although actually not conferring tellurite resistance, and
identifying a mutant variant with properties suggestive of an activated state. Thus, SLAC1
and TehA likely represent a large family of selective anion channels controlled by
environmental stimuli.
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METHODS
Selection of target sequences
TehA from E. coli (EcTehA) was a centrally selected sequence for expansion as a
NYCOMPS targeted family. Target selection criteria utilized at NYCOMPS are described in
details elsewhere46. Briefly, the EcTehA sequence was run against a dataset of about 40,000
predicted alpha helical integral membrane protein sequences from prokaryotic genomes
(NYCOMPS98 dataset46) using PSI-BLAST50. Sequences that matched EcTehA with an E-
value lower than 10−3 in an alignment extending over at least 50% of both predicted TM
regions and passing our post-seed-expansion filtering criteria46 were passed to the protein
production pipeline.
Protein expression screening
Full-length homologs from the following 38 species, including 2 sequences each from 5 of
these, were amplified from genomic DNA by PCR: Thermoplasma acidophilum,
Lactococcus lactis subsp. lactis Il1403, Streptococcus pyogenes M1 GAS, Streptococcus
pneumoniae TIGR4, Haemophilus influenzae Rd KW20, Methanocaldococcus jannaschii
DSM 2661 (2), Escherichia coli K12, Salmonella typhimurium LT2, Clostridium
perfringens ATCC 13124, Xanthomonas campestris pv. campestris str. ATCC 33913,
Streptococcus agalactiae 2603V/R, Shewanella oneidensis MR-1, Streptococcus mutans
UA159, Archaeoglobus fulgidus DSM 4304, Vibrio parahaemolyticus RIMD 2210633 (2),
Pseudomonas syringae pv. tomato str. DC3000 (2), Enterococcus faecalis V583,
Pyrococcus horikoshii OT3, Bordetella bronchiseptica RB50, Streptomyces coelicolor A3,
Corynebacterium glutamicum ATCC 13032, Picrophilus torridus DSM 9790, Acinetobacter
sp. ADP1, Vibrio fischeri ES114, Pseudomonas fluorescens Pf-5, Sulfolobus acidocaldarius
DSM 639, Colwellia psychrerythraea 34H, Neisseria meningitidis MC58, Rhodobacter
sphaeroides 2.4.1, Vibrio cholerae O1 biovar eltor str. N16961 (2), Marinobacter aquaeolei
VT8, Acinetobacter baumannii ATCC 17978, Klebsiella pneumoniae subsp. pneumoniae
MGH 78578, Streptomyces avermitili MA-4680, Bordetella parapertussis 12822 (2),
Streptococcus thermophilus LMG 18311, Salmonella enterica subsp. enterica serovar
Paratyphi A str. ATCC 9150, and Anaeromyxobacter dehalogenans 2CP-C.
Selected cDNAs were cloned into a modified pET vector (Novagen, Inc) that fuses a FLAG
and deca-histidine tag at the C-terminus, which are cleavable by TEV protease. Proteins
were expressed in E. coli BL21(DE3) plysS by a high throughput format (0.6ml in a deep
well block) and purified after lysis by sonication using metal affinity purification in a buffer
containing N-Dodecyl-β-D-Maltopyranoside. Samples were passed over an analytical size
exclusion column in 12 different detergent-containing mobile phases, which included N-
dodecyl-β-D-maltopyranoside (DDM), N-decyl-β-D-altopyranoside (DM), N-nonyl-β-D-
altopyranoside (NM), N-octyl-β-D-altopyranoside (OM), N-octyl-β-D-Glucopyranoside
(OG), N-nonyl-β-D-Glucopyranoside (NG) and lauryl dimethyl amine oxide (LDAO).
Multi-angle light scattering with refractive index detection was used to analyze the
oligomeric state51. The E. coli and H. influenzae proteins were judged to be monodisperse
and stable and were passed to scale up.
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Scaled-up production and purification
For scale up, in brief, transformed BL21 plysS cells were grown at 37°C in 2× YT media to
OD 0.6-0.8 after being inoculated with 1% of the overnight culture. The culture was induced
with 0.4mM IPTG and continued to grow at 37 °C for another 4 hours. The cells were
harvested by centrifugation and stored at −80°C before use. Selenomethionyl (SeMet) TehA
was expressed in a similar way, but using containing SeMet in place of methionine in
defined minimal media. Cells were resuspended in a buffer containing 50 mM Tris-HCl (pH
8.0) and 200 mM NaCl and lysed using a French Press with two passes at 15-20,000 psi.
Cell debris were removed by centrifugation at 10,000g for 20min, and the membrane
fraction was isolated from that supernatant by ultra-centrifugation at 150,000g for 1 hr.
The membrane fraction was homogenized in a solubilization buffer containing 50 mM Tris
(pH 8.0) and 200 mM NaCl, and incubated with final concentration of 1% (w/v) dodecyl-β-
D-maltopyranoside (DDM, Anatrace, Inc) for 1 hr at 4°C. The non-dissolved matter was
remove by ultracentrifugation at 150,000g for 30 min, and the supernatant was loaded to a
5ml Hitrap Ni2+-NTA affinity column (GE Healthcare), pre-equilibrated with the same
solubilization buffer supplemented with 0.05% DDM. After 20 column volume buffer wash,
the protein was eluted with 250mM imidazole in the solublization buffer. The Flag and 10-
His tags were removed by adding super TEV at 1:100 mass ratio and incubating at 4°C
overnight. Tag removal was confirmed by SDS-PAGE, and the resulting sample was
concentrated to around 10mg/ml. Preparative size exclusion chromatography was carried out
on a Superdex-200 column for further purification, removal of TEV protease and the
cleaved tag, and for buffer and detergent exchange. The gel-filtration buffer contained 10
mM Tris (pH 8.0), 200 mM NaCl, 1mM EDTA, 0.5 mM Tris [2-carboxyethyl] phosphine
(TCEP), and 2×CMC of detergent. In the case of HiTehA, the protein was well behaved and
stable in nearly all tested detergents, and we have purified it from DDM, DM, NM, OM, OG
and LDAO.
Protein characterization
We performed N-terminal amino-acid sequencing of purified HiTehA and EcTehA prior to
TEV protease treatment. Results from these analyses proved that true initiating methionine
residue is located 14 residues after one annotated as N-terminal. The intervening nucleotide
sequence contains a Shine-Delgarno sequence.
For cross-linking experiments, purified HiTehA was incubated with 10 mM disuccinimidyl
glutarate at room temperate for 30 min, and 100mM Tris-HCl pH 7.5 was added to stop the
reaction. The incubated sample when run on a 8-25% gradient SDS-PAGE gel showed a
ladder consistent with a trimeric structure.
Crystallization and data collection
Purified protein was concentrated to ~10mg/ml for initial crystal trials in a Mosquito robot
with commercial screens from Hampton research, Emerald Biosystems and Molecular
Dimension. We obtained crystals of HiTehA from protein in detergent DDM, DM, NM,
OM, OG and LDAO, but only those from LDAO and OG gave diffraction to beyond 4Å
spacings. Crystals that proved useful were all grown at 4°C using the sitting-drop vapor
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diffusion method. After extensive optimization we reached conditions supporting very high
resolution. Wild-type HiTehA and most of its variants crystallized from 1mM ZnSO4,
50mM Hepes-Na pH 7.8 and 28% PEG600. HiTehA F262A crystallized from 200mM
Li2SO4, 100mM glycine pH9.3 and 33% PEG400). Addition of 10mM spermidine as an
additive helped to obtain slightly better diffracting crystals. Cryoprotection was achieved by
adding 5% ethylene glycol or PEG400 to the crystallization solution.
Structure determination and refinement
Native and SeMet SAD data set were collected at NSLS beamline X4A and processed using
the software HKL200052. Crystals of HiTehA grew in space group R3 with a=b=c = 85.0Å
and α=β=γ = 93.5°. All subsequent manipulations were done in the hexagonal setting of this
space group with a=b= 96.0Å and c = 136.7Å. The asymmetric unit contains one TehA
protomer and 65% solvent volume. The structure was determined at 2.0Å resolution from
single-wavelength anomalous diffraction (SAD) using selenomethionine-substituted protein
crystals. Assessment of data quality for phasing, location of heavy atom sites and initial
phases were calculated using the HKL2MAP interface to SHELX programs53.
All the secondary structure elements were clearly visible in the experimental electron
density map. Automatic model building was done in Arp/wArp54 and completed manually
in the program COOT55. The model was refined against native data at 1.20Å resolution
using the program Refmac5.5 in CCP456, with anisotropic B factor restrained refinement
applied. Subsequent structural analyses of mutant variants were refined as isomorphous
structures.
Site-directed mutagenesis
Site-directed mutants were constructed using the QuikChange Site-Directed Mutagenesis Kit
(Stratagene, La Jolla, CA 92037) and expressed from pET vectors in E. coli BL21(DE3)
plysS cells as for the wild-type protein.
Electrophysiology
All constructs were cloned into plasmid pGHME2, linearized, and transcribed into cRNA
using T7 polymerase (mMessage mMachine, Ambion). Oocytes were injected with 50 nl of
cRNA solution each, at a constant concentration of 0.5 mg/ml for HiTehA, or 0.5 mg/ml for
AtSLAC1 constructs, with or without 0.5 mg/ml of AtOST1 constructs and recorded in
voltage-clamp experiments 2 days after injection. For mixed expression experiments, 25nl of
cRNA solution was injected for each AtSLAC1 component. Two-electrode voltage clamp
recordings were performed to measure HiTehA or AtSLAC1 currents as described6,7. The
pipette solutions contained 3 M KCl. For voltage-clamp current recordings, the bath solution
contained 1 mM MgCl2, 1 mM CaCl2, 10 mM Mes/Tris (pH 5.6) and 30 mM CsCl; For
anion selectivity measurements, the bath solutions contained 50 mM Cl−, NO −3, malate, or
30 mM SO 2-3 sodium salts, 45 mM Na-gluconate, 1 mM Ca-gluconate2, 1 mM Mg-
gluconate2, 1 mM K-gluconate, as well as 10 mM Tris/Mes (pH 5.6). Osmolality was
adjusted with D-mannitol to 220 mOsmol/kg. The bath electrode was a 3M KCl agar bridge.
The testing potentials were +50 to −110 or −130 mV in 20 mV decrements with 7.5 s
duration. The holding and return potentials were 0 mV with a 1.45 s or 2 s, respectively. I-V
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relations of HiTehA or AtSLAC1 were generated from the current reached after 0.5 s in
testing potentials. The Goldman-Hodgkin-Katz equation was applied to estimate
permeability ratios for monovalent ions as described6. For divalent anions, the permeability
ratios were derived according to Fatt and Ginsborg57.
Bioinformatic analysis of SLAC-related proteins
Sequences related to SLAC1 were analyzed comprehensively by PSI-BLAST50. Searches at
E<10−3 starting from five disparate homologs each identified a common pool of over 900
proteins, which when pooled were used for sub-classification into families and subfamilies.
Details of these analyses are reported in footnotes to Table S1.
Molecular figures were produced in PyMOL58.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We thank Filippo Mancia, Larry Shapiro and Ming Zhou for helpful discussions; Patricia Rodriguez and Arianne
Morrison for help with cloning and expression; Jonah Cheung, Mark Collins, Chris Min, Sheng Ye and Zhen Zhang
for advice in protein chemistry and crystallography; John Schwanof and Randy Abramowitz for help with
synchrotron experiments; John Riley for help with Xenopus oocyte injections; and Min Su for advice on the PSI-
BLAST analysis of SLAC relatives. This project was supported in part by an award to the New York Consortium
on Membrane Protein Structure (NYCOMPS) from the NIGMS Protein Structure Initiative. Beamline X4A at the
National Synchrotron Light Source (NSLS), a DOE facility at Brookhaven National Laboratory, is supported by the
New York Structural Biology Center.
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Figure 1. Sequence analysis for the SLAC1 superfamily
a, Family tree. The presentation was computed by the program COBALT47 from
representative subfamily sequences (Table S1), including Arabidopsis thaliana SLAC1 for
SF1Ai, Haemophilus influenzae TehA for SF1Ci, Escherichia coli TehA for SF1Cii, Vibrio
parahaemolyticus for SF2A, Staphylococcus aureus for SF2B, Aspergillus fumigatus Ssu1
for SF3A, and Schizosaccharomyces pombe Mae1 for SF3B. b, Structure-based sequence
alignment of TehA from Haemophilus influenzae (HiTehA) and SLAC1 from Arabidopsis
thaliana (AtSLAC1). The TehA structure has been used to restrict sequence gaps to inter-
helical segments. Superior coils define extents of the HiTehA helical segments; red letters
mark residue identities; red boxes are drawn for residues that are >95% identical within the
plant subfamily SF1A for AtSLAC1 or within the TehA subfamily for HiTehA; red
diamonds mark HiTehA residues that line the central pore; and the colored inferior bar
encodes ConSurf sequence variability48 for the SF1 family of 204 non-redundant proteins.
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Figure 2. Crystal structure of HiTehA and homology model of AtSLAC1
a, Electron density distribution from the HiTehA crystal structure at 1.2Å resolution. The
map has (2Fo-Fc) coefficients based on the superimposed model. Contours are at 2.5σ. b,
Ribbon diagram of the HiTehA trimer. Each protomer is colored spectrally from blue at its
N-terminus to red at its C-terminus. c, DelPhi49 electrostatic potential at the extracellular
surface. Electronegative and electropositive potential are colored in degrees of red and blue
saturation, respectively. d, Electrostatic potential49 at the intracellular surface. e, Ribbon
diagram of an HiTehA protomer viewed from outside the membrane. The ribbon is colored
spectrally as in 2b. f, Ribbon diagram of an HiTehA protomer viewed from within the
membrane from below the view of e. g, Surface of a homology model of AtSLAC1, viewed
as in f, and colored by sequence variability48. h, Surface of AtSLAC1 as in g, but colored by
electrostatic potential49.
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Figure 3. Putative structure of the SLAC1 conductance pore
a, Cross-section through the homology model of AtSLAC1. The model is viewed as in 2i,
with the electrostatic potential49 shown on the external surface of the molecular envelope.
The side chain of Phe450 is shown as a stick model (red) on the backbone ribbon colored
yellow. b, Cross-section as in a, but colored by surface conservation48 as in 2h. c, Pore-
lining residues in the SLAC1 homology model. A cylinder model (left), spectrally colored as
in 2b, provides a key for viewing the rolled-open structure (left) with pore-lining residues of
AtSLAC1 shown on the TModd helices. d, Ribbon diagrams of HiTehA TM9 (left) and TM7
(right) viewed from within the conductance pore. The side chains of Pro207 and Phe262 are
shown as well as a kink-stabilizing water HOH25 that is coordinated by the NH of Gly263
and by C=O groups of Gly202 and Ala259. Density contours are shown for the water
molecule.
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Figure 4. Ionic conductance measurements
a, Typical microelectrode voltage-clamp current traces from oocytes injected with various
channel cRNAs. Left column, oocytes injected with cRNAs encoding wild-type HiTehA
channels (WT), or F262A, G15D/F262A or G15D mutants. Right two columns, oocytes
injected with cRNAs for WT AtSLAC1, or F450A, G194D/F450A or G194D mutants, with
or without co-injection of AtOST1. Dotted lines represent zero current levels. Extracellular
solution contained 30mM CsCl. Schematic keys at the left place phenyl and/or aspartyl
gates. b, Effects of gating residue mutations. Mean chloride currents, measured at −90 mV,
are shown comparing WT HiTehA and its mutant series F262A,G,T,V,L with WT AtSLAC1
and its corresponding series F450A,G,T,V,L, both alone and co-expressed with AtOST1.
Full I-V relations are shown in Fig. S5. c, Effect of gating residues on relative AtSLAC1
anion permeabilities. Relative permeabilities (P[X]/P[Cl]) for chloride, nitrate, sulfite and
malate of WT, F450A and F450T SLAC1 channels were measured from the change in
current reversal potential with Cl− or anion X− as the sole permeant anion in the bath
solution (Methods, Table S6).
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Figure 5. Structural features at the SLAC1 homolog gate
a,b, Cross-sections through the conductance pores of HiTehA F262A and HiTehA G15D.
The view and presentations are as in 3a, except that helices are colored purple. c, Molecular
basis for conformational strain in gating residue Phe262 of HiTehA. Helices TM1 (left),
TM9 (center) and TM7 (right) are viewed from within the pore and presented as ribbon
diagrams with selected side chains drawn in stick representation. The local low-energy
conformation for the phenyl ring (χ2 = 90°) is shown in thin lines with short contacts
indicated in dashed lines: Leu18 Cδ2 – Phe262 Cδ1, d = 2.4 Å; Val210 Cγ2 – Phe262 Cε2, d
= 2.8 Å. d, Conformational shifts consequent to release of strain in gating residue Phe262.
Cα backbone structures of F262A, F262V and F262L HiTehA were first superimposed onto
WT HiTehA, and residues 258-266 were drawn in stereo as superimposed. All backbone
atoms are shown for peptides 262±1, but only Cα atoms are shown otherwise. The WT
backbone and phenyl group are green; other backbone are all magenta; side chains of
Ala262, Val262 and Leu262 are cyan, blue and red, respectively; oxygen-directed bonds are
red and nitrogen-directed bonds are blue. e, Typical microelectrode voltage-clamp current
traces from oocytes injected with wild-type (WT) or F450L AtSLAC1 cRNA. Experimental
conditions and displays are as in 4a.
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|
3M76
|
Crystal Structure of Plant SLAC1 homolog TehA
|
Homolog Structure of the SLAC1 Anion Channel for Closing
Stomata in Leaves
Yuhang Chen1,6, Lei Hu2, Marco Punta6,7, Renato Bruni6, Brandan Hillerich6, Brian Kloss6,
Burkhard Rost1,6,1, James Love6, Steven A. Siegelbaum2,3,5, and Wayne A.
Hendrickson1,4,5,6
1Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY
10032, USA
2Department of Neuroscience, Columbia University, New York, NY 10032, USA
3Department of Pharmacology, Columbia University, New York, NY 10032, USA
4Department of Physiology and Cellular Biophysics, Columbia University, New York, NY 10032,
USA
5Howard Hughes Medical Institute, Columbia University, New York, NY 10032, USA
6NYCOMPS, New York Structural Biology Center 89 Convent Avenue, New York, NY 10027 USA
7Department of Computer Science and Institute for Advanced Study Technical University of
Munich D-85748 Munich, Germany
Summary
The plant SLAC1 anion channel controls turgor pressure in the aperture-defining guard cells of
plant stomata, thereby regulating exchange of water vapor and photosynthetic gases in response to
environmental signals such as drought or high levels of carbon dioxide. We determined the crystal
structure of a bacterial homolog of SLAC1 at 1.20Å resolution, and we have used structure-
inspired mutagenesis to analyze the conductance properties of SLAC1 channels. SLAC1 is a
symmetric trimer composed from quasi-symmetric subunits, each having ten transmembrane
helices arranged from helical hairpin pairs to form a central five-helix transmembrane pore that is
gated by an extremely conserved phenylalanine residue. Conformational features suggest a
mechanism for control of gating by kinase activation, and electrostatic features of the pore coupled
with electrophysiological characteristics suggest that selectivity among different anions is largely
a function of the energetic cost of ion dehydration.
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Correspondence and request for materials should be addressed to WAH (wayne@convex.hhmi.columbia.edu)..
Supplementary Information is linked to the online version of the paper at www.nature.com/nature.
Author Contributions YC, LH, MP, RB, BH, BK and JL performed experiments; YC, LH, BR, SAS, and WAH analyzed data; YC,
LH, SAS, and WAH prepared the manuscript.
Author Information Atomic coordinates and diffraction data are deposited in the Protein Data Bank with accession codes 3M71,
3M73, 3M74, 3M75, 3M76, and 3M7L. See Table S4 for identifications. Reprints and permissions information is available at
www.Nature.com/reprints.
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Nature. 2010 October 28; 467(7319): 1074–1080. doi:10.1038/nature09487.
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Stomatal pores in the leaves of plants permit the influx of atmospheric carbon dioxide in
exchange for transpirational evaporation of water1,2. A pair of kidney-shaped guard cells
define each pore aperture, and turgor pressure variation in these cells determines the degree
of stomatal pore openness. Depending on diverse environmental factors, the stomata close to
prevent H2O losses and open to admit CO2 for photosynthesis. Environmental stimuli that
lead to stomatal closure include darkness, high CO2 levels, ozone, low air humidity and
drought. The plant hormone abscisic acid (ABA) is critical for signal transduction from
these stimuli. Mutational screens in Arabidopsis thaliana for CO2 and ozone sensitivity
identified a protein with ten predicted transmembrane (TM) helices, now called slow anion
channel 1 (SLAC1), as having a central role in the control of stomatal closure3-5. Recent
studies proved that SLAC1 is indeed an anion channel6-7, with characteristics like those of
slow anion channels found in guard cells8, and that it is activated by phosphorylation from
the OST1 kinase9. OST1 activity is negatively regulated by the ABI1 phosphatase10,11,
which is in turn inhibited by the stomatal ABA receptors PYR/RCAR12,13 when in the
ternary hormone-receptor-phosphatase complex14-18. Thereby, ABA stimulates SLAC1
channel activity. Resulting Cl− efflux through SLAC1 causes membrane depolarization,
which activates outward-rectifying K+ channels, leading to KCl and water efflux to further
reduce turgor and cause stomatal closure.
SLAC1 expression in Arabidopsis is confined to the guard cells of leaves, but other
Arabidopsis tissues do have SLAC1 homologs3, named SLAH1-SLAH4. The identifying
mutations slac1-14 and slac1-23 are, respectively, in predicted transmembrane segments 9
(S456F) and 1 (G194D) of a protein that includes substantial N- and C-terminal extensions
outside a 10-helix transmembrane (TM) domain. SLAH1, which is absent from leaves and
lacks the terminal extensions of SLAC1, fully complements the mutant phenotype in slac1-2
guard cell protoplasts3. SLAC1 and homologs are also present in other plant genomes,
including nine in rice (Oryza sativa) and five in grape vines (Vitis vinifera). SLAC1
relatives, some quite remote, also occur in bacteria, archaea and fungi. Known prokaryotic
homologs contain only the predicted transmembrane domain of SLAC1, but some fungal
homologs do have N- and C-terminal extensions. One homolog, Mae1 from the yeast
Schizosaccharomyces pombe, functions as a malate uptake transporter19; another, Ssu1 from
Saccharomyces cerevisiae and other fungi including Aspergillus fumigatus, is characterized
as a sulfite efflux pump20,21; and TehA from Escherichia coli is identified as a tellurite
resistance protein by virtue of its association in the tehA/tehB operon22,23. Despite a lack of
further biochemical characterization, many homologs are annotated as tellurite resistance/
dicarboxylate transporter (TDT) proteins.
We have undertaken structural and functional characterizations of the SLAC1 anion
channel. We first solved an atomic-resolution crystal structure of the TehA homolog from
Haemophilus influenzae, and we then developed a homology model for Arabidopsis SLAC1.
This model allowed us to conduct mutagenesis for functional testing of structure-inspired
hypotheses on gating and selectivity. We expressed Haemophilus TehA and Arabidopsis
SLAC1 in Xenopus oocytes to characterize channel properties of these proteins and mutant
variants. We also determined crystal structures for several mutant variants, including the
homolog of slac1-2.
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Structure of SLAC1 bacterial homolog TehA
We performed a bioinformatic analysis of SLAC1-related proteins, first clustering nearly
900 non-redundant sequences into a superfamily at the PSI-BLAST level E≤10−3, then into
three distinct families at an initial threshold of E≤10−30, and finally into subfamilies at a
typical initial threshold of E≤10−55. Since previous annotation is not well founded in
experiment and SLAC1 is now the best characterized member, we adopt a nomenclature
defining a SLAC superfamily divided into families identified as SF1-SF3 and subfamilies
SF1A, SF1B etc. Family SF1 comprises the plant SLAC proteins and close bacterial
homologs, family SF2 comprises a distinct set of bacterial proteins often annotated as
exfoliative toxins, and family SF3 comprises the fungal Mae1 and Ssu1 proteins and their
archaeal or bacterial homologs, respectively. SLAC family SF1 has three large subfamilies:
the plant SLAC and SLAH proteins are in subfamily SF1A, closest bacterial homologs are
in SF1B, and the TehA homologs are in SF1C (Fig. 1a). The other families also divide into
subfamilies as detailed in Table S1, and family SF1 is divided into sub-subfamilies (Table
S2). Two pertinent SF1 sequences are aligned in Fig. 1b.
We used a structural genomics approach to obtain structural information, testing expression
and purification for 43 bacterial and archaeal likely homologs, assaying for detergent choice
and stability on 8 of these, finding two with appropriate profiles by size exclusion
chromatography, and obtaining suitable crystals for one. This protein, TehA from H.
influenzae (HiTehA), was found to be trimeric both by size-exclusion multi-angle light
scattering (SEC-MALS) measurements and by chemical cross-linking. When solubilized in
β-octylglucoside, HiTehA crystallized in space group R3 with a=b=96.01Å and c=136.27Å.
Each asymmetric unit contains one subunit and 65% solvent. The structure was solved by
selenomethionyl (SeMet) SAD phasing, ultimately at 1.50Å resolution (Table S3, Fig. S1),
and then refined at 1.20Å resolution (Fig. 2a) to R/Rfree values of 14.1%/15.7% for a model
that includes ordered residues 6-313, 213 water molecules and four detergent molecules
(Table S4).
The crystal structure has TehA trimers aligned with three-fold axes of the lattice (Fig. 2b).
Subunits are tightly associated, burying 8947 Å2 of total surface area within trimer
interfaces. The electrostatic potential surface is largely negative on the extracellular surface
(Fig. 2c) and largely positive on the cytoplasmic surface (Fig. 2d). The membrane
orientation is specified experimentally from GFP tagging of E. coli TehA24. Each TehA
protomer has ten TM helices, as predicted; however, the fold is novel. Tandemly repeated
helical hairpins are arranged with quasi-five-fold symmetry (Figs. 2e,S2). Extracellular
inter-helix loops are short (2-5 residues) whereas intracellular inter-helix connections are
longer, including a nine-residue helix H2,3 between TM2 and TM3 (Fig. 2f). An inner pentad
of outwardly directed, TModd, helices creates an apparent pore through each protomer
perpendicular to the putative membrane plane. TMeven helices from the five hairpins
surround the inner pore and make an outer layer.
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Homology model for plant SLAC1
Arabidopsis SLAC1 (AtSLAC1) is substantially similar to bacterial homologs, notably
HiTehA (Fig. 1b). All HiTehA TM helices are fully aligned to predicted SLAC1 TM
helices, but there are short inter-helical gaps (1-5 residues) in all five extracellular loops and
in two of the intracellular loops. The TM domain of AtSLAC1 (residues 188-504) aligns to
HiTehA with 19% sequence identity and with a PSI-BLAST E-value of 3×10−22. For
comparison, and in keeping with the family tree (Fig. 1a), the TM domain of AtSLAC1
shares sequence identities of 76% with rice SLAC1, 41% with Arabidopsis SLAH1, 25%
with an SL1B homolog from Halorhodospira halophila, 11% with S. cerevisiae Ssu1, and
9% with S. pombe Mae1. A conceptual model with the AtSLAC1 sequence transposed onto
the HiTehA helices sufficed to guide most of our mutational tests, but a detailed AtSLAC1
homology model helped to refine our ideas. Surface variability and electrostatic potential are
plotted onto the surface of this model (Fig. 2g,2h).
The most remarkable feature of the TehA structure and corresponding SLAC1 model is the
central pore through each protomer. As for acetylcholine receptors25, the SLAC1 pore is
formed by five helices; but the SLAC1 helices come from one protein molecule rather than
five. The SLAC1 pore has a relatively uniform diameter of approximately 5Å across nearly
five helical turns (Fig. S3), except for a pronounced constriction in the middle of the
membrane (Fig. 3a) where the pore is occluded by the side chain of Phe 450 (Phe 262 in
HiTehA). This residue is the only absolutely conserved amino acid residue of the SLAC1
family. The pore is lined with highly conserved (86% identity among five SLAC1 orthologs;
32% identity between AtSLAC1 and HiTehA) and generally hydrophobic residues (Figs. 1b,
3b,3c,S4). Despite this hydrophobicity, the electrostatic potential on the pore surface is
polarized (Fig. 3a), presumably due to an invaginated shape adjacent to charged residues
outside the membrane. The generally electropositive character of the cytoplasmic surface
likely contributes to anion efflux.
Kinks in the pore helices contribute to formation of a relatively constant pore diameter
across the membrane. Four of the five HiTehA inner helices have centrally located proline
residues, which necessarily generate kinks, and TM9 is kinked at a backbone-coordinated
water molecule (Fig. 3d). Proline replaces Gly263 in all SF1A and SF1B relatives, including
Pro451 AtSLAC1. This water-displacing change is isostructural26. Centrally located prolines
also prevail in TM3, TM5 and TM7 across the SF1 proteins. The outer helices are longer and
straighter, but more inclined. The only outer-helix proline kink of HiTehA is in TM6 at the
trimer three-fold axis.
Two of the gene-identifying mutations in Arabadopsis SLAC1 are selected point mutations,
others are disruptive transfer DNA (T-DNA) insertions4. In the AtSLAC1 homology model,
the slac1-2 (G194D) mutation3 points into the pore from TM1 and can be accommodated
structurally whereas the slac1-1 (S456F) mutation4 points away from the pore six residues
after pore-blocking Phe450 on TM9 and would be expected to disruptive. Ser456 interacts
with outer helix TM10 in the homology model, and the phenyl bulk from S456F would not
fit. Position 456 has alanine in HiTehA, 66% of all 204 SF1 homologs and another 27%
have threonine or serine (as in all SLAC1 channels); phenylalanine never occurs among all
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814 SLAC superfamily members. Position 194 has glycine in 58% of SF1 homologs and
alanine in another 22%. Asp194, which never occurs naturally, would block the pore and be
expected to repel anions.
Mutational tests of channel function
Mutational studies corroborate the hypothesis that a TehA-based SLAC1 model is
appropriate. First, as discussed above, the slac1-1 mutant (S456F) is expected to be
structurally disruptive, and indeed it is inactive in guard cells4, and the slac1-2 mutation
(G194D) is expected to block the pore, and we show below that this variant is also inactive.
We have also shown that the introduction of SLAC1-conserved proline residues into
HiTehA (A208P/G263P) is accommodated isomorphously26. Moreover, as shown below,
channel conductance properties of several mutants are similar for AtSLAC1 and HiTehA.
To examine characteristics of the SLAC1 channel in light of the structural model, we
performed electrophysiological tests of membrane currents from voltage-clamped Xenopus
oocytes following injection of wild-type or mutant AtSLAC1 or HiTehA cRNAs. We
observed modest-sized chloride currents with wild-type AtSLAC1 cRNA, as found
previously6,7, but did not detect any chloride current following injection of wild-type
HiTehA cRNA. We found that SLAC1 Cl− conductance was enhanced when the OST1
kinase cRNA was co-injected with SLAC1 cRNA, but only to the levels found by Lee et al.6
and not to the much higher levels found by Geiger et al.7 with OST1 physically connected to
SLAC1 by split YFP linkage. In keeping with structural evidence that Phe262 blocks the
HiTehA pore, removal of the phenyl group in HiTehA F262A or in the homologous
AtSLAC1 F450A resulted in very large Cl− currents relative to wild-type levels, and the
SLAC1 currents were now less enhanced by presence of OST1 (Fig. 4a). The tempting
interpretation of an opened gate will require validation with appropriately analyzed single-
channel recordings27. In keeping with the slac1-2 phenotype3, neither the functionally
impaired AtSLAC1 G194D nor its HiTehA G15D homolog showed any substantial
conductance; moreover, further consistent with pore blockage by Asp194 in slac1-2, the
large conductances of HiTehA F262A and AtSLAC1 F450A were stopped in the double
mutants AtSLAC1 G194D/F450A and HiTehA G15D/F262A (Fig. 4a). Here again, the
effects in SLAC1 were independent of OST1.
We also tested conductance characteristics for a series of AtSLAC1 F450X substitution
mutants – F450A,G,T,V,L– and for the corresponding HiTehA F262X series –
F262A,G,T,V,L (Figs. 4b,S5, Table S5). Findings from the two series are roughly parallel;
in particular, the alanine and glycine substitutions lead to large currents for both and in
comparison to the others. There are distinctions, of course, including generally higher
conductances for AtSLAC1 over HiTehA and less conductance of F262T TehA compared to
F450T SLAC1. It is also noteworthy that OST1 activation is very muted for the F450A,G,L
mutants, which is consistent with SLAC1 gating at Phe450.
Crystal structures were also determined for several of the HiTehA mutant variants (Table
S4). The structures of F262A (1.15Å), F262V (1.60Å), F262L (1.65Å) and G15D (1.50Å)
are all essentially isomorphous with the wild-type TehA structure with changes localized
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primarily at the sites of mutation; the same is true for the double mutations of F262A/G15D,
F262G/G15D and A208P/G263P26. The F262A structure has a wide open pore (Fig. 5a)
with a relatively uniform pore diameter of ~5Å through ~30Å across the membrane (Fig.
S3) whereas G15D has a doubly occluded pore (Fig. 5b). The pores of other mutant variants
are consistent with the sizes of constrictive residues and with the observed conductances.
Gating and activation
The crystal structures of TehA and its mutant variants when taken together with the
functional studies in Xenopus oocytes point to a crucial role for Phe450 in gating of the
SLAC1 anion channel. Conservation of this residue across the SLAC1 family implies
functional importance. The occlusion of the pore by the presence of F262 in the structure of
wild-type TehA and the openness of the pore upon its substitution by alanine in the structure
of the F262A mutant provides physical evidence for a gating role of this residue. This
interpretation is supported by the correlated conductance characteristics from variants of the
AtSLAC1 and HiTehA channels (Figs. 4b,S5). While these observations may suffice for
placing the gate within the channel pore, they do not by themselves suggest a mechanism for
gating in response to physiological stimuli. Some insight does come from conformational
details defined at high resolution.
One important structural clue is that the side chain of Phe262 is in a high energy
conformation in the HiTehA structure, with χ1/χ2 at −160°/-4°. Although χ1 is in a preferred
trans conformation, the phenyl ring is restricted by contacts with Val210 and Leu18 to a χ2
value near 0° rather than near to the preferred 90° orientation (Fig. 5c). Further evidence that
Phe262 is restrained from local equilibrium comes from shifts observed in crystal structures
of the F262A, F262V, F262L and F262G/G15D variants, which all show consistent
backbone movements that displace Cβ(262) by 0.47-0.78Å (Fig. 5d). By contrast, Leu262 in
F262L is in a preferred trans/gauche+ conformation at χ1/χ2 = 177°/63° as is Val262 in
F262V at χ1 = −176°. What might control activation of HiTehA is unclear, but for AtSLAC1
activation is by OST1 phosphorylation6,7. The molecular consequences of OST1
phosphorylation of SLAC1 remain unknown, but it is plausible that associated shifts in pore-
helix orientations would unlatch Phe450 in SLAC1 from a TehA-like restrained orientation.
By analogy with Leu262 in F262L, we expect a preferred rotameric state for Leu450 in
AtSLAC1 F450L. Thus, the lack of appreciable OST1 activation of conductance in
AtSLAC1 F450L (Fig. 5e) might be explained by the lack of a restraining latch, whereby the
channel remains closed despite OST1 activation. Puzzles certainly remain since OST1 does
substantially activate AtSLAC1 F450T or F450V, which like HiTehA F262V should also be
unrestrained; presumably activating adjustments widen the pore enough for ion permeation
past threonine and valine but not leucine.
Phosphorylation sites have been discovered in the N- and C-terminal tails of AtSLAC16,7,28
(179 and 51 residues long, respectively), but these alone cannot explain OST1 activation of
SLAC1. First SLAH1, which fully complements the slac1-1 mutation, does not have these
cytoplasmic tails. Secondly, although OST1 phosphorylation of Ser120 in the N-terminal tail
is necessary for SLAC1 activation, it is not sufficient7. Thus, we surmise that direct
phosphorylation of the SLAC1 TM domain must be critical, and SLAC1 has four conserved
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Ser/Thr candidates in its cytoplasmic loops. Moreover, SLAC1 proteins have proline-
mediated kinks at the putative Phe450 gate in helix TM9, and also in adjacent helix TM7;
these features may play a role in phosphorylation-driven unlatching of the Phe450 gate in
SLAC1.
Ion selectivity and discrimination
Our studies of SLAC1 channel relative ion permeabilities, based on measurements of current
reversal potential, are consistent with earlier work demonstrating that AtSLAC1 conducts
anions but not cations and is selective among anions, with greater permeability for nitrate
than for chloride (as in Vicia faba guard cell protoplasts27) and much reduced permeability
for malate, bicarbonate or sulfate6,7. We also find that SLAC1 has little permeability for
sulfite. Additionally, we find that wild-type SLAC1, F450A and F450T all have similar
relative permeabilities to chloride, sulfite and malate, despite having widely different
conductance levels, but the gating mutants do show small but significant decreases in nitrate
permeability (Fig. 4c, Table S6).
The relative insensitivity of anion permeability to gating residue changes suggests that
selectivity for these anions may occur away from the central constriction at the channel gate.
To some extent, ionic discrimination must depend on pore geometry; thus, an organic anion
such as malate may be simply too large to pass through the 5-Å wide pore. Although the
SLAC1 pore is lined largely with hydrophobic sidechains (Fig. 3c), it is sprinkled with
hydroxyl groups from serine and threonine residues (16%); their electropositive hydrogen
atoms may facilitate conductance. Most strikingly, the electrostatic potential within the
AtSLAC1 pore is electropositive throughout (Fig. 3a). This polarization, promoted by
charges on extra-membranous loops, no doubt contributes significantly in discrimination
against cations.
The relative anion permeability sequence of SLAC1 determined by us and others, I− > NO3−
> Br− > Cl− 6,7, corresponds to selectivity sequence 1 compiled by Wright and Diamond29
for a range of anion-selective proteins. This sequence correlates inversely with the hydration
energies of monovalent anions – anions with a lower hydration energy have a greater
channel permeability. It is thought to be generated in proteins with weak, low field-strength,
anion binding sites, where selectivity is largely determined by the energetic cost of anion
dehydration. These selectivity results are thus consistent with the SLAC1 structure, where
the pore lacks any obvious anion binding site.
Distinctiveness of the SLAC1 channel
SLAC1 anion channels are entirely novel in structure and, apparently, in the mechanism for
ion conductance. The best characterized of anion channels belong to the CLC family of Cl−
channels and transporters30-32. CLC channels have an altogether different architecture from
the SLAC1 channel, and the mechanism for selectivity is also very different. Bacterial CLC
transporters bind halide ions at three sites in a highly constricted pore30. By contrast, the
SLAC1 pore has a relatively uniform diameter across the membrane, except where closed by
the gating phenyl group, and we do not find discrete ion binding sites. CLC selectivity is
governed by specific residues surrounding these binding sites30,32. The anion selectivity
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sequence for the CLC channels of Cl−>Br−>NO −3>I−, opposite of that in SLAC1, is
consistent with the high field-strength anion binding sites in CLC channels29. Interestingly,
as for AtSLAC1, an Arabidopsis CLCa channel also preferentially transports nitrate ions33,
and an E. coli CLC channel is converted to preference of nitrate when a generally conserved
serine at the central site is substituted with proline as in AtCLCa32.
SLAC1 also differs radically from other structurally characterized anion channels and
transporters. These include the VDAC1 voltage-gated anion channel from mitochondrial
outer membranes, which has a porin-like beta-barrel structure34-36, and a light-driven
halorhodopsin chloride pump, which has a transmembrane conductance pathway similar to
that of the proton-pumping pore of bacteriorhodopsin37. Although its channel structure is
still only known by homology to other ABC transporters, CFTR is another obviously
distinct chloride channel38. Cys-loop receptors also include anion channels39, and these are
similar to SLAC1 in having five-helix pores25, but here selectivity is governed by charged
groups at the entrance to the pore, which distinguish the anion-selective GABAA and
glycine receptors from the cation-selective acetylcholine and serotonin 5HT3 receptors39.
Finally, the recently identified TMEM16A gene for calcium-activated chloride channels40-42
appears to encode an 8-TM protein that is again distinct from SLAC1.
Stomatal guard cells show both rapidly activated (R-type) and slow (S-type) anion channel
activity43. Although slac1 guard cells have very defective S-type activity, their R-type
currents are normal4. Guard cell protoplasts from the slac1-2 mutant abnormally accumulate
Cl−, K+, malate and fumarate3, whereas SLAC1 shows negligible malate conductance7. As
for SLAC1-associated K+ movements, other channels or transporters must be responsible for
SLAC1-associated malate movements. Recent studies indicate that AtALMT12, an
aluminium-activated malate transporter (ALMT) family member, is a malate-dependent R-
type anion channel44 needed for stomatal closure45.
Conclusions
We find that many functional properties of the plant SLAC1 anion channel are explained
well by the structure of an uncharacterized bacterial TehA protein that has been associated
with tellurite resistance. SLAC1 and TehA belong to distinct subfamilies within one branch
of a larger SLAC1 superfamily, but AtSLAC1 and HiTehA are sufficiently similar (19%
sequence identity) that the SLAC1 homology model is predictive for function, including a
verified placement of the identifying slac1-2 mutation G194D and a phenylalanine gate. One
remaining puzzle concerns the structural change that activating phosphorylation elicits in
SLAC1, and another puzzle is the biochemical role of the TehA homologs in bacteria. In a
companion paper26, we examine functional and structural properties of TehA in bacteria,
showing that it is anion channel, although actually not conferring tellurite resistance, and
identifying a mutant variant with properties suggestive of an activated state. Thus, SLAC1
and TehA likely represent a large family of selective anion channels controlled by
environmental stimuli.
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METHODS
Selection of target sequences
TehA from E. coli (EcTehA) was a centrally selected sequence for expansion as a
NYCOMPS targeted family. Target selection criteria utilized at NYCOMPS are described in
details elsewhere46. Briefly, the EcTehA sequence was run against a dataset of about 40,000
predicted alpha helical integral membrane protein sequences from prokaryotic genomes
(NYCOMPS98 dataset46) using PSI-BLAST50. Sequences that matched EcTehA with an E-
value lower than 10−3 in an alignment extending over at least 50% of both predicted TM
regions and passing our post-seed-expansion filtering criteria46 were passed to the protein
production pipeline.
Protein expression screening
Full-length homologs from the following 38 species, including 2 sequences each from 5 of
these, were amplified from genomic DNA by PCR: Thermoplasma acidophilum,
Lactococcus lactis subsp. lactis Il1403, Streptococcus pyogenes M1 GAS, Streptococcus
pneumoniae TIGR4, Haemophilus influenzae Rd KW20, Methanocaldococcus jannaschii
DSM 2661 (2), Escherichia coli K12, Salmonella typhimurium LT2, Clostridium
perfringens ATCC 13124, Xanthomonas campestris pv. campestris str. ATCC 33913,
Streptococcus agalactiae 2603V/R, Shewanella oneidensis MR-1, Streptococcus mutans
UA159, Archaeoglobus fulgidus DSM 4304, Vibrio parahaemolyticus RIMD 2210633 (2),
Pseudomonas syringae pv. tomato str. DC3000 (2), Enterococcus faecalis V583,
Pyrococcus horikoshii OT3, Bordetella bronchiseptica RB50, Streptomyces coelicolor A3,
Corynebacterium glutamicum ATCC 13032, Picrophilus torridus DSM 9790, Acinetobacter
sp. ADP1, Vibrio fischeri ES114, Pseudomonas fluorescens Pf-5, Sulfolobus acidocaldarius
DSM 639, Colwellia psychrerythraea 34H, Neisseria meningitidis MC58, Rhodobacter
sphaeroides 2.4.1, Vibrio cholerae O1 biovar eltor str. N16961 (2), Marinobacter aquaeolei
VT8, Acinetobacter baumannii ATCC 17978, Klebsiella pneumoniae subsp. pneumoniae
MGH 78578, Streptomyces avermitili MA-4680, Bordetella parapertussis 12822 (2),
Streptococcus thermophilus LMG 18311, Salmonella enterica subsp. enterica serovar
Paratyphi A str. ATCC 9150, and Anaeromyxobacter dehalogenans 2CP-C.
Selected cDNAs were cloned into a modified pET vector (Novagen, Inc) that fuses a FLAG
and deca-histidine tag at the C-terminus, which are cleavable by TEV protease. Proteins
were expressed in E. coli BL21(DE3) plysS by a high throughput format (0.6ml in a deep
well block) and purified after lysis by sonication using metal affinity purification in a buffer
containing N-Dodecyl-β-D-Maltopyranoside. Samples were passed over an analytical size
exclusion column in 12 different detergent-containing mobile phases, which included N-
dodecyl-β-D-maltopyranoside (DDM), N-decyl-β-D-altopyranoside (DM), N-nonyl-β-D-
altopyranoside (NM), N-octyl-β-D-altopyranoside (OM), N-octyl-β-D-Glucopyranoside
(OG), N-nonyl-β-D-Glucopyranoside (NG) and lauryl dimethyl amine oxide (LDAO).
Multi-angle light scattering with refractive index detection was used to analyze the
oligomeric state51. The E. coli and H. influenzae proteins were judged to be monodisperse
and stable and were passed to scale up.
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Scaled-up production and purification
For scale up, in brief, transformed BL21 plysS cells were grown at 37°C in 2× YT media to
OD 0.6-0.8 after being inoculated with 1% of the overnight culture. The culture was induced
with 0.4mM IPTG and continued to grow at 37 °C for another 4 hours. The cells were
harvested by centrifugation and stored at −80°C before use. Selenomethionyl (SeMet) TehA
was expressed in a similar way, but using containing SeMet in place of methionine in
defined minimal media. Cells were resuspended in a buffer containing 50 mM Tris-HCl (pH
8.0) and 200 mM NaCl and lysed using a French Press with two passes at 15-20,000 psi.
Cell debris were removed by centrifugation at 10,000g for 20min, and the membrane
fraction was isolated from that supernatant by ultra-centrifugation at 150,000g for 1 hr.
The membrane fraction was homogenized in a solubilization buffer containing 50 mM Tris
(pH 8.0) and 200 mM NaCl, and incubated with final concentration of 1% (w/v) dodecyl-β-
D-maltopyranoside (DDM, Anatrace, Inc) for 1 hr at 4°C. The non-dissolved matter was
remove by ultracentrifugation at 150,000g for 30 min, and the supernatant was loaded to a
5ml Hitrap Ni2+-NTA affinity column (GE Healthcare), pre-equilibrated with the same
solubilization buffer supplemented with 0.05% DDM. After 20 column volume buffer wash,
the protein was eluted with 250mM imidazole in the solublization buffer. The Flag and 10-
His tags were removed by adding super TEV at 1:100 mass ratio and incubating at 4°C
overnight. Tag removal was confirmed by SDS-PAGE, and the resulting sample was
concentrated to around 10mg/ml. Preparative size exclusion chromatography was carried out
on a Superdex-200 column for further purification, removal of TEV protease and the
cleaved tag, and for buffer and detergent exchange. The gel-filtration buffer contained 10
mM Tris (pH 8.0), 200 mM NaCl, 1mM EDTA, 0.5 mM Tris [2-carboxyethyl] phosphine
(TCEP), and 2×CMC of detergent. In the case of HiTehA, the protein was well behaved and
stable in nearly all tested detergents, and we have purified it from DDM, DM, NM, OM, OG
and LDAO.
Protein characterization
We performed N-terminal amino-acid sequencing of purified HiTehA and EcTehA prior to
TEV protease treatment. Results from these analyses proved that true initiating methionine
residue is located 14 residues after one annotated as N-terminal. The intervening nucleotide
sequence contains a Shine-Delgarno sequence.
For cross-linking experiments, purified HiTehA was incubated with 10 mM disuccinimidyl
glutarate at room temperate for 30 min, and 100mM Tris-HCl pH 7.5 was added to stop the
reaction. The incubated sample when run on a 8-25% gradient SDS-PAGE gel showed a
ladder consistent with a trimeric structure.
Crystallization and data collection
Purified protein was concentrated to ~10mg/ml for initial crystal trials in a Mosquito robot
with commercial screens from Hampton research, Emerald Biosystems and Molecular
Dimension. We obtained crystals of HiTehA from protein in detergent DDM, DM, NM,
OM, OG and LDAO, but only those from LDAO and OG gave diffraction to beyond 4Å
spacings. Crystals that proved useful were all grown at 4°C using the sitting-drop vapor
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diffusion method. After extensive optimization we reached conditions supporting very high
resolution. Wild-type HiTehA and most of its variants crystallized from 1mM ZnSO4,
50mM Hepes-Na pH 7.8 and 28% PEG600. HiTehA F262A crystallized from 200mM
Li2SO4, 100mM glycine pH9.3 and 33% PEG400). Addition of 10mM spermidine as an
additive helped to obtain slightly better diffracting crystals. Cryoprotection was achieved by
adding 5% ethylene glycol or PEG400 to the crystallization solution.
Structure determination and refinement
Native and SeMet SAD data set were collected at NSLS beamline X4A and processed using
the software HKL200052. Crystals of HiTehA grew in space group R3 with a=b=c = 85.0Å
and α=β=γ = 93.5°. All subsequent manipulations were done in the hexagonal setting of this
space group with a=b= 96.0Å and c = 136.7Å. The asymmetric unit contains one TehA
protomer and 65% solvent volume. The structure was determined at 2.0Å resolution from
single-wavelength anomalous diffraction (SAD) using selenomethionine-substituted protein
crystals. Assessment of data quality for phasing, location of heavy atom sites and initial
phases were calculated using the HKL2MAP interface to SHELX programs53.
All the secondary structure elements were clearly visible in the experimental electron
density map. Automatic model building was done in Arp/wArp54 and completed manually
in the program COOT55. The model was refined against native data at 1.20Å resolution
using the program Refmac5.5 in CCP456, with anisotropic B factor restrained refinement
applied. Subsequent structural analyses of mutant variants were refined as isomorphous
structures.
Site-directed mutagenesis
Site-directed mutants were constructed using the QuikChange Site-Directed Mutagenesis Kit
(Stratagene, La Jolla, CA 92037) and expressed from pET vectors in E. coli BL21(DE3)
plysS cells as for the wild-type protein.
Electrophysiology
All constructs were cloned into plasmid pGHME2, linearized, and transcribed into cRNA
using T7 polymerase (mMessage mMachine, Ambion). Oocytes were injected with 50 nl of
cRNA solution each, at a constant concentration of 0.5 mg/ml for HiTehA, or 0.5 mg/ml for
AtSLAC1 constructs, with or without 0.5 mg/ml of AtOST1 constructs and recorded in
voltage-clamp experiments 2 days after injection. For mixed expression experiments, 25nl of
cRNA solution was injected for each AtSLAC1 component. Two-electrode voltage clamp
recordings were performed to measure HiTehA or AtSLAC1 currents as described6,7. The
pipette solutions contained 3 M KCl. For voltage-clamp current recordings, the bath solution
contained 1 mM MgCl2, 1 mM CaCl2, 10 mM Mes/Tris (pH 5.6) and 30 mM CsCl; For
anion selectivity measurements, the bath solutions contained 50 mM Cl−, NO −3, malate, or
30 mM SO 2-3 sodium salts, 45 mM Na-gluconate, 1 mM Ca-gluconate2, 1 mM Mg-
gluconate2, 1 mM K-gluconate, as well as 10 mM Tris/Mes (pH 5.6). Osmolality was
adjusted with D-mannitol to 220 mOsmol/kg. The bath electrode was a 3M KCl agar bridge.
The testing potentials were +50 to −110 or −130 mV in 20 mV decrements with 7.5 s
duration. The holding and return potentials were 0 mV with a 1.45 s or 2 s, respectively. I-V
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relations of HiTehA or AtSLAC1 were generated from the current reached after 0.5 s in
testing potentials. The Goldman-Hodgkin-Katz equation was applied to estimate
permeability ratios for monovalent ions as described6. For divalent anions, the permeability
ratios were derived according to Fatt and Ginsborg57.
Bioinformatic analysis of SLAC-related proteins
Sequences related to SLAC1 were analyzed comprehensively by PSI-BLAST50. Searches at
E<10−3 starting from five disparate homologs each identified a common pool of over 900
proteins, which when pooled were used for sub-classification into families and subfamilies.
Details of these analyses are reported in footnotes to Table S1.
Molecular figures were produced in PyMOL58.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We thank Filippo Mancia, Larry Shapiro and Ming Zhou for helpful discussions; Patricia Rodriguez and Arianne
Morrison for help with cloning and expression; Jonah Cheung, Mark Collins, Chris Min, Sheng Ye and Zhen Zhang
for advice in protein chemistry and crystallography; John Schwanof and Randy Abramowitz for help with
synchrotron experiments; John Riley for help with Xenopus oocyte injections; and Min Su for advice on the PSI-
BLAST analysis of SLAC relatives. This project was supported in part by an award to the New York Consortium
on Membrane Protein Structure (NYCOMPS) from the NIGMS Protein Structure Initiative. Beamline X4A at the
National Synchrotron Light Source (NSLS), a DOE facility at Brookhaven National Laboratory, is supported by the
New York Structural Biology Center.
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58. DeLano, WL. The PyMOL molecular graphics system. DeLano Scientific; San Carlos, CA, USA:
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Figure 1. Sequence analysis for the SLAC1 superfamily
a, Family tree. The presentation was computed by the program COBALT47 from
representative subfamily sequences (Table S1), including Arabidopsis thaliana SLAC1 for
SF1Ai, Haemophilus influenzae TehA for SF1Ci, Escherichia coli TehA for SF1Cii, Vibrio
parahaemolyticus for SF2A, Staphylococcus aureus for SF2B, Aspergillus fumigatus Ssu1
for SF3A, and Schizosaccharomyces pombe Mae1 for SF3B. b, Structure-based sequence
alignment of TehA from Haemophilus influenzae (HiTehA) and SLAC1 from Arabidopsis
thaliana (AtSLAC1). The TehA structure has been used to restrict sequence gaps to inter-
helical segments. Superior coils define extents of the HiTehA helical segments; red letters
mark residue identities; red boxes are drawn for residues that are >95% identical within the
plant subfamily SF1A for AtSLAC1 or within the TehA subfamily for HiTehA; red
diamonds mark HiTehA residues that line the central pore; and the colored inferior bar
encodes ConSurf sequence variability48 for the SF1 family of 204 non-redundant proteins.
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Figure 2. Crystal structure of HiTehA and homology model of AtSLAC1
a, Electron density distribution from the HiTehA crystal structure at 1.2Å resolution. The
map has (2Fo-Fc) coefficients based on the superimposed model. Contours are at 2.5σ. b,
Ribbon diagram of the HiTehA trimer. Each protomer is colored spectrally from blue at its
N-terminus to red at its C-terminus. c, DelPhi49 electrostatic potential at the extracellular
surface. Electronegative and electropositive potential are colored in degrees of red and blue
saturation, respectively. d, Electrostatic potential49 at the intracellular surface. e, Ribbon
diagram of an HiTehA protomer viewed from outside the membrane. The ribbon is colored
spectrally as in 2b. f, Ribbon diagram of an HiTehA protomer viewed from within the
membrane from below the view of e. g, Surface of a homology model of AtSLAC1, viewed
as in f, and colored by sequence variability48. h, Surface of AtSLAC1 as in g, but colored by
electrostatic potential49.
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Figure 3. Putative structure of the SLAC1 conductance pore
a, Cross-section through the homology model of AtSLAC1. The model is viewed as in 2i,
with the electrostatic potential49 shown on the external surface of the molecular envelope.
The side chain of Phe450 is shown as a stick model (red) on the backbone ribbon colored
yellow. b, Cross-section as in a, but colored by surface conservation48 as in 2h. c, Pore-
lining residues in the SLAC1 homology model. A cylinder model (left), spectrally colored as
in 2b, provides a key for viewing the rolled-open structure (left) with pore-lining residues of
AtSLAC1 shown on the TModd helices. d, Ribbon diagrams of HiTehA TM9 (left) and TM7
(right) viewed from within the conductance pore. The side chains of Pro207 and Phe262 are
shown as well as a kink-stabilizing water HOH25 that is coordinated by the NH of Gly263
and by C=O groups of Gly202 and Ala259. Density contours are shown for the water
molecule.
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Figure 4. Ionic conductance measurements
a, Typical microelectrode voltage-clamp current traces from oocytes injected with various
channel cRNAs. Left column, oocytes injected with cRNAs encoding wild-type HiTehA
channels (WT), or F262A, G15D/F262A or G15D mutants. Right two columns, oocytes
injected with cRNAs for WT AtSLAC1, or F450A, G194D/F450A or G194D mutants, with
or without co-injection of AtOST1. Dotted lines represent zero current levels. Extracellular
solution contained 30mM CsCl. Schematic keys at the left place phenyl and/or aspartyl
gates. b, Effects of gating residue mutations. Mean chloride currents, measured at −90 mV,
are shown comparing WT HiTehA and its mutant series F262A,G,T,V,L with WT AtSLAC1
and its corresponding series F450A,G,T,V,L, both alone and co-expressed with AtOST1.
Full I-V relations are shown in Fig. S5. c, Effect of gating residues on relative AtSLAC1
anion permeabilities. Relative permeabilities (P[X]/P[Cl]) for chloride, nitrate, sulfite and
malate of WT, F450A and F450T SLAC1 channels were measured from the change in
current reversal potential with Cl− or anion X− as the sole permeant anion in the bath
solution (Methods, Table S6).
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Figure 5. Structural features at the SLAC1 homolog gate
a,b, Cross-sections through the conductance pores of HiTehA F262A and HiTehA G15D.
The view and presentations are as in 3a, except that helices are colored purple. c, Molecular
basis for conformational strain in gating residue Phe262 of HiTehA. Helices TM1 (left),
TM9 (center) and TM7 (right) are viewed from within the pore and presented as ribbon
diagrams with selected side chains drawn in stick representation. The local low-energy
conformation for the phenyl ring (χ2 = 90°) is shown in thin lines with short contacts
indicated in dashed lines: Leu18 Cδ2 – Phe262 Cδ1, d = 2.4 Å; Val210 Cγ2 – Phe262 Cε2, d
= 2.8 Å. d, Conformational shifts consequent to release of strain in gating residue Phe262.
Cα backbone structures of F262A, F262V and F262L HiTehA were first superimposed onto
WT HiTehA, and residues 258-266 were drawn in stereo as superimposed. All backbone
atoms are shown for peptides 262±1, but only Cα atoms are shown otherwise. The WT
backbone and phenyl group are green; other backbone are all magenta; side chains of
Ala262, Val262 and Leu262 are cyan, blue and red, respectively; oxygen-directed bonds are
red and nitrogen-directed bonds are blue. e, Typical microelectrode voltage-clamp current
traces from oocytes injected with wild-type (WT) or F450L AtSLAC1 cRNA. Experimental
conditions and displays are as in 4a.
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|
3M79
|
A tetrameric Zn-bound cytochrome cb562 complex with covalently and non-covalently stabilized interfaces crystallized in the presence of Cu(II) and Zn(II)
|
Evolution of Metal Selectivity in Templated Protein Interfaces
Jeffrey D. Brodin†, Annette Medina-Morales†, Thomas Ni†, Eric N. Salgado†, Xavier I.
Ambroggio‡, and F. Akif Tezcan†,*
† Department of Chemistry and Biochemistry, University of California, San Diego, La Jolla, CA
92093-0356
‡ Rosetta Design Group LLC, Fairfax, VA 22030
Abstract
Selective binding by metalloproteins to their cognate metal ions is essential to cellular survival.
How proteins originally acquired the ability to selectively bind metals and evolved a diverse array
of metal-centered functions despite the availability of only a few metal-coordinating
functionalities remains an open question. Using a rational design approach (Metal-Templated
Interface Redesign), we describe the transformation of a monomeric electron transfer protein,
cytochrome cb562, into a tetrameric assembly (C96RIDC-1) that stably and selectively binds Zn2+,
and displays a metal-dependent conformational change reminiscent of a signaling protein. A
thorough analysis of the metal binding properties of C96RIDC-14 reveals that it can also stably
harbor other divalent metals with affinities that rival (Ni2+) or even exceed (Cu2+) those of Zn2+
on a per site basis. Nevertheless, this analysis suggests that our templating strategy also introduces
an increased bias towards binding a higher number of Zn2+ ions (4 high affinity sites) versus Cu2+
or Ni2+ (2 high affinity sites), ultimately leading to the exclusive selectivity of C96RIDC-14 for
Zn2 over those ions. More generally, our results indicate that an initial metal-driven nucleation
event followed by the formation of a stable protein architecture around the metal provides a
straightforward path for generating structural and functional diversity.
Introduction
The incorporation of metal ions into correct cellular targets is a formidable chemical task.
Modern-day organisms use a number of strategies to ensure that a metal ion associates with
the right target (frequently a protein), including the control of absolute and relative ambient
metal concentrations,1 active delivery via chaperones,2 and compartmentalization.3
Nevertheless, these strategies still require the target protein to possess an intrinsic affinity
and selectivity for the desired metal ion. This is achieved within the 3-D framework of
proteins despite the availability of only a handful of metal-coordinating side chain
functionalities.
An outstanding question is how protein structures have evolved to stably and selectively
bind metal ions and developed metal-dependent functions, such as signal transduction
(which necessitates conformational flexibility) and electron transfer or catalysis (which
generally require rigid architectures). Some contemporary metalloproteins likely were based
on pre-existing protein folds that acquired the ability to bind metals through random genetic
events and subsequently attained their current structures and functions in the course of
tezcan@ucsd.edu.
Supporting Information Available. Additional experimental details, figures and tables on protein preparation, crystallography,
analytical ultracentrifugation and binding assays. This material is available free of charge via the Internet at http://pubs.acs.org.
NIH Public Access
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Published in final edited form as:
J Am Chem Soc. 2010 June 30; 132(25): 8610–8617. doi:10.1021/ja910844n.
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natural selection.4 In an alternative pathway, metal ions could first have templated the
formation of a protein/peptide aggregate, followed by the evolution of the protein structure
around the metal ion.5 Although this pathway would not have the benefit of re-using
existing genetic material and a preformed scaffold, it may lead to greater flexibility towards
generating bioinorganic diversity. We have developed a rational design approach (Metal-
Templated Interface Redesign, MeTIR), which follows the time course of this latter
hypothetical pathway (Scheme 1).6 Using MeTIR, we describe here the construction of an
oligomeric protein architecture– C96RIDC-14–that stably and selectively binds Zn2+, and
displays a large, metal-dependent conformational change akin to signaling/regulatory
proteins.
Results and Discussion
Design Strategy and Initial Characterization of C96RIDC-14
We have previously shown that variants (termed MBPC’s) of a four-helix-bundle heme
protein, cytochrome cb562, with appropriately installed metal-binding motifs on their
surfaces (represented by Species 2 in Scheme 1) can adopt discrete supramolecular
architectures upon metal coordination.7–9 Because the parent cyt cb562 (1) is a monomeric
protein whose surface carries no bias towards oligomerization, its supramolecular assembly
is largely dictated by metal coordination. Accordingly, all MBPC assemblies initially
formed through metal binding (3) feature interfacial metal ions with saturated coordination
spheres and ligand arrangements that obey the preferred stereochemistry of the metal ions:
Zn2+ – tetrahedral, Cu2+ – tetragonal, Ni2+ – octahedral.9 If the protein assembly
surrounding a particular metal ion can be stabilized without changing the overall
supramolecular architecture, one would expect the binding affinity and specificity to
increase for that metal ion. We decided to put this “template-and-stabilize” strategy to the
test using a tetrameric Zn-mediated assembly, Zn4:MBPC-14 (Figure 1).
MBPC-1 (2) is a cyt cb562 variant with two i, i+4 bis-His metal-binding motifs (H59/H63
and H73/H77) installed on its Helix3. Upon Zn coordination MBPC-1 forms a D2 (222)
symmetrical tetramer, Zn4:MBPC-14, which is held together by four identical Zn ions
coordinated to one bis-His motif (73/77) from one protomer, His63 from a second, and
Asp74 from a third.7 Owing to its twofold dihedral symmetry, Zn4:MBPC-14 presents three
pairs of C2-symmetric interfaces (i1, i2, i3) between its four protomeric constituents. Of
these interfaces, only i1 presents an extensive surface (>1000 Å2) with close protein-protein
contacts. We therefore undertook computationally-guided redesign of i1 to examine if a
favorable set of interactions can be built into i1 to stabilize the entire Zn-driven assembly
(Step b in Scheme 1).
A construct, RIDC-1, which features six mostly hydrophobic mutations (R34A/L38A/
Q41W/K42S/D66W/V69I) in i1, indeed was found to form a considerably stabilized
tetrameric assembly (Zn4:RIDC-14, 4) with an identical supramolecular geometry to the
parent tetramer.6 Despite this stabilization, Zn4:RIDC-14 (like Zn4:MBPC-14) remains a
dynamically exchanging assembly that does not stay intact, for instance, upon passage
through a size-exclusion column. Consequently, it has not been possible to uncouple protein
oligomerization from Zn binding and directly assess whether interface redesign has led to
improvements in Zn affinity and selectivity. In order to obtain a stable tetrameric complex
that would also form in the absence of Zn, we sought to further engineer RIDC-1.
As the dihedral symmetry of the Zn-tetramer dictates, the concurrent stabilization of any two
of the three interfaces could in principle lead to the formation of a persistent tetrameric
assembly. While interface i2 is not as tightly packed as i1 and therefore is less amenable to
redesign, it presents position 96 from two protomers within distance for disulfide (SS)
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crosslinking (Figure 1). Thus, we combined the six mutations in i1 with the T96C mutation
in i2 to generate C96RIDC-1. Under oxidizing conditions C96RIDC-1 readily forms an SS-
crosslinked dimer as shown by SDS-PAGE electrophoresis (Figure S1). Sedimentation
velocity (SV) experiments indicate that the predominant oligomeric form of C96RIDC-1 in
solution is a tetramer (C96RIDC-14, 6), even in the absence of metals (Figure 2a). The
tetramer-dimer dissociation constant (Kd(4mer-2mer)) for C96RIDC-14 has been determined by
sedimentation equilibrium (SE) measurements to be <100 nM (Figure 2b), which to our
knowledge makes it one of the most stable engineered protein complexes.
Zn-dependent changes in C96RIDC-14 conformation
C96RIDC-14 remains tetrameric upon Zn binding. SV population distributions indicate that
the resulting tetramer (5) forms at significantly lower protein and Zn concentrations and
therefore is more stable than its progenitors Zn4:MBPC-14 and Zn4:RIDC-14 (Figure 2a).
SV measurements also suggest that C96RIDC-14 undergoes a Zn-induced rearrangement,
evidenced by a shift in its sedimentation coefficient from 4.25 S to 4.5 S. To elucidate this
conformational change, we solved the crystal structures of C96RIDC-14 and its Zn adduct at
2.1 and 2.4-Å resolution, respectively (PDB ID’s 3IQ5 and 3IQ6).
As illustrated in Figure 3, these structures reveal a remarkable double-clothespin motion of
the four protomers upon Zn-coordination, measuring ~16 Å at the N-terminus of Helix3.
One of the keys to the simultaneous stability and conformational plasticity of C96RIDC-14
lies with the redesigned interface i1. Each of two equivalent i1 interfaces in C96RIDC-14 is
formed between two protomers oriented in an antiparallel fashion. These interfaces feature
an extensive network of hydrophobic interactions (~1300 Å2 buried surface), the main
contributors being the two pairs of engineered Trp (41 and 66) and His (59 and 77) residues
(Figure S3). Not surprisingly, this hydrophobic interface was predicted and found to be
rather fluid.6 Upon binding Zn, the fluidity of the engineered hydrophobic interactions
allows the four protomers to pivot around i1 and undergo the double-clothespin motion. The
resulting architecture of Zn4:C96RIDC-14 features a well-packed hydrophobic core in i1
(~1400 Å2 buried surface) built around the engineered residues originally proposed by
computation. Zn4:C96RIDC-14 is superimposable onto both Zn4:MBPC-14 and Zn4:RIDC-14
structures with respective root-mean-square deviations of 0.59 and 0.63 Å over 424 Cα’s
(Figure S4). Importantly, the four equivalent Zn coordination environments remain
unchanged as intended by the template-and-stabilize strategy.
The interfacial SS bonds incorporated into i2 are the second key component for the
bistability of C96RIDC-14. The electron density maps for C96RIDC-14 and Zn4:C96RIDC-14
clearly outline the C96-C96′ linkages, which are found in distinct conformations to
accommodate the two different supramolecular arrangements (Figure 3). Using the SS-
bonds as “loose hinges”, the pairs of protomers that share i2 undergo a significant
translational and rotational motion relative to one another, while maintaining the ideal bond
distance (2.05 Å) and the stereochemical requirements for an SS bond. Operating together,
the fluid non-covalent interactions in i1 and the adaptable SS bonds in i2 allow the
formation of two stable and interconvertible tetrameric architectures in the absence or
presence of Zn.
Zn binding by C96RIDC-14
Having thus uncoupled protein oligomerization from metal binding, we examined if our
interface templating strategy leads to increased Zn affinity. Because Zn is spectroscopically
silent, its binding to C96RIDC-14 was assessed through two different indirect methods. In the
first method modeled after a procedure used for determining uranyl binding to NikR,10 we
used nitrilotriacetic acid (NTA) as a competing Zn ligand (Kd(Zn-NTA) = 18.2 nM at 22°C
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and μ = 0.15 M) and 4-(2-pyridylazo)resorcinol (PAR) as an indicator, which exhibits an
increase in absorbance at 500 nm upon binding Zn.11 Briefly, increasing amounts of NTA
were added to samples of nearly equimolar Zn and C96RIDC-14. The resulting mixtures
were subjected to centrifugation in 10-kDa-cutoff protein concentrators, which allowed the
separation of the Zn fraction bound to C96RIDC-14 from that bound to NTA at each NTA
concentration. This, in turn, permitted the calculation of the extent of Zn binding
to C96RIDC-14 versus free Zn concentration, which was inferred from the Zn-NTA binding
equilibrium. The NTA titrations are reasonably well described by an n-equivalent binding
sites model, and indicate that each C96RIDC-14 complex binds four Zn equivalents with an
apparent dissociation constant of 3.3 nM (Figure 4a, see Figure S9 for fits to alternative
binding models).
In a second, “higher-resolution” method that obviates the separation step and the secondary
indicator, we used the chromophore Fura-2 (Kd, Zn = 5.7 nM)12 as a competing Zn ligand/
indicator in a similar fashion to previously published procedures.13 In these experiments,
increasing amounts of Zn were added to samples that contained C96RIDC-14 and Fura-2, and
the Zn-dependent changes in the Fura-2 spectrum were directly monitored (Figure 4b).
These titrations reveal that Zn binding to C96RIDC-14 is best described by four individual,
consecutive binding equilibria or nearly equally well by two consecutive binding events by
pairs of Zn2+ ions, with corresponding dissociation constants that range from 0.5 nM to 60
nM in the case of the 1+1+1+1 model and from 0.5 to 40 nM in the case of the 2+2 model
(Table 1).14
Owing primarily to its flexibility, C96RIDC-14 apparently can accommodate Zn binding
through several different modes; in other words, the intermediate Zn bound states may and
most likely do utilize different ligand sets from one another. In any case, given that i, i+4
bis-His motifs on α-helices display Zn dissociation constants in the low μM range,15 the Zn
binding titrations suggest that the pre-formation of a tetrameric, templated acceptor complex
results in a ≥1000-fold increase in Zn-binding affinity relative to the monomeric parent
species, MBPC-1.
Zn binding selectivity of C96RIDC-14 over other divalent metal ions
To elucidate if C96RIDC-14 also displays increased selectivity for Zn binding, its
interactions with several other divalent metal ions (M2+) were examined, including the
neighboring Co2+, Ni2+ and Cu2+, which typically are effective competitors for Zn binding
sites. To this end, C96RIDC-14 was incubated with the metal ion of interest in a
noncoordinating buffer solution (20 mM 3-(N-morpholino)propanesulfonic acid, MOPS),
followed by the separation of the C96RIDC-14-metal complex via gel filtration and
subsequent metal analysis by ICP-OES. These experiments show that C96RIDC-14 retains
~1 equivalent of Co2+, ~2 equivalents of Ni2+ and ~4 equivalents of Cu2+ (Figure S5a).
In competition studies, the initial C96RIDC-14/metal mixture was additionally incubated
with Zn at various metal/Zn ratios prior to gel filtration and metal analysis. Each
competition experiment was also carried out in reverse order - incubation with Zn followed
by addition of other metals - to ascertain the formation of thermodynamic products (Figure
S5b). These experiments reveal that C96RIDC-14 displays significant Zn selectivity over all
ions except Cu2+, especially relative to its parent structure, MBPC-1 (Figures 5, S5b and
S7). Previous studies have shown the affinity of the i, i+4 bis-His motif for Zn2+ to be
comparable to that for Ni2+ and 5–10 fold higher than that for Co2+,15,16 following the
Irving-Williams(IW) series.17 In contrast, Zn2+ completely outcompetes Co2+
for C96RIDC-14 binding at all ratios measured (up to 100 Co:1 Zn), and has an effective
affinity roughly 100-fold higher than Ni2+.
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Cu2+ presents a special case in terms of Zn selectivity. Due to a combination of its d9
configuration and high Lewis acidity, Cu2+ is situated at the top of the IW series, leading to
its higher affinity for most ligand platforms designed for specific Zn binding13,18 and even
natural Zn enzymes.19 The results shown in Figure 5 initially indicated that neither Cu2+
nor Zn2+ outcompete each other for C96RIDC-14 binding; rather, each
tetrameric C96RIDC-14 unit appeared to bind ~3 equivalents of each ion in the non-
coordinating MOPS buffer solution. To ascertain whether this apparent oversaturation
of C96RIDC-14 is due to the binding of Cu2+ or Zn2+ ions to the C96RIDC-14 surface (in
addition to the core binding sites) or due to the formation of an actual Cu-Zn heterometallic
core species, we obtained crystals of C96RIDC-14 grown in the presence of equimolar
amounts of both ions. The resulting 2.1-Å resolution diffraction data reveal a structure (PDB
ID: 3M79) identical to that of Zn4:C96RIDC-14. To identify the four observed interfacial
metal ions in this complex, we collected full data sets at the Zn and Cu K edges (1.28 Å and
1.38 Å), respectively. The corresponding anomalous difference maps clearly show that the
interfacial ions are Zn2+, and that there are no detectable Cu2+ ions associated with the core
or the surface of the tetramer (Figures 6a and b), despite the fact the crystal clearly contained
Cu as indicated by an X-ray fluorescence excitation scan (Figure 6c). These observations
suggest that Zn2+ outcompetes Cu2+ completely for binding to the core sites. This is further
supported by the finding that when the Zn-Cu competition experiments are carried out in a
weakly coordinating buffer solution (20 mM Tris(hydroxymethyl)aminomethane, TRIS), the
amount of Cu2+ associated with C96RIDC-14 is significantly diminished, whereas the
amount of bound Zn2+ stays constant (see the last two rightmost bars in Figure 5).
To describe the Zn selectivity of C96RIDC-14 in a more quantitative fashion, we examined
its affinity for Co2+, Ni2+ and Cu2+, again using Fura-2 as a competing ligand (Kd, Fura-Co =
8.6 nM,20 Kd, Fura-Ni = 6.9 nM,12 Kd, Fura-Cu = 0.3 pM12). The titrations indicate
that C96RIDC-14 has one weak Co2+ binding site that barely competes with Fura-2 (Figure
S11 and Table 1), whereas it can accommodate two equivalents of Ni2+ and Cu2+ with
comparable affinities to Fura-2 (Figure 7 and Table 1).21 Both Ni2+ and Cu2+ binding
curves are well described by two consecutive binding equilibria. Although on a per site basis
the derived affinities are either similar to those for Zn2+ (in the case of Ni2+) or considerably
higher (in the case of Cu2+) in line with the IW series, the higher multiplicity for Zn2+
binding apparently results in a more favorable overall free energy (Table 1), and ultimately
in the Zn selectivity of C96RIDC-14 over these ions.21 Thus, templated interface engineering
leads to increased bias not only towards Zn coordination geometry but also towards Zn
binding multiplicity, which, to our knowledge, is a rare, and perhaps unique, case in
designed/synthetic systems. Studies are currently underway to elucidate the coordination
modes/environments of Co2+, Ni2+ and Cu2+ to C96RIDC-14 (which we expect to be
different from each other and from Zn2+) and associated changes in protein structure.
Conclusions
Metal-templated synthesis has been a powerful approach to construct ligands with enforced
coordination geometries that provide stable and specific metal binding.22–24 We have
shown here that such templating strategies used for smaller systems can also be applied to
proteins, which, owing to their extensive surfaces rich in chemical functionality, allow the
formation of an extensive network of covalent and non-covalent interactions around the
template. Importantly, the templating of protein interfaces around the target metal ion, Zn2+,
has led to the evolution of a flexible, multi-14 stable protein complex that not only presents
an increased bias towards the tetrahedral Zn2+ coordination geometry, but also towards the
Zn2+ binding multiplicity of four, ultimately resulting in significantly increased selectivity
over other divalent ions including Cu2+. It is well established that Cu2+ can readily
outcompete Zn2+ (or any other physiologically important metal ion) for binding even rigid,
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tetrahedral coordination environments that favor Zn2+.19 Such thermodynamic dominance
by the Cu2+ ion indeed necessitates various cellular strategies – often operating under
kinetic control – to be employed for the incorporation of other metal ions into their intended
protein targets.25 Our findings suggest that control of metal binding multiplicity may be a
viable thermodynamic strategy alongside the design and restraint of the inner-sphere
coordination environment to enhance metal selectivity in natural or synthetic systems.
Our study further demonstrates that cyt cb562 – a monomeric, putative electron transfer
protein – can be transformed into a Zn-responsive complex through a minimal number of
mutations that amount to less than 10% of its amino acid sequence. It is conceptually
straightforward to envision how the Zn4:C96RIDC-14 architecture can be further rigidified
and modified around the Zn centers to promote metal-based functions such as Lewis acid
catalysis. While the role of metal templating in the evolution of metalloproteins can only be
postulated, MeTIR clearly provides a practical route to generating structural and functional
diversity. It remains to be seen if this approach can also be extended to non-metallic
substrates, which could have served as nucleants in the early emergence of protein folds.
26,27
Experimental Section
Site-Directed Mutagenesis and Protein Expression/Purification
The T96C mutation was introduced into pET-ridc16 using QuikChange (Stratagene) site-
directed mutagenesis and primers obtained from Integrated DNA technologies, yielding the
expression vector pET-C96ridc1. pET-C96ridc1 was transformed into XL-1 blue E. coli cells,
purified using the QIAprep Spin Miniprep Kit (QIAGEN) and sequenced by Retrogen.
pET-C96ridc1 was transformed into BL21(DE3) E. coli cells along with the ccm heme
maturation gene cassette plasmid, pEC86.28 Cells were plated on LB agar containing 100
μg/ml ampicillin and 34 μg/ml chloramphenicol and grown overnight. 3 mL starter cultures
were inoculated from the resulting colonies, grown to an Abs600 of 0.6 and used to inoculate
1 L of LB medium. 1-L cultures were then incubated for 16 hours with rotary shaking at 250
rpm. No induction was necessary.
Protein was obtained by sonicating cells in the presence of lysozyme, bringing the lysate to
pH 5 with HCl and isolating the soluble fraction by centrifugation at 16,000 g, 4° C, for 1 hr.
Initial purification was by cation-exchange chromatography on a CM-Sepharose matrix
(Amersham Biosciences) using an NaCl step gradient in sodium acetate (pH 5). After
dialysis into sodium phosphate (pH 8), the protein was further purified by anion exchange
on an Uno-Q (BioRad) column using a DuoFlow chromatography workstation (BioRad) and
a linear NaCl gradient. A protein sample was then exchanged into water, mixed 1:1 with
sinapinic acid matrix (Agilent Technologies) and subjected to MALDI mass spectrometry to
verify the T96C mutation (expected mass 12305 amu, observed 12299 amu).
Dimeric C96RIDC-12 was separated from monomeric protein on a preparative scale size
exclusion chromatography column (GE Healthcare) packed with Superdex 75 (GE
Healthcare) resin equilibrated in 20 mM Tris HCl (pH 7) and 150 mM NaCl. Separation of
dimer (which effectively is a tetramer at concentrations > 1 μM) from monomer was verified
by non-reducing SDS-PAGE gel electrophoresis using a 15% acrylamide gel. Typical
protein preparations yielded 8–10 mg of protein per liter of culture based on an extinction
coefficient of 148,000 M−1cm−1 at 415 nm. After purification, dimeric C96RIDC-12 was
concentrated to ~2 mM using an Amicon stirred cell (Millipore), flash frozen in liquid
nitrogen and stored at −80° C.
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Analytical Ultracentrifugation
SV and SE samples were prepared in 20 mM Tris (pH 7) and 150 mM NaCl using
appropriate volumes of 5 mM metal stock solutions or 50 mM EDTA. After incubating in
the presence of M2+ or EDTA for 1 hour, SV measurements were carried out at 25° C on a
Beckman XL-I Analytical Ultracentrifuge using an An-60 Ti rotor at 41,000 rpm for a total
of 250 scans per sample. The following wavelengths were used to monitor C96RIDC-12
sedimentation at different protein concentrations: 415 nm (5 μM), 440 nm (30 μM). All data
were processed using SEDFIT29 software with the following fixed parameters: buffer
density (r) = 1.0049 g/ml; bufferviscosity = 0.010214 poise; Vbar = 0.73084.
SE measurements were carried out at 25° C using speeds between 10,000 and 20,000 rpm.
Scans were taken at 14 and 16 hours and visually inspected to verify that sedimentation
equilibrium was achieved. The following wavelengths were used to monitor C96RIDC-12
sedimentation at different protein concentrations: 415 nm (1 μM), 420 nm (2.5 μM), and 500
nm (12.5 μM). 16-hr scans were fit to a monomer-dimer (where monomer = C96RIDC-12) or
a dimer-only model using SEDPHAT.30 The molecular mass of C96RIDC-12 (24610 Da)
and the menisci were fixed while floating the association constant. Standard deviation for
the resulting log10(K) value was determined through Monte-Carlo analysis within
SEDPHAT.30
Crystallography
All crystals were grown by sitting drop vapor diffusion at room temperature (20–25° C) in
drops consisting of 2 μL of protein and 1 μL of precipitant solution. For apo and Zn2+
crystals, a 2.1 mM protein stock solution in 20 mM Tris (pH 7) and 150 mM NaCl was used.
The precipitant solution for apo-C96RIDC-14 crystals consisted of 100 mM Bis-Tris (pH 6.5)
and 30% PEG 400. The precipitant solution for Zn4:C96RIDC-14 crystals was 100 mM Tris
(pH 7.5), 20% PEG 2000 and 2.5 mM ZnCl2. Crystallization of C96RIDC-14 in the presence
of Zn and Cu (Zn/CuC96RIDC-14) was performed by incubating 60 μM protein with one
molar equivalent of Zn for one hour followed by addition of one equivalent of Cu and
incubation overnight to ensure the formation of the thermodynamic product. The mixture
was then concentrated to 2.6 mM using 4 mL Amicon Ultra (Millipore) centrifugal filters.
Crystallization was performed as above with a precipitant solution consisting of 24% PEG
2000 and 100 mM Bis-Tris (pH 6.5). Crystals used for diffraction were exchanged stepwise
into a solution containing 20% glycerol as a cryoprotectant and frozen in liquid nitrogen or
directly in the cryostream.
Diffraction data were collected at the Stanford Synchrotron Radiation Laboratory (SSRL)
Beamline 9-2 for C96RIDC-14 and Zn4:C96RIDC-14 or Beamline 7-1 for Zn/CuC96RIDC-14
at 100 K. 1.0-Å radiation datasets were collected for all crystals and additional datasets were
collected using 1.28-Å and 1.38-Å radiation, corresponding to Zn and Cu K edges,
respectively, for Zn/CuC96RIDC-14. The data were processed using MOSFLM and SCALA.
31 The structures of C96RIDC-14, Zn4:C96RIDC-14 and Zn/CuC96RIDC-14 were determined
at 2.05, 2.35 and 2.1 Å resolution, respectively, by molecular replacement with MOLREP,32
using the RIDC-1 monomeric structure (PDB ID: 3HNI) as the search model. Rigid-body,
positional and thermal refinement with CNS33 or REFMAC,34 along with manual model
rebuilding and water/ligand placement with XFIT35 or COOT36 produced the final models.
For all structures, non-crystallographic symmetry (NCS) restraints (tight main-chain and
medium side-chain restraints) were applied throughout the positional/thermal refinement
process. The Ramachandran plots were calculated with PROCHECK.37 All figures were
produced with PYMOL (www.pymol.org). Data collection and refinement statistics are
summarized in Table S1.
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Inductively-coupled plasma-optical emission (ICP-OES) spectroscopy
ICP-OES samples were in 20 mM MOPS (pH 7) and 150 mM NaCl or 20 mM Tris (pH 7)
and 150 mM NaCl. To determine the stoichiometry of metal binding to C96RIDC-14, 60 μM
protein was incubated with one or two molar equivalents of the desired metal ion. For
competition studies, 60 μM protein was incubated with one molar equivalent of Zn2+ for one
hour, after which, one, ten or one hundred molar equivalents of competing metal (M2+) were
added, bringing the final volume to 1 ml. The same experiments were also done in reverse
order. All samples were allowed to equilibrate overnight at room temperature and free/
loosely bound metal was subsequently removed using a 10DG gel filtration column (Bio-
Rad). Samples were prepared for ICP-OES by diluting 1.3 mL of protein solution collected
off the 10DG column to a final volume of 2 ml and adding 90 μl of 69% reagent grade nitric
acid (Fluka) to achieve a final concentration of 3%. Standards were prepared from 1000 ppm
certified ICP-OES metal stock solutions (Ricca) by mixing equal volumes of all metal
analytes and diluting to a final concentration of 200 ppm of each metal. A standard curve
with eleven points between 0.05 and 10 ppm was then constructed by diluting appropriate
volumes of the 200-ppm stock to 10 ml with 3% nitric acid in deionized water. Data were
collected on a Perkin-Elmer Optima 3000 DV ICP-OES spectrometer located at the
Analytical Facility of the Scripps Institute of Oceanography. Because each C96RIDC-1
monomer contains one Fe atom as part of the covalently linked heme molecule, the
experimentally determined M(II):Fe ratios directly yielded the M2+:protein ratios.
Wavelengths used for the detection of various metal ions were as follows: Mg (279.077,
280.271 and 285.213 nm), Ca (315.887 and 317.993), Fe (234.349, 238.204, 239.562,
259.939 and 273.055 nm), Co (228.616 and 238.892 nm), Ni (221.648 and 231.604 nm), Cu
(224.7, 222.778, 221.459, 327.393 and 324.752 nm), and Zn (202.548, 206.2 and 213.857
nm). Values reported for each metal are averages of those for all wavelengths indicated.
Competitive binding assays using nitrilotriacetic acid (NTA)
The stability constant for Zn2+ binding to C96RIDC-14 was determined based on a
previously published protocol for the determination of uranyl binding to NikR,10 with the
exception that NTA was substituted as the competing ligand. Titrations were performed in
20 mM MOPS (pH 7) and 150 mM NaCl at 22° C. Trace metal was removed from all
buffers by passage through a Chelex 100 (Bio-Rad) column.
For each titration point, 22 μM protein was mixed with 28 μM of Zn and variable amounts
of NTA. Samples were allowed to equilibrate overnight at room temperature and
subsequently centrifuged in 0.5 mL Amicon Ultra (Millipore) protein concentrators (10 KDa
MW cutoff) for ten minutes at 2000 rpm to allow free Zn, free NTA and NTA:Zn to pass
through the membrane. Zn concentrations were determined for the flow-through and protein
chamber at each titration point using 4-(2-pyridylazo)resorcinol (PAR), which exhibits an
increase in aborbance at 500 nm upon binding Zn (Δε500 = 59100 M−1cm−1 in buffer and
54800 M−1cm−1 in 5 M guanidine hydrochloride), essentially as previously described.11
1800 μL of 1 mM PAR was added to a 1-cm pathlength cuvette, and the initial absorbance
spectrum was recorded. Subsequently, 100 ml of flow-through was added, allowed to stir for
a minimum of five minutes to ensure that equilibrium had been reached, and absorbance at
500 nm was again recorded. Metal content in the protein chamber was determined similarly
to flow-through samples, except that 1800 μl of 5 M buffered guanidine hydrochloride was
used to denature protein and allow the complete release of bound metal for measurement.
Absorbance at 500 nm was corrected for dilution and protein absorbance, and Zn
concentrations determined using the Δε500 values listed above.
The fraction of protein bound to metal as a function of competing ligand concentration was
determined by the following equation:
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where [M]P.C. is the concentration of metal in the protein chamber, [M]F.T. is the
concentration of metal in the flow-through and [P]T is the total concentration of protein
measured after centrifugation. For calculating dissociation constants, the concentration of
free metal was determined based on the concentration and dissociation constant of the
competing ligand (NTA) and metal in the flow-through using the program MaxChelator
(http://maxchelator.stanford.edu). The data were fit to the following models38 using Igor
Pro v. 6.02a (Wavemetrics, Inc.).
(1)
(2)
(3)
where B is the molar equivalents of metal bound per C96RIDC-14, n is the number of binding
sites, [M] is the concentration of free metal and K1–4 are stoichiometric association
constants. Equation (1) assumes four identical binding sites with invariant affinities.
Equation (2) assumes two stepwise binding events each of which involves binding of two
Zn2+. Equation (3) assumes four stepwise binding events.
Binding affinity of Fura-2 for Ni2+, Cu2+ and Zn2+
1 mg of lyophilized Fura-2 (Invitrogen) was suspended in 1 mL of deionized water and its
concentration was determined based on a published extinction coefficient of 27,000
M−1cm−1 at 362 nm.39 EGTA competition binding assays were used to determine
dissociation constants for Ni2+, Cu2+ and Zn2+ using the following EGTA:M2+ logK values
obtained from the online program MaxChelator (http://maxchelator.stanford.edu):
Cu2+:EGTA − 13.2, Ni2+ − 9.0, Zn2+ − 8.1. These values are corrected for pH, temperature
and ionic strength. In a 3 mL quartz cuvette, 11 μM Fura-2 was mixed with 100 μM EGTA
in 20 mM MOPS (pH 7) and 150 mM NaCl to a final volume of 2 mL at 22° C. M2+ was
titrated into the solution with an equilibration time of 10 minutes at ambient conditions
between each addition, and absorbance spectra were recorded from 190–800 nm using a
Hewlett Packard 8452A diode array spectrophotometer (Figure S10). For Cu2+, a separate
titration into a solution of 100 μM buffered EGTA was conducted and subtracted from the
Fura-2 titration points to account for absorbance from aqueous Cu2+ and the Cu2+:EGTA
complex. Absorbance and M2+ concentrations were corrected for dilution and data was fit to
a 1:1 M2+:Fura-2 model using DynaFit40 scripts (Figure S10) modeling the competition
between EGTA and Fura-2 for M2+. The Zn2+:Fura-2 value determined by this method
(logK = 8.2) compares favorably with a value previously determined at pH 7.15 (logK =
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8.5).41 The logK values determined for the Fura-2 complexes of Cu2+ and Ni2+ complexes
are 12.5 and 8.2, respectively.
Competitive binding assays using Fura-2
Competition assays using the Fura-2:M2+ dissociation constants determined above were
used to measure C96RIDC-14:M2+ affinities. In a 3-mL quartz cuvette, a stock solution
of C96RIDC-14 was diluted to 7.5 μM in 20 mM MOPS (pH 7) and 150 mM NaCl at 22° C
and absorbance was recorded. Fura-2 was added to a final concentration of 10 μM and
absorbance again recorded. Stock metal solutions were then titrated into the
Fura-2/C96RIDC-14 mixture in ~2 μM steps, allowed to equilibrate with stirring for 10
minutes at ambient conditions and absorbance recorded. Protein and metal concentrations
were corrected for dilution, protein absorbance was subtracted from total absorbance and
plots of Abs373 versus total metal added were fit to various models using custom DynaFit
scripts (see Supporting Information). Co titrations were performed similarly to Zn, Ni and
Cu, but with the following modifications due to its lower affinity
for C96RIDC-14: C96RIDC-14 concentration was increased to 21 μM and the Fura-2
concentration was decreased to 8 μM. Also, titrations were performed in a 0.5 cm pathlength
cuvette to allow for the use of a higher protein concentration without saturating the detector.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We thank Prof. Thomas O’Halloran for helpful discussions. Support for this work was provided by NIH (training
grants to J.D.B. and A.M.M.; predoctoral fellowship to E.N.S.), NSF (CHE-0908115, structural work), DOE (DE-
FG02-10ER46677, competitive metal binding titrations) and the Arnold and Mabel Beckman Foundation (F.A.T.).
Portions of this research were carried out at SSRL, operated by Stanford University on behalf of DOE.
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Figure 1.
Interprotomeric interfaces in the D2-symmetric Zn4:MBPC-14 complex. Residues subjected
to redesign as well as those that coordinate Zn2+ ions are shown as sticks.
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Figure 2.
(a) Sedimentation coefficient distributions for various cyt cb562 constructs at 5 μM
monomeric concentration and equimolar Zn2+ where indicated. For MBPC-1 and RIDC-1,
the Zn-induced tetrameric species are fully populated at >1 mM and >20 μM protein and Zn,
respectively. At 5 μM protein and Zn, MBPC-1 is still predominantly monomeric (Smax=
1.8), while RIDC-1 is a mixture of dimeric (Smax= 2.8) and tetrameric (Smax= 4.5) forms.
(b) Sedimentation equilibrium profile for 2.5 μM C96RIDC-12 obtained at 20000 rpm (see
Figure S2 for other concentrations and speeds). The sedimentation data are equally well
described by a dimer-tetramer equilibrium (Kd = 52 nM) or a tetramer-only model, which
suggests the tetramer dissociation constant to be <100 nM.
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Figure 3.
Crystal structures of C96RIDC-14 (left) and Zn4:C96RIDC-14 (right). Redesigned residues in
i1 and i2 are shown as sticks. Interfacial SS-bond configurations are shown below each
structure along with corresponding Fo – Fc omit difference maps (cyan mesh − 4.5 σ (apo),
3 σ (Zn); purple mesh − 11 σ (apo), 5 σ (Zn)). See Figure S4 for detailed views of interfacial
residues.
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Figure 4.
(a) Zn-binding isotherm of C96RIDC-14 determined using NTA as a competing ligand. (b)
Zn-binding isotherm for Fura-2–C96RIDC-14 competition experiments; corresponding
changes in the Fura-2 absorbance spectrum are shown in the inset. The data are corrected for
dilution and background absorbance by the protein. The sample contained 7.5
μM C96RIDC-14 and 11 μM Fura-2. The tick marks shown on the top x-axis correspond to
theoretical endpoints for titration if C96RIDC-14 bound to one, two, three or four equivalents
of Zn. The fits obtained using DynaFit are shown for the following different models: solid
line, four consecutive Zn binding equilibria (1+1+1+1); dashed line, two consecutive
binding equilibria (2+2); dotted line; single binding equilibrium (4×1). Equilibrium
constants obtained with these different models are listed in Table 1.
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Figure 5.
Extent of divalent metal ion binding to C96RIDC-14 in competition experiments.
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Figure 6.
Anomalous difference maps (4σ) for the C96RIDC-14 structures grown in the presence of
equimolar Cu2+ and Zn2+ obtained at the Zn (a) or Cu (b) K edges. As expected, the heme
Fe centers show anomalous signals at both wavelengths, whereas the core metal sites do the
same only at the higher energy Zn edge, unambiguously identifying them as Zn ions. (c) X-
ray fluorescence excitation scans of the same crystal – which was thoroughly washed with
non-metal containing solutions – indicate the presence of both Zn and Cu.
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Figure 7.
(a) Cu2+ and (b) Ni2+ binding isotherms for Fura-2–C96RIDC-14 competition experiments;
corresponding changes in the Fura-2 absorbance spectrum are shown in the inset. The
samples contained 7.5 μM C96RIDC-14 and 11 μM Fura-2. The tick marks shown on the top
x-axis correspond to theoretical endpoints for titration if C96RIDC-14 bound to one, two,
three or four equivalents of metal. The fits obtained using DynaFit are shown for the
following different models: solid line, two consecutive binding equilibria (1+1); dashed line,
single binding equilibrium (2). Equilibrium constants obtained with these different models
are listed in Table 1.
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Scheme 1.
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Table 1
Association constants for various metal binding equilibrium models for C96RIDC-14 determined through competitive Fura-2 titrations (pH 7, 295 K). The
total free energies for metal binding correspond to the free energy sums of individual equilibria (times their multiplicity) for every model. Corresponding
titrations and fits are shown in Figure 4b (Zn2+), Figure 7 (Cu2+ and Ni2+) and Figure S11 (Co2+). Numbers in parentheses correspond to standard
deviation in the last reported significant figure, were obtained through DynaFit, and do not include any experimental errors.
Total metal equivalents
Number of Consecutive Binding Equilibria
Kd1 (M)
Kd2 (M)
Kd3 (M)
Kd4 (M)
Total -ΔG for metal binding (kJ mol−1)
4 Zn2+
2
5.2(4) × 10−10
4.3(2) × 10−8
189
4
1.3(3) × 10−9
5.3(7) × 10−10
3.3(8) × 10−8
5.8(8) × 10−8
186
2 Cu2+
1
1.0(1) × 10−12
136
2
2.5(3) × 10−13
1.4(1) × 10−12
138
2 Ni2+
1
8.0(9) × 10−9
92
2
9.0(1) × 10−10
4.9(5) × 10−9
93
1 Co2+
1
9(4) × 10−7
34
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|
3M7D
|
Crystal structure of an N-terminal 44 kDA fragment of topoisomerase V in the presence of dioxane
|
Structures of minimal catalytic fragments of topoisomerase V
reveals conformational changes relevant for DNA binding
Rakhi Rajan*, Bhupesh Taneja*,†, and Alfonso Mondragón*,‡
* Department of Biochemistry, Molecular Biology and Cell Biology, Northwestern University, 2205
Tech Dr, Evanston, IL 60208
Summary
Topoisomerase V is an archaeal type I topoisomerase that is unique among topoisomerases due to
presence of both topoisomerase and DNA repair activities in the same protein. It is organized as an
N-terminal topoisomerase domain followed by 24 tandem helix hairpin helix (HhH) motifs.
Structural studies have shown that the active site is buried by the (HhH) motifs. Here we show that
the N-terminal domain can relax DNA in the absence of any HhH motifs and that the HhH motifs
are required for stable protein-DNA complex formation. Crystal structures of various
topoisomerase V fragments show changes in the relative orientation of the domains mediated by a
long bent linker helix, and these movements are essential for the DNA to enter the active site.
Phosphate ions bound to the protein near the active site helped model DNA in the topoisomerase
domain and shows how topoisomerase V may interact with DNA.
Introduction
DNA topoisomerases are enzymes found in all forms of life (bacteria, eukarya, and archaea)
and they regulate the topological state of DNA inside the cell. They form a transient break in
a single or double stranded DNA and allow the passage of another single or double DNA
strand through the break, before resealing the break (Champoux, 2001) (Schoeffler and
Berger, 2008). As a result of this, topoisomerases can relax supercoiled DNA, help in the
segregation of DNA strands following replication, and lead to the formation and resolution
of knots and catenates (Gellert, 1981). Topoisomerases participate in many aspects of DNA
metabolism, such as replication, recombination, and transcription (Champoux, 2001). In
addition, they are targets of various anti-cancerous drugs and anti-bacterial agents (Maxwell,
1999; Pommier, 1998; Rothenberg, 1997; Wang et al., 1997).
DNA topoisomerases are broadly classified into two types, type I and type II enzymes. Type
I enzymes cleave a single strand of a DNA molecule and pass another single or double
stranded DNA through the break before resealing the opening. Type II enzymes cleave both
‡Corresponding author: Phone: 847-491-7726, Fax: 847-467-6489, a-mondragon@northwestern.edu.
†Present address: Institute of Genomics and Integrative Biology, CSIR, Delhi, India
Protein data bank accession codes
The final structure factors and coordinates of Topo-31, Topo-44 Form I, Form II, and Form III have been deposited in the Protein Data
Bank with accession codes 3M7G, 3M7D, 3M6K, and 3M6Z respectively.
Supplementary data
Supplementary data are available at Structure Journal Online.
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Author Manuscript
Structure. Author manuscript; available in PMC 2011 July 14.
Published in final edited form as:
Structure. 2010 July 14; 18(7): 829–838. doi:10.1016/j.str.2010.03.006.
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strands of a double stranded DNA in concert and pass another double stranded DNA through
the break. Type I enzymes use the torsional energy stored in the supercoiled DNA to drive
DNA relaxation and hence they do not require high energy cofactors, such as ATP, for their
activity (Baker et al., 2009) Type II enzymes, on the other hand, require ATP and Mg2+ for
their activity. Type I topoisomerases are further subdivided into three subtypes: IA, IB, and
IC (Forterre et al., 2007). Type IA and IB enzymes have been studied extensively (Baker et
al., 2009) and there is ample information available about their general mechanism of DNA
relaxation and the mode of DNA binding. Type IC, on the other hand, is a relatively new
subtype. Currently topoisomerase V is the only member of this family and it has been
identified only in the Methanopyrus genus. Previously, topoisomerase V had been
considered as a type IB enzyme based on its biochemical characteristics (Slesarev et al.,
1993), but the crystal structure of an N-terminal 61 kDa of topoisomerase V (Topo-61)
(Taneja et al., 2006) revealed a completely new fold without similarity to other
topoisomerases or any other known protein. Furthermore, the orientation of the putative
active site residues is also different from other type I topoisomerases, suggesting a different
mechanism of cleavage and religation of DNA. These observations, together with the lack of
sequence similarity, indicated that topoisomerase V defines a new subtype of type I enzymes
(Forterre, 2006).
Topoisomerase V was identified in Methanopyrus kandleri, an extremophile isolated from a
deep-water ‘black smoker’ chimney in the Gulf of California (Huber et al., 1989). The
enzyme is active at very high temperatures (122°C) and high salt concentrations (0.65 M
NaCl and 3.1 M potassium glutamate). The unusual characteristic of topoisomerase V is that
it has both topoisomerase and DNA repair activities in the same polypeptide (Belova et al.,
2001). Based on the sequence analysis of topoisomerase V, it has been predicted that the
protein contains 24 helix-hairpin-helix (HhH) DNA binding motifs arranged as 12 (HhH)2
domains around the N-terminal topoisomerase domain (Belova et al., 2002) (Figure 1A).
Some of these (HhH)2 domains are involved in the apurinic/apyrimidinic (AP) site-
processing activity, but the exact location of the repair active site is not known yet.
Topoisomerase V can relax both positively and negatively supercoiled DNA without the
need for metal cations or high energy cofactors. Single molecule experiments have shown
that topoisomerase V relaxes DNA by a constrained swiveling mechanism, relaxing around
12 turns of DNA per relaxation cycle (Taneja et al., 2007). Type IB enzymes, which also use
a constrained swiveling mechanism for DNA relaxation, relax around 19 turns of DNA per
relaxation cycle (Koster et al., 2005).
The structure of Topo-61 showed that the topoisomerase domain is mainly alpha helical and
that the first four (HhH)2 domains curl around the topoisomerase domain (Taneja et al.,
2006) (Figure 1B). The topoisomerase and (HhH)2 domains are joined by a long bent helix,
termed the “linker helix”. Three of the five putative active site residues are present in a
helix-turn-helix (HTH) domain and the other two are present in an intervening loop and a
helix. The active site residues are buried by the first (HhH)2 domain and it has been
suggested that large conformational changes will be needed for the DNA to access the active
site of topoisomerase V (Taneja et al., 2007). Here we present data that shows that the N-
terminal 31 kDa fragment of topoisomerase V (Topo-31) has topoisomerase activity,
consistent with previous predictions based on the structure. In addition, we show that the
Topo-44 fragment (N-terminal 44 kDa fragment of topoisomerase V) can form a stable
protein-DNA complex, emphasizing the need of the (HhH)2 domains for binding DNA. We
determined a crystal structure of (Topo-31) fragment, which has only the topoisomerase
domain, and three different crystal structures of the Topo-44 fragment, which includes the
topoisomerase domain and three tandem HhH motifs. In all structures, the topoisomerase
domain is very similar. In contrast, the structures of Topo-44 show conformational changes
in the linker helix resulting in variable orientations of the (HhH)2 domains when compared
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to the Topo-61 structure. Phosphate ions are present in the vicinity of the topoisomerase
active site in two of the Topo-44 structures. Some of the catalytic residues interact with the
phosphate ions and may mimic contacts with DNA. These observations suggest that the
movement of the (HhH)2 domains is mediated by the linker helix and helps expose the
topoisomerase active site to facilitate DNA binding. In addition, the location of the
phosphate ions suggests a possible path for the DNA and the way the active site residues
interact with it.
Results
The topoisomerase domain can relax DNA
DNA relaxation assays using different topoisomerase V fragments showed that the
topoisomerase domain alone is capable of relaxing DNA. Topoisomerase V fragments with
different numbers of (HhH)2 domains, Topo-31, Topo-44, and Topo-78, were studied using
relaxation assays. Topo-31 has no (HhH)2 domains, Topo-44 has one full and one partial
(HhH)2 domain, while Topo-78 has eight full (HhH)2 domains, including a putative DNA
repair domain. In addition to standard conditions, the effect of different pH conditions and
presence of magnesium ions were also tested. The experiments show that Topo-31 is
capable of relaxing DNA, despite the absence of the (HhH)2 domains (Figure 2B). A pH
profile analysis for the DNA relaxation assays showed that Topo-78 relaxes DNA over a
wider pH range (pH 5 to 9), while Topo-31 and Topo-44 relax DNA optimally at pH 5
(Figure 2A, 2B, 2C). In addition, magnesium is not required for the reaction, but stimulates
it at all pH values (Figure 2B, 2C). Topo-78 can relax DNA to the same extent with lower
amounts of protein (0.1 μg/reaction) compared to Topo-44 (~1.5 μg/reaction) and Topo-31
(~9 μg/reaction). This could be due to the enhanced DNA binding facilitated by the (HhH)2
domains. Together, these results suggest that, even though the (HhH)2 domains are
dispensable for topoisomerase activity, they enhance DNA relaxation activity. In addition,
the pH dependence of the DNA relaxation activity indicates that the reaction is likely to
involve side chains with ionizable groups in the low pH range, such as glutamates. Finally,
the magnesium independence of the reactions confirms that even the smallest fragments do
not require metals for activity, although magnesium has a stimulatory effect. This may be
due to favorable interactions of the cations with DNA.
The (HhH)2 domains enhance DNA binding affinity
EMSA experiments with different fragments of topoisomerase V and DNA showed that
(HhH)2 domains could help in the formation of a stable protein-DNA complex. Various
topoisomerase V fragments (Topo-31, Topo-44, and Topo-78) and single and double
stranded DNA were analyzed by EMSA experiments. Topo-44 and Topo-78 formed stable
complexes with a 39mer double stranded DNA (Figure 2D), while no DNA binding was
observed for the Topo-31 fragment (data not shown). These observations indicate that
(HhH)2 domains are necessary for a stable protein-DNA complex and that as few as one and
half (HhH)2 domains are enough for formation of a stable protein-DNA complex. EMSA
with single stranded DNA showed that Topo-31 and Topo-44 cannot bind to single stranded
DNA, while Topo-78 can bind to single stranded DNA (data not shown).
Overall Structures
The topoisomerase domain of topoisomerase V is a helical-rich compact domain that has no
structural similarity to any other known protein. The only recognizable structural element is
a HTH that contains some of the active site residues. Not surprisingly, the topoisomerase
domain of the four structures (Topo-31, Topo-44 (Forms I, II, and III)) superimpose very
well on each other and also to that from the Topo-61 structure. In the Topo-31 structure, two
surface loops, residues 39-49 and 120-124, adopt a different conformation compared to the
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Topo-61 and Topo-44 structures. These two loops are not always visible in the Topo-44
structures, suggesting that they are mobile regions. The r.m.s.d. for the superposition of the
topoisomerase core domain of all the new structures on to the Topo-61 structure range from
0.2 Å to 0.7 Å if the two mobile surface loops are not included (Figure 3A). In general, the
topoisomerase domain remains unchanged and is identical in all structures. The (HhH)2
domains also remain largely unchanged, with r.m.s.d. for the superposition of only the
(HhH)2 domains from the three Topo-44 crystal forms and equivalent domains in the
Topo-61 structure ranging from 0.31 Å to 0.56 Å.
The five crystallographically independent structures of Topo-44 (Form I, Form II A and B
monomers, and Form III A and B monomers) were compared with each other and to the two
crystallographically independent Topo-61 monomers to understand the conformational
changes in the protein. The r.m.s.d. for the superposition of all the Topo-44 structures
(residues 3-375) on to the Topo-61 fragment or on each other vary between 0.9 Å and 2.7 Å,
with the majority above 1.5 Å, showing that in general the structures have slightly different
conformations. As mentioned above, the different domains behave as rigid or almost rigid
subunits and the only change in the structure is the relative orientation between the
topoisomerase and the (HhH)2 domains. The change in orientation of the domains starts at
the linker helix (residues 269-295), which acts as a hinge region, and follows into the
(HhH)2 domains. At the start of the linker helix, the structures superimpose very well for all
five Topo-44 and two Topo-61 structures. In the middle of the linker helix there is a kink
after which the linker helix from all the structures shows different orientations (Figure 3B).
The flexibility of the linker helix is also evident by the fact that the linker helix in the B
subunit of Form III crystals appears in two alternate conformations. The change in the
relative orientation of the (HhH)2 and topoisomerase domains (Figure 3C and 3D), suggests
that these domains can adopt different orientations and these movements might be necessary
for the DNA to access the active site.
The topoisomerase domain has a positively charged groove adjacent to the active site
The structure of the Topo-31 as well as the structures of the Topo-44 fragment reveals the
presence of a positively charged groove in the protein that encompasses the active site
region (shown later in Figure 6C). This charged groove had been observed before in the
structure of the Topo-61 fragment, although several (HhH)2 motifs partially obstruct it
(Taneja et al., 2006). The structure of the Topo-31 confirms the presence of the groove even
in the absence of the (HhH)2 motifs. The groove is long and can be deep in some areas. It
includes regions of the HTH motifs and extends all the way to the linker helix. All the
residues forming the active site pentad point towards the groove. The active site tyrosine,
Tyr226, is found near one of the ends of the groove, a region where it widens. The positively
charged character of the groove and its presence by the active site strongly suggest that it
may be involved in DNA binding.
Phosphate ions bind in the groove near the topoisomerase active site
An interesting observation stemming from the Form II and Form III Topo-44 structures is
the presence of phosphate ions near the positively charged DNA binding groove. All three
Topo-44 crystal forms were crystallized in the presence of phosphate-citrate buffer, but only
Form II and Form III structures showed phosphate ions bound to the protein, which were
assigned based on electron density consistent with a tetrahedral phosphate ion (Figure 4A).
Form II and Form III crystals include 1–1.2 M guanidium hydrochloride in the
crystallization solution. The high resolution Form III structure shows clear density for three
guanidium ions bound to the protein, two very well ordered and one with weak density. The
presence of guanidium hydrochloride in the crystals appears to trigger a conformational
change allowing the binding of phosphate ions to the protein. It is interesting to note that
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Form I crystals did not show any bound phosphate albeit its presence in the crystallization
condition. This could be due to the absence of guanidium hydrochloride to trigger the
binding of phosphate ions as observed in Form II and Form III structures. There are three
phosphate ions in the B subunit and none in the A subunit of the Form II Topo-44 structure.
Two of the phosphates are in the topoisomerase active site and one of them forms close
contacts with the putative active site residues in the topoisomerase domain (Figure 4B).
Form III crystal has seven phosphate ions, three in each subunit and one between both the
subunits. In the Form III structure, the phosphate ion near the active site Tyr226 is absent,
but it shows several new locations for phosphate ions, especially in the positively charged
groove containing the topoisomerase active site (Figure 5A). An overlay of the A and B
subunits of the Topo-44 Form III structure with the B subunit of Topo-44 Form II structure
shows eight unique phosphate ions (Figure 5A). It clearly shows that there are more
phosphate ions bound in the positively charged groove compared to other regions of the
protein.
Taking into account all structures, there are five unique phosphate ion binding sites in the
putative DNA binding groove and an additional one near its end and close to the start of the
linker helix. Several pairs of phosphates in the groove are separated by a distance of around
7 Å (Figure 5B), which would be consistent with the phosphate-phosphate distance in
adjacent nucleotides in a DNA double helix. One of the phosphates (P1) is found near the
active site tyrosine and is coordinated by Tyr226 and Arg131, two residues that have been
implicated in cleavage and religation of the DNA (Taneja et al., 2006), and by Glu215,
whose charge may be important for interactions with DNA (R.R. and A.M., unpublished
observations). The side chains of the tyrosine and the glutamate residues are in contact with
Arg144 and His200, the other putative active site residues, and these interactions may help
to orient them for the catalytic reaction. Adjacent to P1, there is a second phosphate (P2) at a
distance of 7.5 Å which is trapped between the topoisomerase domain and an HhH motif. P2
is coordinated by Arg131, an active site residue, in addition to Arg108 from the
topoisomerase domain and Arg293 and serines 322 and 324 from the second HhH motif
(Figure 6C). Three more phosphates are found in the groove (P3, P4, and P5) coordinated
mainly by positively charged residues, such as Arg37, Lys47, Arg108, Lys134, and Arg135
from the topoisomerase domain and also residues from the linker helix such as Tyr289 and
Arg293. In general, some of the side chains can contact more than one phosphate. The
distance between P3 and P4 and P4 and P5 is 6.8 Å and 6.5 Å respectively. A final
phosphate (P6) is located at the start of the linker helix and on the edge of the groove
(Figure 5A).
Discussion
Topoisomerase V is active at very high temperatures (122°C) and high salt concentrations.
DNA relaxation assays with various topoisomerase V fragments (Topo-44 and Topo-61)
show that a temperature above 60° C is required for optimal activity, although longer
fragments of topoisomerase V can relax DNA at lower temperatures (Taneja et al., 2007).
Topo-44 was first identified by limited proteolytic digestion of the full length topoisomerase
V protein (Belova et al., 2002) at 80°C. In contrast, Topo-61 is the shortest fragment
showing topoisomerase activity when the proteolytic reaction is performed at 37°C (Belova
et al., 2002). The N-terminal Topo-31 fragment, which contains neither HhH motifs nor the
linker helix, was identified as the smallest region spanning the topoisomerase domain from
the crystal structure of Topo-61 fragment (Taneja et al., 2006) and it was suggested that it
could represent the minimal domain capable of relaxing DNA. Relaxation experiments with
this minimal domain show that this is indeed the case, although the activity is not as robust
as with longer fragments. As expected, Topo-31 does not require magnesium for activity,
but magnesium enhances it, as is the case for type IB topoisomerases, which also uses a
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swiveling mechanism for DNA relaxation (Stewart et al., 1996). The optimal pH for activity
for the Topo-31 and the Topo-44 fragments is around 5. This pH dependence is not observed
for the Topo-78 fragment. The DNA relaxation by shorter fragments of topoisomerase V at
pH 5 could point to the involvement of some ionizable side chains in the relaxation activity.
It could also be simply due to the effects of various side chains on DNA binding. Further
experiments with different active site mutations in both longer and shorter fragments of
topoisomerase V will be required to probe the pH dependence of the relaxation reaction by
shorter topoisomerase V fragments.
Gel shift experiments show that Topo-44 and also longer fragments (Topo-78) can bind
double stranded DNA. Surprisingly, Topo-31 does not show DNA binding activity in these
assays even though it is still capable of relaxing DNA. It appears that the presence of the
(HhH)2 domains stabilizes the DNA/protein complex. One possibility is that the (HhH)2
domains could play a similar role to the cap domain present in type IB enzymes, which helps
to encircle the DNA during the swiveling reaction (Redinbo et al., 1998). In addition, both
short fragments of topoisomerase V do not bind single stranded DNA, whereas Topo-78 can
form a stable complex with single stranded DNA (data not shown). (HhH)2 domains binding
to single stranded DNA has been observed before. For instance, the N-terminal 8 kDa of
mammalian polymerase β, which contains a single HhH motif, binds to single stranded DNA
through both helices (Kumar et al., 1990; Liu et al., 1994). The exact mode of single
stranded DNA binding by Topo-78 or the possible role in relaxation or repair activities is not
yet clear.
The structure of Topo-61 showed that the topoisomerase active site of topoisomerase V is
buried by one of the (HhH)2 domains suggesting that conformational changes are essential
for the protein to bind DNA. The present structures of Topo-44 reinforce this observation
and show that the (HhH)2 domains can change their position relative to the topoisomerase
domain and that this change is mediated by the movement of the linker helix. The (HhH)2
domains act as rigid individual units, as evidenced by the fact that in different structures
they show the same structure and relative orientation of the two HhH motifs. The
topoisomerase domain also appears to be rigid showing the same structure even in the total
absence of the rest of the protein. The linker helix (residues 269-295), which is a long bent
helix, serves as a hinge for the movement of the (HhH)2 domains away from the rigid
topoisomerase domain, possibly by responding to interactions with double stranded DNA.
This movement has to be quite large. The Topo-44 structures in the absence of DNA capture
the regions that move, but do not show the full extent of the movement or indicate the way
the HhH motifs interact with DNA.
As mentioned before, topoisomerase V binds double stranded DNA and has a groove wide
enough to accommodate double strand DNA (Figure 6C). The presence of an HTH domain
normally associated with DNA binding, the positively charged nature of it, and several
phosphates bound along it suggest that this groove could be involved in DNA binding. In
addition, the active site is found in this groove and some residues form part of the HTH
domain. Previously, DNA was modeled bound to the topoisomerase domain (Taneja et al.,
2006) based on the structures of HTH domains in complex with DNA but there was no
evidence to support it. Using the phosphates present in the groove in the current structures, it
is possible to refine this model. A superposition of the B subunit of Form II and the A and B
subunits of Form III Topo-44 structures shows five different phosphate ions in the positively
charged groove which are separated by a distance of around 7 Å, consistent with the
distance of consecutive phosphates in B DNA of ~6.4 Å. A sixth phosphate ion is found
outside the groove near the linker helix. A double stranded DNA molecule was modeled into
the groove based on the positions of the phosphate ions (Figure 6). Only five out of the six
phosphates could be placed on the DNA molecule, as one of them was inconsistent with a
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double stranded molecule. Phosphate ions P3, P4, and P5 would correspond to three
adjacent phosphates in one DNA strand, while P1, located near the active site, would belong
to the opposite strand. A final phosphate (P6) is away from the groove and near the linker
helix (Figure 6A, 6B). The fit to the four inner phosphates is excellent and the DNA can be
accommodated in the groove of the Topo-31 structure without the need for any major
rearrangements of the protein backbone. The fifth phosphate (P6) does not fit as well and a
better fit would require movement of either the protein or the DNA, but the change would be
relatively modest. Several side chains would need to move, but these changes would also be
minor. The major change needed to accommodate the DNA in the structures with the
(HhH)2 domains present is the movement of the (HhH)2 domains away from the
topoisomerase domain (Figure 6B). The movement of (HhH)2 domains should be feasible as
is evident from the Topo-44 structures showing different orientations of the (HhH)2
domains. The location of the (HhH)2 domains after DNA binding is not evident, but one
possibility is that they would help enclose the DNA to form a clamp around it, similar to the
arrangement in type IB enzymes.
In the model of the topoisomerase domain in complex with DNA, the active site residues are
in close contact with the backbone of DNA. The catalytic Tyr226 is pointing towards the
phosphate of the DNA backbone; Arg131 and Arg144 are positioned to stabilize the protein-
DNA covalent complex. Surprisingly Glu215 also appears to interact directly with the DNA
phosphate backbone. The other active site residues like His200 and Lys 218 are also near the
DNA. The active site is located near the end of the groove, where it widens. At this end, the
DNA fits loosely in the groove, which is spacious to accommodate the movement of the
strands. The ‘constrained swiveling’ mechanism employed by type IB and IC enzymes
necessitates rotation of one strand about the other after forming the covalent protein-DNA
intermediate. The position of the active site at the wider end of the putative DNA binding
groove would facilitate the rotation of the DNA strand at this end, while holding the rest of
the DNA in place through extensive interactions along the groove.
Even though type IB and IC enzymes have a similar overall mechanism of action, the
structures of fragments of topoisomerase V suggest many differences. Type IB enzymes
have two domains which come together to form a C-shaped clamp around the DNA (Perry et
al., 2006; Redinbo et al., 1998; Stewart et al., 1998) The protein has an open stage where
these domains are separate and this helps in the entry and release of the DNA from the
protein active site. A wide DNA binding cavity is not observed in the topoisomerase V
structures. Instead, the structures show a positively charged groove which is always present
in the protein and does not require domain rearrangements to form. DNA can access this
groove after a conformational change involving the movement of the (HhH)2 domains
exposing the active site. The (HhH)2 domains could help enclose DNA during the swiveling
of the DNA, forming a similar enclosure to the one observed for type IB enzymes. It is not
known whether all HhH motifs contact DNA simultaneously, but this appears unlikely
without a major rearrangement of the motifs. It is likely that only some of the HhH motifs
contact DNA at any given time or that some of the motifs do not have the capacity to bind
DNA. Finally, similar to type IB enzymes (Cheng and Shuman, 1998), the putative domain
enclosing the DNA is dispensable for activity, although it enhances the relaxation activity
markedly. Thus, it is likely that type IB and IC enzymes have several overall similarities in
the way that they interact with DNA, but the atomic details are markedly different.
There are still many details of the atomic mechanism of type IC topoisomerases that need to
be understood. The present functional and structural studies provide new information about
topoisomerase V including the observations that the Topo-31 is the minimal fragment
capable of DNA relaxation, the (HhH)2 domains enhance binding of the protein to DNA, the
changes in relative orientation of the domains is mediated by the linker helix, and several
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phosphate ions bind in a positively charged groove. Furthermore, the position of the
phosphate ions in the groove helped in the placement of DNA in the topoisomerase domain
and this provides an initial model of how topoisomerase V interacts with DNA. Thus the
present study helps to establish the role of different domains more clearly, to illustrate a
mechanism to drive the conformational changes needed for activity, and to suggest a
possible manner of binding DNA. Additional work on structures of protein/DNA complexes
and intermediates in the swiveling reaction are needed to understand the way this new type
of topoisomerases interacts with DNA to perform a complex reaction.
Experimental Procedures
Protein purification
The N-terminal 31 kDa (Topo-31: residues 1-269), and 44 kDa (Topo-44: residues 1 to 380)
fragments of topoisomerase V protein were cloned into the pET15b plasmid and
transformed into Escherichia coli BL21 Rosetta (DE3) cells. The N-terminal 78 kDa
(Topo-78: residues 1 to 685) fragment of topoisomerase V protein was cloned into the
pET14b plasmid (Belova et al., 2002) and transformed into Escherichia coli BL21(DE3)
cells. For protein production, cells were grown at 37° C in LB medium containing 100 μg/ml
ampicillin and 100 μg/ml chloramphenicol for Rosetta cells and LB medium with 100 μg/ml
ampicillin for BL21(DE3) cells to an optical density (OD600) of 0.6. The cells were then
cooled down on ice, followed by the addition of isopropyl β-D-1-thiogalactopyranoside
(IPTG) to a final 0.5 mM concentration, and grown overnight at 16° C. Cells were harvested
and resuspended in 50 mM Tris pH 8, 500 mM NaCl, 0.5 mM EDTA, 1 mM DTT, flash
frozen in liquid nitrogen and stored at −80° C. After thawing the pellet, pepstatin (1μg/ml),
benzamidine (1mM), PMSF (1mM), and Brij 58 (0.1%) were added to the cells and the
protein was purified as described earlier (Taneja et al., 2006) The protein was further
purified by anion exchange and gel filtration chromatography. Pure protein was
concentrated and stored in 50 mM Tris pH 8, 250 mM NaCl, and 1 mM DTT. The seleno-
methionine substituted Topo-44 was prepared from cells grown in a minimal medium
supplemented with nutrients and salts (Doublie, 1997); protein purification followed the
same procedure as for the native protein except that 5mM DTT was used in all the
purification steps and for storage.
Relaxation assays
Relaxation assays with the different topoisomerase V fragments were carried out at pH
values ranging from 4 to 10. The pH of the buffers was adjusted at 65 °C to account for the
change in pH at higher temperature. The different buffers used were: sodium acetate for pH
4 and 5, MES for pH 6, HEPES for pH 7, TRIS for pH 8, CHES for pH 9, and CAPS for pH
10. Topoisomerase activity assays were performed by incubating varying amounts of protein
(Topo-31, Topo-44 or Topo-78) with 0.2 μg negatively supercoiled pUC19 DNA in 50 mM
of the required buffer, 30 mM NaCl, 0.2 mM or 5 mM EDTA or 1 mM MgCl2. The
reactions were carried out at 65 °C for 15 min and terminated by cooling and addition of
SDS to a final 1% concentration. The products were resolved on a 1% agarose gel and
visualized by ethidium bromide staining.
Electrophoretic Mobility Shift Assay
For Electrophoretic Mobility Shift Assay (EMSA), 4 μM of a 39mer double stranded DNA
oligonucleotide (5′ GCGACGCGAGGCTGGATGGCCTTCCCCATTATGATTCTT3′) was
incubated with different concentrations of topoisomerase V fragments in 50 mM sodium
acetate pH 5, 30 mM NaCl, 1 mM MgCl2 at 65 °C for 30 minutes. Glycerol was added to
the reaction mixture to a final concentration of 8% and the products were separated on a 4 %
acrylamide native gel. The gel was stained with ethidium bromide to detect the DNA. When
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a stable protein-DNA complex was formed, there was an upward shift in the band indicating
a higher molecular weight complex.
Crystallization
Topo-31 crystals were grown using the sitting drop vapor diffusion method equilibrated
against, 23% PEG 6000, 0.1 M Na citrate pH 5.5, at 22°C. For data collection, the Topo-31
crystals were cryo-protected by adding glycerol to the mother liquor to a final 20%
concentration. Topo-44 was crystallized by the hanging drop vapor diffusion method under
three different crystallization conditions (Forms I, II, and III). Crystal Form I grew under 0.1
M phosphate citrate pH 5, 0.2 M NaCl, 15% PEG 3350 and 8% dioxane. The crystals were
cryo-protected by increasing the PEG concentration to 30%. Form II crystals grew under 0.1
M phosphate citrate pH 5, 0.2 M NaCl, 16% PEG 8000 and 1M guanidium hydrochloride.
For cryo-protection, they were transferred to a solution with 1.5X reservoir solution and
20% 2,3 butanediol or 20% DMSO for 10 seconds and immediately flash frozen under
liquid nitrogen. Form III crystals grew under 0.1 M phosphate citrate pH 5.5, 0.15 M sodium
sulfate, 0.01 M MgCl2, 1 M guanidium hydrochloride, and 28 % PEG 3350. The crystals
were grown at 30°C and were cryo-protected by increasing the PEG concentration to 40%.
Further details of crystallization are presented in the Supplementary Information.
Data collection and structure determination
Diffraction data were collected at the Dupont Northwestern Dow and Life Science
Collaborative Access Team stations (DND and LS CAT) at the Advanced Photon Source in
Argonne National Laboratory. Data collection and refinement statistics are shown in Table I.
All data were processed and integrated using XDS (Kabsch, 1993) and scaled with SCALA
(Collaborative-Computational-Project-4, 1994). Data on the Topo-31 crystals were collected
to 2.4 Å resolution. The structure was solved by Molecular Replacement (McCoy et al.,
2007) using the topoisomerase domain from the Topo-61 structure (residues 1-266) (Taneja
et al., 2006) as the search model. It was refined with refmac5 (Murshudov et al., 1997) and
Phenix (Afonine et al., 2005) to a final Rwork of 20.0 % and Rfree of 24.8 %. Topo-44 Form I
crystals diffract to 1.8 Å. The structure of Form I crystals was solved by Molecular
Replacement (McCoy et al., 2007) using the topoisomerase domain from the Topo-61
structure as the search model. Model rebuilding was performed using coot (Emsley and
Cowtan, 2004), and refinement using refmac5 (Murshudov et al., 1997). The final Rwork and
Rfree are 17.5 % and 22.0 % respectively.
For Topo-44 Form II and Form III crystals, seleno-methionine derivatized crystals were
used for single-wavelength anomalous dispersion (SAD) phasing. AutoSharp (Vonrhein et
al., 2007) was used for locating the selenium atoms; model building was done using coot
(Emsley and Cowtan, 2004), and refinement was carried out using refmac5 (Murshudov et
al., 1997) Three phosphate ions were noticed in the Form II structure; two of which present
in the topoisomerase active site and are separated by a distance of ~7.5 Å. The structure was
refined to a final Rwork of 24.1 % and Rfree of 28.9 %. Topo-44 Form III crystals diffracted
to 1.4 Å. The final Rwork and Rfree are 16.5 % and 18.4%, respectively. An interesting
observation is the presence of both phosphate and guanidium ions in the Form III Topo-44
structure. The linker helix and part of the first HhH motif of the B monomer show alternate
conformations and were built as two separate chains with occupancy of 0.5 each. Further
details on data collection and structure determination are given in the Supplementary
Information.
All figures were made with Pymol (DeLano, 2002) and the electrostatic surfaces were
calculated with APBS (Baker et al., 2001).
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Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We acknowledge staff and instrumentation support from the Keck Biophysics Facility and the Center for Structural
Biology at Northwestern University, and DND and LS-CAT at the Advanced Photon Source (APS) at Argonne
National Laboratory. Support from the R.H. Lurie Comprehensive Cancer Center of Northwestern University to the
Structural Biology Facility is also acknowledged. DND-CAT is supported by Dupont, DOW and the NSF. LS-CAT
was supported by the Michigan Economic Development Corporation and the Michigan Technology Tri-Corridor.
Use of the APS is supported by the Department of Energy (DOE). Research was supported by NIH grant GM51350
(to AM).
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Figure 1. Organization of topoisomerase V
Topoisomerase V is a multi-domain protein consisting of 24 helix-hairpin-helix (HhH) DNA
binding motifs arranged as 12 (HhH)2 domains following the N-terminal topoisomerase
domain. A) Schematic diagram of various topoisomerase V fragments. The topoisomerase
domain is shown in red, the (HhH)2 domains are shown in alternating colors of cyan and
yellow. The (HhH)2 domains with repair activity are shown in green. All fragments shown
have topoisomerase activity, but only the full length protein and the Topo78 fragment have
repair activity. B) Crystal structure of Topo-61 fragment (Taneja et al., 2006). The coloring
scheme is the same as in Figure 1A, except that the linker helix is shown in grey.
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Figure 2. DNA relaxation activity and EMSA for Topo-31, Topo-44 and Topo-78 fragments of
topoisomerase V
A) pH profile of the DNA relaxation activity of Topo-78 and Topo-44 fragments. 0.2 μg of
pUC19 DNA were incubated with 0.1 μg of Topo-78 or 1.5 μg of Topo-44 proteins at 65°C
for 15 minutes for the relaxation reaction. The reaction buffer contained 50 mM of the
appropriate buffer, 30 mM NaCl and 0.2 mM EDTA. Topo-78 relaxes DNA at a wider pH
range (5 to 9) than Topo-44, which relaxes DNA efficiently only at pH 5. DNA relaxation
activity of Topo-31 (B) and Topo-44 (C) fragments in the absence and presence of MgCl2.
0.2 μg of pUC19 DNA were incubated with 9 μg of Topo-31 or 1.5 μg of Topo-44 proteins
at 65°C for 15 minutes for the relaxation reaction. The reaction buffer contained 50 mM of
the appropriate buffer, 30 mM NaCl and 5 mM EDTA or 1 mM MgCl2. Both Topo-31 and
Topo-44 fragments can relax DNA in the absence of MgCl2, but MgCl2 enhances the DNA
relaxation activity of the topoisomerase V fragments. The black triangle in panels A, B and
C represents increasing pH from 4 to 10 by one pH unit. D) EMSA of Topo-44 and Topo-78
fragments with a 39mer double stranded DNA. Both Topo-44 and Topo-78 form stable
complexes with DNA, although Topo-78 seems to saturate DNA binding while Topo-44
does not. In addition, Topo-44 shows some cleavage of the DNA (bottom free DNA band),
while the cleavage is not apparent in Topo-78. The numbers at the bottom represent the
molar ratio of protein to DNA.
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Figure 3. Structure of Topo-44 fragments
A) Overlay of Form I (green), Form II (B subunit: blue), and Form III (B subunit: magenta)
structures onto the Topo-61 structure (B subunit: orange). The topoisomerase domains
superimpose very well for all the structures, while the linker helix and (HhH)2 domains
show differences in orientation. B) Overlay of the linker helices of Form I, II, and III
structures with that of Topo-61. The color scheme is same for all the figures unless
mentioned otherwise. Note that the linker helices have the same orientation at the start and
they change as they move further down the helix. C) Superposition of Form I, II, and III
Topo-44 structures with that of Topo-61. Only the (HhH)2 domains are colored while the
remaining parts are shown in gray for clarity. The active site residues are shown as orange
sticks. Note that the (HhH)2 domains adopt different orientations in all the structures. D)
Orientation of the (HhH)2 domains of Form I, II and Topo-61 structures. In Form I and II
structures, the (HhH)2 domains are moved away from the topoisomerase domain. For clarity,
the (HhH)2 domains of Form III are not shown. In panels C and D, the topoisomerase
domains were superposed to emphasize the different orientation of the other domains.
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Figure 4. Phosphate ions present near the active site of the Topo-44 structure
A) Stereo view of a Form III difference electron density map calculated with a model not
including the phosphates. The electron density is contoured at 3.7σ and shows the
tetrahedral shape of the phosphate ions. The active site residues are shown in stick. B)
Stereo view of the interaction of the phosphate ions with the putative active site residues.
The B subunit of Form II structure was superimposed onto the B subunit of Form III
structure and the phosphates ions from both structures are shown together with the Form II
B subunit protein backbone. The interactions made by the phosphate ion with the active site
residues and the corresponding distances in Å are represented as black dotted lines.
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Figure 5. Representation of the unique phosphate ions from Form II and Form III Topo-44
structures
A) An overlay of the A (magenta) and B (brown) subunits of the Form III structure and B
(blue) subunit of Form II Topo-44 structures. The positions of eight unique phosphate ions
(orange spheres) are shown. Note that most phosphate ions are found along the DNA
binding groove of the topoisomerase domain. B) The phosphate ions in the DNA binding
groove are separated by distances of around 7 Å. The protein backbone is that of the B
subunit of Form III structure. The active site residues are represented as sticks and distances
in Å between adjacent phosphate ions are shown as black dotted lines.
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Figure 6. Model showing DNA bound to the topoisomerase domain
A) Model of a 17-mer double stranded DNA bound to the Topo-31 structure (teal). The
DNA is represented as green sticks, where as phosphate ions are represented as orange
sticks. DNA binds along the DNA binding groove and five of the eight phosphate ions noted
in the Topo-44 structures coincide with the DNA backbone. B) Model of Topo-44 (Form II,
B subunit: blue) binding to 17-mer double stranded DNA. Note that the linker helix and the
(HhH)2 domains interfere with DNA binding to the topoisomerase domain and are likely to
move away to allow binding. C) Electrostatic surface representation of the Topo-31
structure. The positively charged DNA binding groove is clearly visible and the phosphate
ions are bound in this groove. The orientation corresponds to a 90° rotation of the one shown
in Figure 6A in the direction of the arrow. Note that the DNA binding groove goes from one
end of the molecule to the other and it is narrower at one end (start of the linker helix) and
wider at the other end. The putative active site residues (green sticks) are located at the
wider end of the groove. Other residues lining the groove and interacting with the phosphate
ions are shown as cyan sticks. D) Electrostatic surface representation of Topo-31 with
phosphate ions (orange) and DNA (green). Three phosphate ions (P3, P4, and P5) coincide
with the phosphates of one of the DNA strands, where as P1 coincides with a phosphate of
the opposite DNA strand. The model shows that the DNA binding groove of topoisomerase
V is wide enough to bind DNA and that the movement of linker helix and (HhH)2 domains
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are required to accommodate the DNA. The electrostatic potential was calculated with a
dielectric constant of 80 for solvent and 2 for protein. The surface is colored with a blue to
red gradient from +10 to −10 KbT/ec.
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Table 1
Data collection and refinement statistics
Topo-31
Topo-44 Form I
Topo-44 Form II
Topo-44 Form III
Data Collection
Space group
C2221
C121
P41212
P212121
Cell dimensions
a=106.7 Å, b=119.4
Å, c=63.7 Å
a=104.2 Å, b=47.7 Å,
c=81.2 Å (β=112.48)
a=b=70.1 Å, c=349.6 Å
a=63.6 Å, b=80.1 Å,
c=137.2 Å
Resolution (Å)a
79.56 – 2.4 (2.53 –
2.4)
75.05 – 1.82 (1.91 –
1.82)
29.5- 2.6 (2.72-2.6)
28.9-1.4 (1.46-1.4)
Number of observed
reflections
78,729 (11,538
134,411 (13,220)
227,408 (19,917)
1,157,917 (126,319)
Number of unique reflections
16,259 (2,346)
32,998 (4,301)
28,151 (3,331)
136,662 (15,986)
Completeness (%)
99.8 (99.8)
98.3 (88.6)
99.9 (100.0)
98.8 (95.5)
Multiplicity
4.8 (4.9)
4.1 (3.1)
8.1 (6.0)
8.5 (7.9)
Rmerge (%)b
4.7 (71.1)
4.0 (16.3)
7.4 (52.2)
4.5 (37.9)
Rmeas (%)c
5.3 (79.6)
4.6 (19.4)
7.9 (57.2)
4.8 (40.5)
≪I>/σ(<I>)>d
20.5 (2.5)
23.0 (6.8)
19 (3.2)
27.5 (5.3)
Refinement
Resolution (Å)
79.56 - 2.4 (2.46 -
2.4)
28.06 -1.82 (1.87 –
1.82)
29.14 – 2.6 (2.67 – 2.6)
28.9 - 1.4 (1.44 - 1.4)
Number of reflections
working/test
15,419/821
31,317/1,673
26,710/1,438
129,802/6,859
Rwork (%)e
20.0(24.3)
17.5 (17.9)
24.1(36.6)
16.5 (19.3)
Rfree(%)f
24.8 (31.1)
22.0 (24.8)
28.9 (45.1)
18.4 (22.1)
Protein residues/atomsg
269/2,203
376/3212
727/5,970
738/7,511
Atoms in alternate
conformations
0
258 (20 protein
residues)
8 (1 protein residue)
2846 (157 protein
residues)
Water molecules
29
238
30
573
Other atoms
-
-
3 PO4
7 PO4, 3 Gmh, 3 Mg++, 2
Cl−
B-factor (Å2)
Protein atoms (chain)
68.4
22.8
A:53.8; B:58.2
A:13.4; B:14.9
Water molecules
59.1
29.3
40.0
23.7
r.m.s. deviations
bond lengths (Å)
0.015
0.006
0.01
0.009
bond angles (°)
1.42
0.920
1.2
1.2
Ramachandran ploti
Favored regions (%)
94.3
98.9
96.2
98.5
Outliers (%)
0.0
0.0
0.3
0
aNumbers in parenthesis correspond to highest resolution shell.
bRmerge= Σ|I − <I>|/ΣI, where I is the observed intensity and <I> the average intensity obtained from multiple measurements.
Structure. Author manuscript; available in PMC 2011 July 14.
NIH-PA Author Manuscript
NIH-PA Author Manuscript
NIH-PA Author Manuscript
Rajan et al.
Page 20
cRmeas as described in Diederichs and Karplus (Diederichs and Karplus, 1997).
d≪I>/σ(<I>)> = Mean Ih over the standard deviation of the mean Ih averaged over all reflections in a resolution shell.
eRwork= Σ ||Fo| − |Fc||/Σ|Fo|, where |Fo| is the observed structure factor amplitude and |Fc| the calculated structure factor amplitude.
fRfree: Rfactor based on 5% of the data excluded from refinement.
gTotal number of protein atoms, including those in alternate conformations.
hGm: guanidinum ion.
iAs reported by Molprobity (Davis et al., 2004).
Structure. Author manuscript; available in PMC 2011 July 14.
|
3M7F
|
Crystal structure of the Nedd4 C2/Grb10 SH2 complex
|
Structural Basis for the Interaction between the Growth
Factor-binding Protein GRB10 and the E3 Ubiquitin Ligase
NEDD4*
Received for publication,May 10, 2010, and in revised form, October 11, 2010 Published, JBC Papers in Press,October 26, 2010, DOI 10.1074/jbc.M110.143412
Qingqiu Huang1 and Doletha M. E. Szebenyi
From MacCHESS, Cornell University, Ithaca, New York 14853
In addition to inhibiting insulin receptor and IGF1R kinase
activity by directly binding to the receptors, GRB10 can also
negatively regulate insulin and IGF1 signaling by mediating
insulin receptor and IGF1R degradation through ubiquitina-
tion. It has been shown that GRB10 can interact with the C2
domain of the E3 ubiquitin ligase NEDD4 through its Src ho-
mology 2 (SH2) domain. Therefore, GRB10 might act as a con-
nector, bringing NEDD4 close to IGF1R to facilitate the ubiq-
uitination of IGF1R by NEDD4. This is the first case in which
it has been found that an SH2 domain could colocalize a ubiq-
uitin ligase and its substrate. Here we report the crystal struc-
ture of the NEDD4 C2-GRB10 SH2 complex at 2.0 A˚ . The
structure shows that there are three interaction interfaces be-
tween NEDD4 C2 and GRB10 SH2. The main interface centers
on an antiparallel -sheet composed of the F -strand of
GRB10 SH2 and the C -strand of NEDD4 C2. NEDD4 C2
binds at nonclassical sites on the SH2 domain surface, far from
the classical phosphotyrosine-binding pocket. Hence, this in-
teraction is phosphotyrosine-independent, and GRB10 SH2
can bind the C2 domain of NEDD4 and the kinase domain of
IGF1R simultaneously. Based on these results, a model of how
NEDD4 interacts with IGF1R through GRB10 has been pro-
posed. This report provides further evidence that SH2 do-
mains can participate in important signaling interactions be-
yond the classical recognition of phosphotyrosine.
The GRB7 (growth factor receptor-binding protein) family
of adaptor proteins includes GRB7, GRB10, and GRB14.
These proteins share a conserved molecular architecture: a
proline-rich N-terminal region, a Ras-associating-like do-
main, a pleckstrin homology domain, a family-specific BPS
region, and a conserved C-terminal Src homology 2 (SH2)2
domain (1, 2). Their SH2 domains have the ability to recog-
nize phosphotyrosine-containing peptides on a variety of acti-
vated tyrosine kinase receptors. GRB7 has been shown to in-
teract with EGF receptor, ErB2 receptor, EphB1, focal
adhesion kinase, and platelet-derived growth factor receptor
and to be involved in regulating cell migration (3, 4). GRB10
and GRB14 have been shown to interact with insulin receptor
(IR), IGF1R (insulin-like growth factor 1 receptor), EGF re-
ceptor, Raf1 kinase, and MEK1 kinase and to be involved in
cell growth regulation (5–11). The Grb10 gene is maternally
imprinted in mice. When the Grb10 gene was disrupted by a
gene trap insertion, the mutant mice were 30% greater in
size than normal, with disproportionately large livers (5, 12).
As adults, these mutant mice had improved glucose tolerance,
increased muscle mass, and reduced adiposity (13, 14). Fur-
thermore, Grb10 transgenic mice overexpressing GRB10
showed growth retardation and insulin resistance (15). These
results indicate that GRB10 plays a negative role in cell
growth, as a consequence of hypernegative regulation of the
IR and IGF1R. Mice lacking the Grb14 gene were of normal
size and had improved glucose tolerance and increased insu-
lin signaling in muscle and liver (16). Therefore, GRB10 and
GRB14 are tissue-specific negative regulators of insulin and
IGF1 signaling.
Additional research results indicate that GRB10 and
GRB14 might contribute to type 2 (non-insulin-dependent)
diabetes in humans (5, 17, 18). From a genome-wide associa-
tion scan in the Old Order Amish, the GRB10 gene has been
identified as having the strongest association between type 2
diabetes and a single nucleotide polymorphism (SNP) (19). In
subcutaneous adipose tissue, the GRB14 mRNA levels in type
2 diabetes patients were 43% higher than those in normal per-
sons (18).
The mechanisms of the negative regulation of IR and
IGF1R by GRB10 and GRB14 are not yet clear. Biochemical
studies have shown that GRB10 and GRB14 can bind IR and
IGF1R through their BPS and SH2 domains (20–22). The
GRB14 BPS region binds as a pseudosubstrate inhibitor in the
tyrosine kinase domain of IR to suppress insulin signaling
(20). Suppressing endogenous GRB10 expression led to in-
creased IR protein levels, whereas overexpression of GRB10
led to reduced IR protein levels (7). Reduced IR levels were
observed in cells with prolonged insulin treatment, and this
reduction was inhibited in GRB10-deficient cells (7). The in-
sulin-induced IR reduction was largely reversed by MG-132, a
proteasomal inhibitor, but not by chloroquine, a lysosomal
inhibitor. IR undergoes insulin-stimulated ubiquitination in
cells, and this ubiquitination was inhibited in the GRB10-sup-
* This work was supported, in whole or in part, by National Institutes of
Health, NCRR, Grant RR-01646. This work is based upon research con-
ducted at the Cornell High Energy Synchrotron Source (CHESS), which is
supported by the National Science Foundation under Award DMR
0225180.
The atomic coordinates and structure factors (code 3M7F) have been deposited
in the Protein Data Bank, Research Collaboratory for Structural Bioinformat-
ics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).
1 To whom correspondence should be addressed: MacCHESS, Cornell Uni-
versity, Ithaca, NY 14853. Tel.: 607-255-9386; Fax: 607-255-9001; E-mail:
qh24@cornell.edu.
2 The abbreviations used are: SH2, Src homology 2; IR, insulin receptor; IPTG,
isopropyl 1-thio--D-galactopyranoside; r.m.s., root mean square.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 53, pp. 42130–42139, December 31, 2010
© 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
42130
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 53•DECEMBER 31, 2010
pressed cell line (7). Therefore, in addition to inhibiting IR
kinase activity by directly binding to IR through its BPS and
SH2 domains, GRB10 also negatively regulates insulin signal-
ing by affecting insulin-stimulated degradation of the recep-
tor, possibly through regulation of ubiquitination of the
receptor.
GRB10 and GRB14 do not themselves have ubiquitin ligase
activity. However, it has been shown that GRB10 binds the C2
domain of the E3 ubiquitin ligase NEDD4 through its SH2
domain, and formation of the complex promotes IGF1-stimu-
lated multiubiquitination, internalization, and degradation of
the IGF1R (23–25). GRB10 and NEDD4 remain associated
with the IGF1R in early endosomes and caveosomes (23).
Therefore, the SH2 domain of GRB10 might act as a connec-
tor between the C2 domain of NEDD4 and the kinase domain
of IGF1R; proximity of NEDD4 and IGF1R would facilitate
the ubiquitination of IGF1R by NEDD4. However, SH2 is typ-
ically highly specialized for the recognition of phosphoty-
rosine and has only one binding site for phosphotyrosine-
containing peptides (the Tyr(P)-binding pocket). It was not
clear how the SH2 domain of GRB10 can bind the C2 domain
of NEDD4 and the kinase domain of IGF1R simultaneously,
although the possibility of a second binding site was suggested
by the observation that the NEDD4 C2-GRB10 SH2 interac-
tion is phosphorylation-independent (25). Here we report the
crystal structure of the NEDD4 C2-GRB10 SH2 complex,
which clearly shows that the SH2 domain of GRB10 binds the
C2 domain of NEDD4 through a surface region distinct from
the classical Tyr(P)-binding pocket.
EXPERIMENTAL PROCEDURES
Protein Expression and Purification—The cDNA encoding
the mouse GRB10 SH2 domain (residues 429–536) was am-
plified from the plasmid pcDNA-Grb10 (a kind gift from Dr.
Junlin Guan) and inserted into the expression vector pQE80
to make a pQE80-SH2 expression construct. This construct
was transferred into BL21(DE3) (Novagen) for protein expres-
sion. Protein expression was induced at 37 °C for 4 h with 0.3
mM IPTG. The cells were harvested by centrifugation and
then suspended in binding buffer (500 mM NaCl, 50 mM Tris-
HCl, pH 8.5, 10 mM imidazole, 5 mM -mercaptoethanol, and
1 mM benzamidine chloride). Cell lysis was carried out by son-
ication. After centrifugation, the supernatant was applied to a
nickel affinity column. After protein binding, the column was
washed thoroughly with 100 volumes of binding buffer fol-
lowed by 10 volumes of washing buffer (500 mM NaCl, 50 mM
Tris-HCl, pH 8.5, 40 mM imidazole, 5 mM -mercaptoethanol,
and 1 mM benzamidine chloride). The protein was then eluted
from the column with 5 volumes of elution buffer (200 mM
NaCl, 300 mM imidazole-HCl, pH 7.5, 5 mM -mercaptoetha-
nol, and 1 mM benzamidine chloride). The protein solution
was concentrated and further purified by FPLC using a
Superdex 200 column (GE Healthcare) with an elution buffer
containing 0.15 M NaCl and 5 mM Tris-HCl (pH 7.5).
The cDNA encoding the full-length mouse GRB10 (an
isoform of GRB10) was inserted into the expression vector
pMAL-2C to make a pMAL-2C-Grb10 expression construct.
This construct was transferred into BL21(DE3) for protein
expression. The harvested cells were lysed in the binding
buffer (100 mM NaCl, 50 mM Tris-HCl, pH 8.5, 5 mM -mer-
captoethanol, and 1 mM benzamidine chloride) by sonication,
and the supernatant was applied to a maltose-agarose column.
The column was washed thoroughly with 100 volumes of
binding buffer, and the maltose-binding protein-GRB10 fu-
sion protein was eluted with 3 volumes of elution buffer (100
mM NaCl, 50 mM maltose, 50 mM Tris-HCl, pH 8.0, 2 mM
CaCl2). After the addition of Factor Xa, this elution solution
was incubated at 4 °C for 48 h to cleave off the maltose-bind-
ing protein tag. The protein solution was then concentrated
and purified with a Superdex 200 column.
The cDNA encoding the mouse NEDD4 C2 domain (resi-
dues 108–287) was amplified from the plasmid pBS-Nedd4 (a
kind gift from Dr. Andrea Morrione) and inserted into the
expression vector pSUMO (Invitrogen) to make a pSUMO-
Nedd4 C2 expression construct. This construct was trans-
ferred into BL21(DE3) for protein expression. Protein expres-
sion was induced at 22 °C overnight with 0.3 mM IPTG. The
protein was purified with a nickel column using the method
described above. After the column was washed thoroughly
with 100 volumes of binding buffer followed by 10 volumes of
washing buffer, the protease SUMOase was added to the col-
umn and incubated at 4 °C for 24 h to cleave off the
SUMOHis tag. The free protein C2 was eluted, concentrated,
and further purified with a Superdex 200 column. Trunca-
tions of NEDD4 C2 (residues 108–250, 200–300, and 115–
287, respectively) were cloned, expressed, and purified using
the same procedure as for wild type NEDD4 C2.
To express GRB10 SH2 without any tag, the cDNA encod-
ing the mouse GRB10 SH2 domain (residues 429–536) was
inserted into the expression vector pCDFDuet-1 to make
pCDFDuet-SH2. To prepare the NEDD4 C2-GRB10 SH2
complex for crystallization, the plasmids pSUMO-C2 and
pCDFDuet-SH2 were co-transferred into the host cell
BL21(DE3) for co-expression. In this system, the expression
level of NEDD4 C2-SUMOHis was higher than that of
GRB10 SH2. Protein expression was induced at 15 °C for 24 h
with 0.3 mM IPTG. The cell pellet was suspended in binding
buffer (200 mM NaCl, 50 mM Tris-HCl, pH 8.5, 10 mM imidaz-
ole, 5 mM -mercaptoethanol, and 1 mM benzamidine chlo-
ride) and lysed by sonication. After centrifugation, the super-
natant was applied to a nickel affinity column. After protein
binding, the column was washed thoroughly with 100 vol-
umes of binding buffer followed by 10 volumes of washing
buffer (200 mM NaCl, 50 mM Tris-HCl, pH 8.5, 40 mM imidaz-
ole, 5 mM -mercaptoethanol, and 1 mM benzamidine chlo-
ride). Then protease SUMOase was added to the column and
incubated at 4 °C for 24 h to cleave off the SUMOHis tag. The
free proteins (NEDD4 C2-GRB10 SH2 complex and isolated
NEDD4 C2) were eluted, concentrated, and purified with a
Superdex 200 column. The fractions containing the NEDD4
C2-GRB10 SH2 complex were collected and further purified
using a SourceQ column (GE Healthcare) for FPLC.
Effect of Calcium on the Interaction—GRB10 SH2 (with an
N-terminal His tag, GRB10 SH2His) was mixed with NEDD4
C2 (without tag) in a 1:1 molar ratio (as judged by A280) in the
incubating buffer (150 mM NaCl and 50 mM Tris-HCl, pH 8.5)
Structure of the NEDD4 C2-GRB10 SH2 Complex
DECEMBER 31, 2010•VOLUME 285•NUMBER 53
JOURNAL OF BIOLOGICAL CHEMISTRY 42131
containing 20 mM EGTA or 20 mM CaCl2, on ice, for 2 h. Sim-
ilarly, GRB10 (without tag) was mixed with NEDD4
C2SUMOHis in a 1:1 molar ratio in the incubating buffer
containing 20 mM EGTA or 20 mM CaCl2, on ice, for 2 h.
Then EGTA or CaCl2 in the mixtures was removed by a sizing
column. The fraction containing proteins from the sizing col-
umn was collected and applied onto a nickel affinity column.
After protein binding, the column was washed with 100 vol-
umes of incubating buffer and further washed with 10 vol-
umes of washing buffer (150 mM NaCl, 50 mM Tris-HCl, and
40 mM imidazole, pH 8.5). 15 l of gel slurry was drawn for
analysis by SDS-PAGE.
Effect of Ionic Strength on the Interaction—GRB10 SH2His
and NEDD4 C2 were mixed in a 1:1 molar ratio in the incu-
bating buffer (50 mM Tris-HCl, pH 8.5). After being incubated
on ice for 2 h, the NEDD4 C2-GRB10 SH2His complex was
bound to a nickel column. Then the column was eluted with
incubating buffer containing different NaCl concentrations
(0, 100, 200, 300, 400, 500, and 1000 mM, sequentially). For
each concentration, 2 volumes of incubating buffer were used.
The eluted solutions were analyzed by SDS-PAGE.
The Specificity of the Interaction—The cDNAs encoding
mouse GRB14 SH2 domain and GRB7 SH2 domain were sep-
arately inserted into the expression vector pGEX4T-1. The
resulting constructs pGEX4T-GRB14 SH2 and pGEX4T-
GRB7 SH2 were co-transferred, respectively, with pSUMO-
NEDD4 C2 into the host cell BL21(DE3). Protein expression
was induced with 0.3 mM IPTG at 15 °C for 24 h. The cells
were collected and resuspended in PBS for lysis by sonication.
The supernatant from the cell lysis was purified using a nickel
affinity column and a glutathione-agarose column sequen-
tially. After the column was washed thoroughly, 15 l of gel
slurry was drawn and analyzed by SDS-PAGE.
The cDNAs encoding mouse GRB14 and GRB7 were sepa-
rately inserted into the expression vector pQE80 to express
the target proteins with N-terminal His tags. Purified
GRB14His (or GRB7His) was mixed with NEDD4 C2 (with-
out tag) in a 1:1 molar ratio in the incubating buffer (150 mM
NaCl and 50 mM Tris-HCl, pH 8.5). After being incubated on
ice for 2 h, the mixture was applied to a nickel affinity col-
umn. After protein binding, the column was washed by 100
volumes of incubating buffer followed by 10 volumes of wash-
ing buffer (150 mM NaCl, 50 mM Tris-HCl, and 40 mM imid-
azole, pH 8.5). Then 15 l of gel slurry was drawn for analysis
by SDS-PAGE.
Crystallization and Data Collection—The purified NEDD4
C2-GRB10 SH2 complex showed two protein bands on SDS-
polyacrylamide gel, corresponding to NEDD4 C2 and GRB10
SH2, respectively (Fig. 1A). The NEDD4 C2-GRB10 SH2 com-
plex solution was desalted and concentrated to 20 mg/ml.
Crystals were grown by the hanging drop vapor diffusion
method at 18 °C. Typically, 2 l of the protein stock was
mixed with 2 l of the reservoir solution consisting of 35%
MPD (v/v) and 0.1 M Hepes-HCl buffer, pH 7.5. Crystals were
observed after 2 days and reached a typical size of 100
100 200 m3 1 month later. Diffraction data were collected
at 100 K on beamline A1 at MacCHESS. The diffraction data
were reduced using the HKL package (26), and the statistics of
data collection and processing are summarized in Table 1.
Structure Refinement—The structure of the NEDD4 C2-
GRB10 SH2 complex was solved by the molecular replace-
ment program Phaser (27) using the structure of human
GRB10 SH2 (28) (Protein Data Bank code 1NRV) and the
structure of human NEDD4 C2 (Protein Data Bank code
3B7Y) as the search models. The structure was refined using
Refmac5 (29) and PHENIX (30). The refinement statistics are
given in Table 1. The B factors are constant throughout the
protein. For the GRB10 SH2 subunit (average B factor 33.7
Å2), the B factors of its N terminus (residues 429–438) and C
terminus (residues 526–535) are 32.1 and 37.6 Å2, respec-
tively. For the NEDD4 C2 subunit (average B factor 39.3
Å2), the B factors of its N terminus (residues 112–120) and C
terminus (residues 281–287) are 41.1 and 40.5 Å2, respec-
tively. The interface between GRB10 SH2 and NEDD4 C2 was
analyzed using the program PISA (31) from the CCP4 suite
(32). SDS-PAGE analysis of the NEDD4 C2-GRB10 SH2 crys-
tals showed that the NEDD4 C2 and the GRB10 SH2 in the
crystals had the same molecular weights as the original
NEDD4 C2 and the original GRB10 SH2, respectively (Fig.
1B), suggesting that both NEDD4 C2 and GRB10 SH2 in the
crystal are complete and have not been degraded.
Mutation—Site-directed mutations of GRB10 SH2 (K505P,
N519A, R533A, R431A and H434A, respectively) were carried
out using the Phusion mutation kit (New England Biolabs).
The interaction between the GRB10 SH2 mutants and
NEDD4 C2 was analyzed using the co-expression system de-
scribed above.
RESULTS
The NEDD4 C2-GRB10 SH2 Interaction Is Ca2-indepen-
dent and Phosphorylation-independent—As shown by the yeast
two-hybrid method, GRB10 can interact with the C2 domain
of NEDD4 through its SH2 domain (25). The C2 domain is a
conserved module of about 120 residues, which was first
found in protein kinase C (33). In most proteins, the C2 do-
main binds phospholipids in a Ca2-dependent manner (34,
35); in some proteins, it mediates protein-protein interac-
tions (36). The C2 domain of NEDD4 has been shown to
target NEDD4 to the plasma membrane in response to
Ca2 (37). However, the interaction between NEDD4 and
GRB10 has been demonstrated to be Ca2-independent by
the co-immunoprecipitation method (25). We performed a
series of in vitro interaction experiments using purified C2
domain and purified SH2 domain (or full-length GRB10).
As shown in Fig. 1A, NEDD4 C2 could form a complex
with GRB10 SH2 in the presence of either 20 mM EGTA or
20 mM CaCl2, suggesting that the interaction between
NEDD4 C2 and GRB10 SH2 was Ca2-independent.
NEDD4 C2 could also form a complex with full-length
GRB10 (an isoform of GRB10), and this interaction was
also Ca2-independent (Fig. 1A).
The SH2 domain is a conserved module of about 100 resi-
dues, which is highly specialized for the recognition of phos-
photyrosine, with only a few exceptions to date (38). How-
ever, GRB10 has been suggested to preferentially associate
Structure of the NEDD4 C2-GRB10 SH2 Complex
42132
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 53•DECEMBER 31, 2010
with unphosphorylated NEDD4 (based on co-immunopre-
cipitation experiments) (25). In our work, NEDD4 C2 was
expressed in Escherichia coli, in which expressed proteins
cannot be phosphorylated. The NEDD4 C2 domain produced
in this system could form a complex with GRB10 SH2 or
GRB10, indicating that binding of GRB10 SH2 to NEDD4 C2
is phosphorylation-independent.
Both the NEDD4 C2-GRB10 SH2 complex and the NEDD4
C2-GRB10 complex were stable in low ionic strength solu-
tion (200 mM NaCl). The complexes dissociated in a solu-
tion containing more than 500 mM NaCl, in the presence or
absence of 20 mM EGTA (or 20 mM CaCl2) (data not shown).
These results suggested that the main interaction between
NEDD4 C2 and GRB10 SH2 was not hydrophobic and was
unaffected by Ca2.
The molecular mass of GRB10 SH2 is about 12 kDa. The
molecular mass of NEDD4 C2 is about 19 kDa. As determined
by a Superdex200 sizing column, the molecular mass of the
NEDD4 C2-GRB10 SH2 complex was about 30 kDa, suggest-
ing that this complex is a heterodimer in solution (Fig. 1C).
The Specificity of the Interaction—All members of the GRB7
adaptor protein family (GRB7, GRB10, and GRB14) contain a
Ras-associating-like domain, a pleckstrin homology domain, a
family-specific BPS region, and a C-terminal SH2 domain (1,
2). Because the SH2 domains are highly conserved among this
family (Fig. 2), we investigated whether NEDD4 C2 could also
bind the SH2 domains of GRB7 and GRB14, using the co-
expression method. In the E. coli co-expression system,
NEDD4 C2 was expressed with a SUMOHis tag, which could
FIGURE 2. Alignment of the amino acid sequences of the SH2 domains
of GRB7, GRB10, and GRB14. The -helices and -strands of GRB10 SH2
are highlighted with red lines and blue arrows, respectively. The underlined
peptide of GRB10 is the F -strand, which forms an antiparallel -sheet with
the C -strand of NEDD4 C2 in the NEDD4 C2-GRB10 SH2 complex. The resi-
dues buried in the interface with NEDD4 C2 are shown with red letters. The
two residues in interface I (Lys505 and Asn519) chosen for mutation are high-
lighted as red boldface letters.
TABLE 1
Data collection and structure refinement statistics
Parameters
Values
Data collection
Space group
P212121
Cell dimensions a, b, c (Å)
52.31 70.30 86.47
Resolution (Å)
50-2.0 (2.03-2.00)a
No. of unique observations
21,866 (1023)
Redundancy
6.5 (4.4)
Completeness (%)
99.4 (96.8)
Average I/I
35.7 (3.4)
Rmerge (%)
5.2 (36.2)
Structure refinement
No. of protein atoms/waters
2011/95
Resolution (Å)
50-2.0
Rwork (%) /Rfree (%)
19.2/23.2
r.m.s. deviations
Bonds (Å)/Angles (degrees)
0.006/1.057
Average B factor (Å2)
36.78
Ramachandran plot
Most favored (%)
95.34
Allowed (%)
4.66
a Values in parentheses are for the highest resolution shell.
FIGURE 1. Interaction between NEDD4 C2 and GRB10 SH2. The two pro-
teins were mixed and incubated in the incubating buffer for 2 h. After being
desalted with a sizing column, the mixture was applied onto a nickel affinity
column. After being washed with washing buffer, 15 l of gel slurry was
drawn for analysis by SDS-PAGE (for details, see “Experimental Procedures”).
A, GRB10 could form a stable complex with NEDD4 C2SUMOHis in the
presence of 20 mM EGTA (lane 3) or 20 mM CaCl2 (lane 4) or without any ad-
ditive (lane 5); GRB10 alone did not bind to the nickel beads (lane 2);
NEDD4 C2SUMOHis alone is shown in lane 1. GRB10 SH2His could form a
stable complex with NEDD4 C2 in the presence of 20 mM EGTA (lane 7) or 20
mM CaCl2 (lane 8) or without any additive (lane 9); NEDD4 C2 alone did not
bind to the nickel beads (lane 10); GRB10 SH2His alone is shown in lane 11.
Lane 6, standards. B, wild type (lane 1) and the mutants N519A (lane 3),
R533A (lane 12), R431A (lane 13), and H434A (lane 14) of GRB10 SH2His
could form a stable complex with NEDD4 C2, whereas the K505P mutant
(lane 2) could not form a complex with NEDD4 C2. NEDD4 C2 alone could
not bind to the nickel beads (lane 4); wild type GRB10 SH2His alone could
bind to the nickel beads (lane 10). Neither the truncated NEDD4 C2 (resi-
dues 108–250) (lane 5) nor the truncated NEDD4 C2 (residues 200–300)
(lane 6) could bind to GRB10 SH2His; the truncated NEDD4 C2 (residues
115–287) could bind to GRB10 SH2His (lane 7). Crystals of the NEDD4 C2-
GRB10 SH2 complex were picked out from the drop and analyzed by SDS-
PAGE (lane 9). Lanes 8 and 11, standards. C, elution profile of the NEDD4
C2-GRB10 SH2His complex from a Superdex 200 sizing column. The inset
shows the analysis of the peak fractions by SDS-PAGE.
Structure of the NEDD4 C2-GRB10 SH2 Complex
DECEMBER 31, 2010•VOLUME 285•NUMBER 53
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Structure of the NEDD4 C2-GRB10 SH2 Complex
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 53•DECEMBER 31, 2010
bind to a nickel-agarose column, whereas GRB7 SH2 and
GRB14 SH2 were expressed with a GST tag, which could bind
to a glutathione-agarose column but not a nickel column. If
the SH2 domain can form a complex with C2SUMOHis, it
should be co-purified with C2SUMOHis by a nickel column.
As shown by SDS-PAGE analysis, the protein purified by a
nickel column contained only NEDD4 C2SUMOHis,
whereas the protein purified by a glutathione-agarose column
contained only GRB7 SH2GST or GRB14 SH2GST. Al-
though both NEDD4 C2SUMOHis and GRB7 SH2GST (or
GRB14 SH2GST) were co-expressed in the same cell at high
levels, neither of the SH2 domains could be co-purified with
NEDD4 C2SUMOHis by a nickel column, suggesting that
neither GRB7 SH2 nor GRB14 SH2 could form a complex
with NEDD4 C2. As a control, we showed that GRB10
SH2GST could form a complex with NEDD4 C2SUMOHis,
suggesting that neither the GST tag nor the SUMO tag could
inhibit the interaction between NEDD4 C2 and the SH2
domain.
In vitro interactions between NEDD4 C2 and full-length
GRB7 (or GRB14) were also investigated using purified pro-
teins. As shown by SDS-PAGE analysis, NEDD4 C2 could not
form a complex with GRB7 or GRB14.
Crystal Structure of the NEDD4 C2-GRB10 SH2 Complex—
To gain further insights into the interaction between NEDD4
C2 and GRB10 SH2, we co-expressed NEDD4 C2 and GRB10
SH2 in E. coli and determined the crystal structure of the
NEDD4 C2-GRB10 SH2 complex at 2.0 Å resolution (Protein
Data Bank code 3M7F). Data collection and refinement statis-
tics are given in Table 1.
The NEDD4 C2-GRB10 SH2 complex exists as a het-
erodimer in the crystal. The most interesting features of the
NEDD4 C2-GRB10 SH2 complex structure are the interaction
interfaces between the two proteins (Fig. 3A). Interface I is the
major interface, with a buried area of 530 Å2. Central to this
interface is a small antiparallel -sheet formed from the F
-strand (residues 502–508) of GRB10 SH2 and the C
-strand (residues 153–160) of NEDD4 C2 (Fig. 3B). It has
been shown that a Pro residue can disrupt a -strand in pro-
teins (39). Therefore, the Lys505 residue of GRB10 SH2 was
mutated to Pro to disrupt the F -strand. The resultant
K505P mutant of GRB10 SH2 could not form a complex with
NEDD4 C2 in the E. coli co-expression system as wild type
GRB10 SH2 did (Fig. 1B). This result suggests that this antipa-
rallel -sheet plays an important role in stabilizing the
NEDD4 C2-GRB10 SH2 complex. Residues in -strand E and
helix B of GRB10 SH2 and in -strands A, B, and D of NEDD4
C2 also contribute to interface I. There are a total of eight
hydrogen bonds between SH2 and C2 in this interface, four
main chain-main chain in the small -sheet and four side
chain-main chain elsewhere. Two of the latter are from the
side chain of Asn519 in helix B of SH2 to main chain nitrogen
and oxygen atoms of Leu177 in the D -strand of C2 (Fig. 3C).
The N519A mutant of GRB10 SH2 could form a complex
with NEDD4 C2 (Fig. 1B), suggesting that this pair of hydro-
gen bonds is not critical to the formation of the GRB10 SH2-
NEDD4 C2 complex. There is a possible salt bridge between
the acid residue Asp514 of GRB10 SH2 and the alkaline resi-
due Arg179 of NEDD4 C2 (distance 4 Å). In this interface,
there are also hydrophobic interactions between the two pro-
teins, involving residues Phe496, Phe506, Leu512, Phe515,
Tyr516, and Leu518 of GRB10 SH2 with Leu148, Ile155, Leu156,
Val159, Ile176, Leu177, and Phe178 of NEDD4 C2 (Fig. 3C).
Interface II is the smallest interface, with an area of 90 Å2,
comprising the N terminus (residues 429–434) and part of
the B helix (residue Ile510) of GRB10 SH2 and the N terminus
(residues 112–114) of NEDD4 C2 (Fig. 3A). There is one hy-
drogen bond between the side chain of residue Gln433 of
GRB10 SH2 and the carbonyl group of residue Glu112 of
NEDD4 C2 (Fig. 4B). A truncated NEDD4 C2 (residues 115–
287) lacking the N terminus could form a stable complex with
GRB10 SH2 (Fig. 1B), suggesting that interface II does not
play an important role in stabilizing the NEDD4 C2-GRB10
SH2 complex.
Interface III is a medium-sized interface, with an area of
312 Å2. The proline-rich C terminus (residues 283–287) of
NEDD4 C2 rests against a surface made up of an N-terminal
part (residues 431–440) and a C-terminal part (residues 532–
535) of GRB10 SH2 (Fig. 3A). There is a hydrogen bond be-
tween the side chain of residue Arg533 of GRB10 SH2 and the
carbonyl group of residue Pro284 of NEDD4 C2 (Fig. 4C) and a
salt bridge between the side chain of Arg431 in SH2 and the
C-terminal carboxyl group of C2 (residue Pro287) (Fig. 4D).
The particular importance of this interface is suggested by the
observation that a 28-residue stretch preceding the C-termi-
nal Pro-rich cluster of C2 is disordered (and therefore invisi-
ble) in the crystal, but the cluster itself is well ordered and
clearly defined in electron density maps. A truncated NEDD4
C2 (residues 108–250), lacking the C-terminal 37 residues,
could not form a stable complex with GRB10 SH2 (Fig. 1B),
suggesting that interface III plays an important role in stabi-
lizing the NEDD4 C2-GRB10 SH2 complex. A single site mu-
tation of SH2 at this interface (R533A, R431A, or H434A)
could not disrupt the NEDD4 C2-GRB10 SH2 complex (Fig.
1B), indicating that interface III involves a number of inter-
acting residues and is not easily disrupted. NEDD4 C2 trun-
cated to residues 200–300, however, could not form a stable
complex with GRB10 SH2 (Fig. 1B), suggesting that interface
III by itself is not sufficient for the formation of the NEDD4
C2-GRB10 SH2 complex.
FIGURE 3. Crystal structure of the NEDD4 C2-GRB10 SH2 complex. A, ribbon diagram of the entire complex. There are three interaction interfaces be-
tween NEDD4 C2 and GRB10 SH2. Interface I includes the F -strand and the B helix of GRB10 SH2 and (primarily) the C and D -strands of NEDD4 C2. Inter-
face II includes the N termini of both NEDD4 C2 and GRB10 SH2. Interface III includes the N- and C-terminal regions of GRB10 SH2 and the C terminus of
NEDD4 C2. Both the Tyr(P)-binding pocket on GRB10 SH2 and the Ca2-binding site on NEDD4 C2 are far from these interfaces. The peptides without elec-
tron density are shown as dashed lines. B, the antiparallel -sheet formed between the C -strand of NEDD4 C2 and the F -strand of GRB10 SH2 in interface
I, containing four hydrogen bonds (shown as red lines with lengths in angstroms). Only the main chains are shown. C, the hydrophobic interactions be-
tween NEDD4 C2 and GRB10 SH2 in interface I. The two hydrogen bonds between Asn519 of GRB10 SH2 and Leu177 of NEDD4 C2 are shown as red lines with
lengths in angstroms. Only the side chains are shown (except for residue Leu177 of NEDD4 C2, whose main chain and side chain are shown).
Structure of the NEDD4 C2-GRB10 SH2 Complex
DECEMBER 31, 2010•VOLUME 285•NUMBER 53
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Structure of the NEDD4 C2-GRB10 SH2 Complex
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Because the main interactions between NEDD4 C2 and
GRB10 SH2 are hydrogen bonds and salt bridges, the complex
is expected to be unstable in high salt solution. In fact, as
shown above, the complex dissociates in a solution containing
more than 500 mM NaCl.
Phosphotyrosine-containing peptides/proteins bind to the
Tyr(P)-binding pocket on the surface of an SH2 domain (the
classical SH2 binding site) (Fig. 3A). On the surface of GRB10
SH2, this pocket is more than 15 Å away from the three
NEDD4 C2-GRB10 SH2 interfaces and has no overlap with
them (Fig. 3A); NEDD4 C2 binds to a nonclassical binding site
on GRB10 SH2, which explains why the interaction between
NEDD4 C2 and GRB10 SH2 is phosphorylation-independent.
Usually, a C2 domain binds Ca2 through its AB and EF
loops (Fig. 3A). This Ca2-binding site is more than 12 Å
away from the three NEDD4 C2-GRB10 SH2 interaction in-
terfaces, and no Ca2 was found in the interfaces. Hence, the
NEDD4 C2-GRB10 SH2 interaction is expected to be Ca2-
independent, as observed in binding studies (Fig. 1A).
Structure Comparison—Superposition of the GRB10 SH2
subunit of the NEDD4 C2-GRB10 SH2 complex with the free
GRB10 SH2 domain shows that binding of NEDD4 C2 has not
resulted in significant conformational change of GRB10 SH2
(r.m.s. deviation 0.37 Å) (Fig. 4A), except that the side
chain of residue Gln511 has moved from a position that would
clash with the main chain of residue Leu177 of NEDD4 C2 to a
location well clear of Leu177.
The structure of GRB10 SH2 is similar to those of GRB14
SH2 (r.m.s. deviation 0.92 Å) and GRB7 SH2 (r.m.s. deviation
0.95 Å). Superposition of GRB14 SH2 and GRB7 SH2 onto the
GRB10 SH2 subunit of the NEDD4 C2-GRB10 SH2 complex
shows that the main differences among these three structures
are located at the BC loop, the DE loop, and the N and the C
termini (Fig. 4A).
Almost all of the residues involved in interface I between
SH2 and NEDD4 C2 are identical or very similar among the
three species. One point of difference is the contact between
residues 494–496 (QTF) of GRB10 SH2 and residues 153–
155 (SGI) of NEDD4 C2. The sequence of the corresponding
SH2 residues is EMF in GRB14 and RLY in GRB7 (i.e. a
charged rather than merely polar residue is introduced in the
first position, and, in the case of GRB7, the large Tyr replaces
Phe in the third position). Additionally, the position of the
main chain is slightly shifted, relative to the central -sheet
contact, in the other two proteins versus GRB10.
In the other two interfaces, significant differences are ap-
parent among the superposed structures. In interface II, the
conformation of the N terminus of GRB7 SH2 is quite differ-
ent from that of GRB10 and would clash with NEDD4 C2
(Fig. 4B). GRB14 SH2 is less discrepant but does show a posi-
tional shift relative to GRB10 as well as a sequence difference
in one position, which could affect binding.
In interface III, both GRB7 and GRB14 exhibit significant
differences in their SH2 C-terminal conformations from
GRB10, such that the fit of the NEDD4 C2 Pro-rich cluster
against SH2 becomes very poor (Fig. 4, C and D). The possi-
bility exists that the GRB7 and/or GRB14 SH2 conformation
could be modified in the presence of NEDD4 C2 to create
more favorable interactions with C2, although in the case of
GRB14, a sequence change from Gly to His at position 438
(GRB10 numbering) would create a clash with Pro286 in
NEDD4 C2 even if the main chain conformation were the
same as in GRB10 (Fig. 4D). In any case, the binding studies
described above argue strongly against such a conformational
change.
NEDD4 C2 contains eight -strands and no -helices.
Comparison of the NEDD4 C2 subunit of the NEDD4 C2-
GRB10 SH2 complex with the free NEDD4 C2 domain (su-
perposition r.m.s. deviation 0.53 Å) shows that the main dif-
ferences between the two structures are located at the BC
loop and the N-terminal part, regions that interact with the
GRB10 SH2 subunit.
DISCUSSION
SH2 domains are well known as binding modules for phos-
photyrosine-containing peptides. The classical SH2/Tyr(P)-
containing peptide interactions play important roles in nu-
merous cell signaling pathways (38, 40). A typical SH2 domain
comprises a seven-stranded -sheet core flanked by two
-helices (41). SH2 domains are highly specialized for the rec-
ognition of Tyr(P) residues. On the classical substrate-binding
site of an SH2 domain, the target Tyr(P)-containing peptide is
usually bound in an extended conformation. The highly con-
served Tyr(P)-binding site contains an invariant arginine and
a second positively charged residue coordinating the phos-
phate moiety (42). Residues in the target peptide that are lo-
cated downstream of the Tyr(P) residue confer specificity on
the interaction (43).
It has been shown that the SH2 domains of GRB10 and
GRB14 can bind the activation loop of the kinase domains of
FIGURE 4. Superposition of free GRB10 SH2 (blue; Protein Data Bank code 1NRV), GRB7 SH2 (yellow; Protein Data Bank 2QMS), and GRB14 SH2 (red;
Protein Data Bank 2AUG) with the GRB10 SH2 subunit (cyan) of the NEDD4 C2-GRB10 SH2 complex. The NEDD4 C2 subunit of the NEDD4 C2-GRB10
SH2 complex is shown in magenta. A, the structure of GRB10 SH2 is similar to those of GRB14 SH2 and GRB7 SH2, with significant conformational differences
located at the BC loop, the DE loop, the N-terminal region, and the C-terminal region (marked with arrows). B, detailed structure of the N-terminal region
around residue Gln433. The side chain of residue Gln433 of GRB10 SH2 can form a hydrogen bond with the carbonyl group of residue Glu112 of NEDD4 C2
(shown with a red line), whereas the side chain of residue Gln437 of GRB14 SH2 is too close to the carbonyl group of residue Glu112 of NEDD4 C2 (shown with
a blue arrow, 1.49 Å). The N-terminal part of GRB7 SH2 (side chains of His426 and Arg427) clashes with the N-terminal part of NEDD4 C2 (residues Leu113 and
His114). C, detailed structure of the C-terminal region around residue Arg533. The side chain of residue Arg533 of GRB10 SH2 can form a hydrogen bond with
the carbonyl group of residue Pro284 of NEDD4 C2 (shown with a red line), whereas the side chain of residue Arg537 of GRB14 SH2 is too far away from the
carbonyl group of residue Pro284 of NEDD4 C2 (shown with a blue arrow, 3.91 Å) to form a hydrogen bond. Moreover, the side chain of residue Arg537 of
GRB14 SH2 is too close to residue Pro286 of NEDD4 C2 (1.5 Å). D, detailed structure of the N-terminal region around residue Arg431. There is a salt bridge
interaction between the side chain of residue Arg431 of GRB10 SH2 and the carboxyl group of residue Pro287 of NEDD4 C2 (shown with a red line, 3.22 Å),
whereas the side chain of residue Arg435 of GRB14 SH2 is too far away from the carboxyl group of residue Pro287 of NEDD4 C2 (shown with a blue arrow, 8.01
Å) to form a salt bridge interaction. Moreover, the side chain of residue Arg537 of GRB14 SH2 is too close to residue Pro286 of NEDD4 C2 (1.5Å). The side
chain of residue His442 of GRB14 SH2 clashes with residue Pro286 of NEDD4 C2, whereas the corresponding residues in GRB10 SH2 and GRB7 SH2 are gly-
cines that have no side chain.
Structure of the NEDD4 C2-GRB10 SH2 Complex
DECEMBER 31, 2010•VOLUME 285•NUMBER 53
JOURNAL OF BIOLOGICAL CHEMISTRY 42137
IR and IGF1R through classical Tyr(P)-SH2 interactions (44).
On the other hand, there was evidence showing that GRB10
could form a complex with the E3 ubiquitin ligase NEDD4
through the GRB10 SH2-NEDD4 C2 interaction. This inter-
action is phosphotyrosine-independent and Ca2-indepen-
dent (25). Furthermore, there was also evidence suggesting the
existence of a GRB10-NEDD4-IGF1R complex (23). The crys-
tal structure of the GRB10 SH2-NEDD4 C2 complex, re-
ported here, provides a structural basis for how the SH2 do-
main of GRB10 can bind the C2 domain of NEDD4 and the
kinase domain of IGF1R simultaneously. All of the three
NEDD4 C2 recognition sites on GRB10 SH2 are far away
(more than 15 Å) from, and do not overlap with, the classical
Tyr(P)-containing peptide binding pocket (Fig. 3A); binding
of the kinase domain of IGF1R at the Tyr(P)-binding pocket
(the classical site) does not interfere with binding of the C2
domain of NEDD4. In the NEDD4-GRB10-IGF1R complex,
GRB10 serves as a connector to form a bridge between
NEDD4 and IGF1R.
Although GRB10 can form a complex with the E3 ubiquitin
ligase NEDD4, GRB10 is not ubiquitinated by NEDD4 inside
the cell. However, IGF1R is ubiquitinated by NEDD4 inside
the cell, and binding of GRB10 to NEDD4 is critical for this
ubiquitination (25). This is explained by the predicted struc-
ture of the NEDD4-GRB10-IGF1R complex, in which GRB10
acts as an adaptor to bring NEDD4 close enough to IGF1R to
facilitate ubiquitination of IGF1R by NEDD4, through the
C2-SH2-kinase domain interaction (Fig. 5A).
There are a few other cases in which an SH2 domain binds
proteins using binding sites other than the classical Tyr(P)-
binding pocket (45, 46). For example, the Itk kinase domain
docking site on the PLC1 SH2C domain surface, which in-
cludes residues Glu709, Arg748, Met750, Lys751, and Arg753, is
far from and does not overlap with the classical Tyr(P)-bind-
ing pocket (47).
Some instances have been reported in which SH2 domains
use binding sites different from and not overlapping with the
classical Tyr(P)-binding pocket to colocalize a kinase and sub-
strate (45, 46). As shown in Fig. 5B, the SAP SH2 domain
binds simultaneously to the SH3 domain of Fyn kinase and to
the Tyr(P)-containing peptide of SLAM. This interaction re-
sults in the colocalization of Fyn with its SLAM substrate to
facilitate the phosphorylation of SLAM by Fyn (45). In the
complex, SLAM binds to the classical Tyr(P)-binding pocket
of SAP SH2, whereas Fyn SH3 binds SAP SH2 in a phosphoty-
rosine-independent manner, at a binding site outside of the
classical Tyr(P)-binding pocket, involving the DE loop and
part of the B helix (45). Another example is the interaction
between the fibroblast growth factor receptor kinase (FGFR1)
and the N-terminal SH2 domain (SH2N) of PLC1 (Fig. 5C)
(46). In this complex, there are two interaction sites between
PLC1 SH2N and the kinase domain of FGFR1; the Tyr(P)-
containing tail of the kinase domain binds the classical
Tyr(P)-binding pocket on SH2N, whereas a second interac-
tion site involves the BC and DE loops of SH2N.
Unlike the above cases in which the SH2 domain colocal-
izes a kinase and its substrate, in the GRB10-NEDD4-IGF1R
complex, GRB10 SH2 colocalizes a ubiquitin ligase (NEDD4)
and its substrate (IGF1R). This case provides further evidence
that SH2 domains have a diverse set of other interaction sur-
faces besides the classical Tyr(P)-binding pocket and that SH2
domains can colocalize an enzyme and its substrate to facili-
tate the reaction between them. Our work also supports the
FIGURE 5. Model for the interaction of NEDD4 with IGF1R through
GRB10 and examples of SH2 domains that use binding sites different
from and not overlapping with the classical Tyr(P)-binding pocket to
colocalize a kinase and substrate. A, the interaction between GRB10 BPS
(cyan) and the IGF1R kinase domain (green) is modeled based on the crystal
structure of the GRB14 BPS-IR kinase complex (25). The interaction between
GRB10 SH2 (cyan) and the IGF1R kinase domain is modeled based on the
crystal structure of the APS SH2-IR kinase complex (48). The dashed red oval
highlights the interaction between GRB10 SH2 and the activation loop of
the kinase domain. The missing linker between the BPS and the SH2 do-
main of GRB10 is shown as a dashed cyan line. B, structure of the SAP SH2-
Fyn SH3-SLAM complex (45). The two dashed red ovals highlight the Tyr(P)-
binding pocket and the interaction interface between SAP SH2 and Fyn
SH3, respectively. C, structure of the PLC1 SH2N-SH2C-FGFR1 kinase do-
main complex (46). The dashed blue oval highlights the Tyr(P)-binding
pocket. The dashed purple oval highlights the secondary interaction inter-
face between PLC1 SH2N and the FGFR1 kinase domain.
Structure of the NEDD4 C2-GRB10 SH2 Complex
42138
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 285•NUMBER 53•DECEMBER 31, 2010
conclusion that SH2 domains can participate in important
signaling interactions beyond the recognition of
phosphotyrosine.
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Structure of the NEDD4 C2-GRB10 SH2 Complex
DECEMBER 31, 2010•VOLUME 285•NUMBER 53
JOURNAL OF BIOLOGICAL CHEMISTRY 42139
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3M7G
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Structure of topoisomerase domain of topoisomerase V protein
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Structures of minimal catalytic fragments of topoisomerase V
reveals conformational changes relevant for DNA binding
Rakhi Rajan*, Bhupesh Taneja*,†, and Alfonso Mondragón*,‡
* Department of Biochemistry, Molecular Biology and Cell Biology, Northwestern University, 2205
Tech Dr, Evanston, IL 60208
Summary
Topoisomerase V is an archaeal type I topoisomerase that is unique among topoisomerases due to
presence of both topoisomerase and DNA repair activities in the same protein. It is organized as an
N-terminal topoisomerase domain followed by 24 tandem helix hairpin helix (HhH) motifs.
Structural studies have shown that the active site is buried by the (HhH) motifs. Here we show that
the N-terminal domain can relax DNA in the absence of any HhH motifs and that the HhH motifs
are required for stable protein-DNA complex formation. Crystal structures of various
topoisomerase V fragments show changes in the relative orientation of the domains mediated by a
long bent linker helix, and these movements are essential for the DNA to enter the active site.
Phosphate ions bound to the protein near the active site helped model DNA in the topoisomerase
domain and shows how topoisomerase V may interact with DNA.
Introduction
DNA topoisomerases are enzymes found in all forms of life (bacteria, eukarya, and archaea)
and they regulate the topological state of DNA inside the cell. They form a transient break in
a single or double stranded DNA and allow the passage of another single or double DNA
strand through the break, before resealing the break (Champoux, 2001) (Schoeffler and
Berger, 2008). As a result of this, topoisomerases can relax supercoiled DNA, help in the
segregation of DNA strands following replication, and lead to the formation and resolution
of knots and catenates (Gellert, 1981). Topoisomerases participate in many aspects of DNA
metabolism, such as replication, recombination, and transcription (Champoux, 2001). In
addition, they are targets of various anti-cancerous drugs and anti-bacterial agents (Maxwell,
1999; Pommier, 1998; Rothenberg, 1997; Wang et al., 1997).
DNA topoisomerases are broadly classified into two types, type I and type II enzymes. Type
I enzymes cleave a single strand of a DNA molecule and pass another single or double
stranded DNA through the break before resealing the opening. Type II enzymes cleave both
‡Corresponding author: Phone: 847-491-7726, Fax: 847-467-6489, a-mondragon@northwestern.edu.
†Present address: Institute of Genomics and Integrative Biology, CSIR, Delhi, India
Protein data bank accession codes
The final structure factors and coordinates of Topo-31, Topo-44 Form I, Form II, and Form III have been deposited in the Protein Data
Bank with accession codes 3M7G, 3M7D, 3M6K, and 3M6Z respectively.
Supplementary data
Supplementary data are available at Structure Journal Online.
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Author Manuscript
Structure. Author manuscript; available in PMC 2011 July 14.
Published in final edited form as:
Structure. 2010 July 14; 18(7): 829–838. doi:10.1016/j.str.2010.03.006.
NIH-PA Author Manuscript
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strands of a double stranded DNA in concert and pass another double stranded DNA through
the break. Type I enzymes use the torsional energy stored in the supercoiled DNA to drive
DNA relaxation and hence they do not require high energy cofactors, such as ATP, for their
activity (Baker et al., 2009) Type II enzymes, on the other hand, require ATP and Mg2+ for
their activity. Type I topoisomerases are further subdivided into three subtypes: IA, IB, and
IC (Forterre et al., 2007). Type IA and IB enzymes have been studied extensively (Baker et
al., 2009) and there is ample information available about their general mechanism of DNA
relaxation and the mode of DNA binding. Type IC, on the other hand, is a relatively new
subtype. Currently topoisomerase V is the only member of this family and it has been
identified only in the Methanopyrus genus. Previously, topoisomerase V had been
considered as a type IB enzyme based on its biochemical characteristics (Slesarev et al.,
1993), but the crystal structure of an N-terminal 61 kDa of topoisomerase V (Topo-61)
(Taneja et al., 2006) revealed a completely new fold without similarity to other
topoisomerases or any other known protein. Furthermore, the orientation of the putative
active site residues is also different from other type I topoisomerases, suggesting a different
mechanism of cleavage and religation of DNA. These observations, together with the lack of
sequence similarity, indicated that topoisomerase V defines a new subtype of type I enzymes
(Forterre, 2006).
Topoisomerase V was identified in Methanopyrus kandleri, an extremophile isolated from a
deep-water ‘black smoker’ chimney in the Gulf of California (Huber et al., 1989). The
enzyme is active at very high temperatures (122°C) and high salt concentrations (0.65 M
NaCl and 3.1 M potassium glutamate). The unusual characteristic of topoisomerase V is that
it has both topoisomerase and DNA repair activities in the same polypeptide (Belova et al.,
2001). Based on the sequence analysis of topoisomerase V, it has been predicted that the
protein contains 24 helix-hairpin-helix (HhH) DNA binding motifs arranged as 12 (HhH)2
domains around the N-terminal topoisomerase domain (Belova et al., 2002) (Figure 1A).
Some of these (HhH)2 domains are involved in the apurinic/apyrimidinic (AP) site-
processing activity, but the exact location of the repair active site is not known yet.
Topoisomerase V can relax both positively and negatively supercoiled DNA without the
need for metal cations or high energy cofactors. Single molecule experiments have shown
that topoisomerase V relaxes DNA by a constrained swiveling mechanism, relaxing around
12 turns of DNA per relaxation cycle (Taneja et al., 2007). Type IB enzymes, which also use
a constrained swiveling mechanism for DNA relaxation, relax around 19 turns of DNA per
relaxation cycle (Koster et al., 2005).
The structure of Topo-61 showed that the topoisomerase domain is mainly alpha helical and
that the first four (HhH)2 domains curl around the topoisomerase domain (Taneja et al.,
2006) (Figure 1B). The topoisomerase and (HhH)2 domains are joined by a long bent helix,
termed the “linker helix”. Three of the five putative active site residues are present in a
helix-turn-helix (HTH) domain and the other two are present in an intervening loop and a
helix. The active site residues are buried by the first (HhH)2 domain and it has been
suggested that large conformational changes will be needed for the DNA to access the active
site of topoisomerase V (Taneja et al., 2007). Here we present data that shows that the N-
terminal 31 kDa fragment of topoisomerase V (Topo-31) has topoisomerase activity,
consistent with previous predictions based on the structure. In addition, we show that the
Topo-44 fragment (N-terminal 44 kDa fragment of topoisomerase V) can form a stable
protein-DNA complex, emphasizing the need of the (HhH)2 domains for binding DNA. We
determined a crystal structure of (Topo-31) fragment, which has only the topoisomerase
domain, and three different crystal structures of the Topo-44 fragment, which includes the
topoisomerase domain and three tandem HhH motifs. In all structures, the topoisomerase
domain is very similar. In contrast, the structures of Topo-44 show conformational changes
in the linker helix resulting in variable orientations of the (HhH)2 domains when compared
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to the Topo-61 structure. Phosphate ions are present in the vicinity of the topoisomerase
active site in two of the Topo-44 structures. Some of the catalytic residues interact with the
phosphate ions and may mimic contacts with DNA. These observations suggest that the
movement of the (HhH)2 domains is mediated by the linker helix and helps expose the
topoisomerase active site to facilitate DNA binding. In addition, the location of the
phosphate ions suggests a possible path for the DNA and the way the active site residues
interact with it.
Results
The topoisomerase domain can relax DNA
DNA relaxation assays using different topoisomerase V fragments showed that the
topoisomerase domain alone is capable of relaxing DNA. Topoisomerase V fragments with
different numbers of (HhH)2 domains, Topo-31, Topo-44, and Topo-78, were studied using
relaxation assays. Topo-31 has no (HhH)2 domains, Topo-44 has one full and one partial
(HhH)2 domain, while Topo-78 has eight full (HhH)2 domains, including a putative DNA
repair domain. In addition to standard conditions, the effect of different pH conditions and
presence of magnesium ions were also tested. The experiments show that Topo-31 is
capable of relaxing DNA, despite the absence of the (HhH)2 domains (Figure 2B). A pH
profile analysis for the DNA relaxation assays showed that Topo-78 relaxes DNA over a
wider pH range (pH 5 to 9), while Topo-31 and Topo-44 relax DNA optimally at pH 5
(Figure 2A, 2B, 2C). In addition, magnesium is not required for the reaction, but stimulates
it at all pH values (Figure 2B, 2C). Topo-78 can relax DNA to the same extent with lower
amounts of protein (0.1 μg/reaction) compared to Topo-44 (~1.5 μg/reaction) and Topo-31
(~9 μg/reaction). This could be due to the enhanced DNA binding facilitated by the (HhH)2
domains. Together, these results suggest that, even though the (HhH)2 domains are
dispensable for topoisomerase activity, they enhance DNA relaxation activity. In addition,
the pH dependence of the DNA relaxation activity indicates that the reaction is likely to
involve side chains with ionizable groups in the low pH range, such as glutamates. Finally,
the magnesium independence of the reactions confirms that even the smallest fragments do
not require metals for activity, although magnesium has a stimulatory effect. This may be
due to favorable interactions of the cations with DNA.
The (HhH)2 domains enhance DNA binding affinity
EMSA experiments with different fragments of topoisomerase V and DNA showed that
(HhH)2 domains could help in the formation of a stable protein-DNA complex. Various
topoisomerase V fragments (Topo-31, Topo-44, and Topo-78) and single and double
stranded DNA were analyzed by EMSA experiments. Topo-44 and Topo-78 formed stable
complexes with a 39mer double stranded DNA (Figure 2D), while no DNA binding was
observed for the Topo-31 fragment (data not shown). These observations indicate that
(HhH)2 domains are necessary for a stable protein-DNA complex and that as few as one and
half (HhH)2 domains are enough for formation of a stable protein-DNA complex. EMSA
with single stranded DNA showed that Topo-31 and Topo-44 cannot bind to single stranded
DNA, while Topo-78 can bind to single stranded DNA (data not shown).
Overall Structures
The topoisomerase domain of topoisomerase V is a helical-rich compact domain that has no
structural similarity to any other known protein. The only recognizable structural element is
a HTH that contains some of the active site residues. Not surprisingly, the topoisomerase
domain of the four structures (Topo-31, Topo-44 (Forms I, II, and III)) superimpose very
well on each other and also to that from the Topo-61 structure. In the Topo-31 structure, two
surface loops, residues 39-49 and 120-124, adopt a different conformation compared to the
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Topo-61 and Topo-44 structures. These two loops are not always visible in the Topo-44
structures, suggesting that they are mobile regions. The r.m.s.d. for the superposition of the
topoisomerase core domain of all the new structures on to the Topo-61 structure range from
0.2 Å to 0.7 Å if the two mobile surface loops are not included (Figure 3A). In general, the
topoisomerase domain remains unchanged and is identical in all structures. The (HhH)2
domains also remain largely unchanged, with r.m.s.d. for the superposition of only the
(HhH)2 domains from the three Topo-44 crystal forms and equivalent domains in the
Topo-61 structure ranging from 0.31 Å to 0.56 Å.
The five crystallographically independent structures of Topo-44 (Form I, Form II A and B
monomers, and Form III A and B monomers) were compared with each other and to the two
crystallographically independent Topo-61 monomers to understand the conformational
changes in the protein. The r.m.s.d. for the superposition of all the Topo-44 structures
(residues 3-375) on to the Topo-61 fragment or on each other vary between 0.9 Å and 2.7 Å,
with the majority above 1.5 Å, showing that in general the structures have slightly different
conformations. As mentioned above, the different domains behave as rigid or almost rigid
subunits and the only change in the structure is the relative orientation between the
topoisomerase and the (HhH)2 domains. The change in orientation of the domains starts at
the linker helix (residues 269-295), which acts as a hinge region, and follows into the
(HhH)2 domains. At the start of the linker helix, the structures superimpose very well for all
five Topo-44 and two Topo-61 structures. In the middle of the linker helix there is a kink
after which the linker helix from all the structures shows different orientations (Figure 3B).
The flexibility of the linker helix is also evident by the fact that the linker helix in the B
subunit of Form III crystals appears in two alternate conformations. The change in the
relative orientation of the (HhH)2 and topoisomerase domains (Figure 3C and 3D), suggests
that these domains can adopt different orientations and these movements might be necessary
for the DNA to access the active site.
The topoisomerase domain has a positively charged groove adjacent to the active site
The structure of the Topo-31 as well as the structures of the Topo-44 fragment reveals the
presence of a positively charged groove in the protein that encompasses the active site
region (shown later in Figure 6C). This charged groove had been observed before in the
structure of the Topo-61 fragment, although several (HhH)2 motifs partially obstruct it
(Taneja et al., 2006). The structure of the Topo-31 confirms the presence of the groove even
in the absence of the (HhH)2 motifs. The groove is long and can be deep in some areas. It
includes regions of the HTH motifs and extends all the way to the linker helix. All the
residues forming the active site pentad point towards the groove. The active site tyrosine,
Tyr226, is found near one of the ends of the groove, a region where it widens. The positively
charged character of the groove and its presence by the active site strongly suggest that it
may be involved in DNA binding.
Phosphate ions bind in the groove near the topoisomerase active site
An interesting observation stemming from the Form II and Form III Topo-44 structures is
the presence of phosphate ions near the positively charged DNA binding groove. All three
Topo-44 crystal forms were crystallized in the presence of phosphate-citrate buffer, but only
Form II and Form III structures showed phosphate ions bound to the protein, which were
assigned based on electron density consistent with a tetrahedral phosphate ion (Figure 4A).
Form II and Form III crystals include 1–1.2 M guanidium hydrochloride in the
crystallization solution. The high resolution Form III structure shows clear density for three
guanidium ions bound to the protein, two very well ordered and one with weak density. The
presence of guanidium hydrochloride in the crystals appears to trigger a conformational
change allowing the binding of phosphate ions to the protein. It is interesting to note that
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Form I crystals did not show any bound phosphate albeit its presence in the crystallization
condition. This could be due to the absence of guanidium hydrochloride to trigger the
binding of phosphate ions as observed in Form II and Form III structures. There are three
phosphate ions in the B subunit and none in the A subunit of the Form II Topo-44 structure.
Two of the phosphates are in the topoisomerase active site and one of them forms close
contacts with the putative active site residues in the topoisomerase domain (Figure 4B).
Form III crystal has seven phosphate ions, three in each subunit and one between both the
subunits. In the Form III structure, the phosphate ion near the active site Tyr226 is absent,
but it shows several new locations for phosphate ions, especially in the positively charged
groove containing the topoisomerase active site (Figure 5A). An overlay of the A and B
subunits of the Topo-44 Form III structure with the B subunit of Topo-44 Form II structure
shows eight unique phosphate ions (Figure 5A). It clearly shows that there are more
phosphate ions bound in the positively charged groove compared to other regions of the
protein.
Taking into account all structures, there are five unique phosphate ion binding sites in the
putative DNA binding groove and an additional one near its end and close to the start of the
linker helix. Several pairs of phosphates in the groove are separated by a distance of around
7 Å (Figure 5B), which would be consistent with the phosphate-phosphate distance in
adjacent nucleotides in a DNA double helix. One of the phosphates (P1) is found near the
active site tyrosine and is coordinated by Tyr226 and Arg131, two residues that have been
implicated in cleavage and religation of the DNA (Taneja et al., 2006), and by Glu215,
whose charge may be important for interactions with DNA (R.R. and A.M., unpublished
observations). The side chains of the tyrosine and the glutamate residues are in contact with
Arg144 and His200, the other putative active site residues, and these interactions may help
to orient them for the catalytic reaction. Adjacent to P1, there is a second phosphate (P2) at a
distance of 7.5 Å which is trapped between the topoisomerase domain and an HhH motif. P2
is coordinated by Arg131, an active site residue, in addition to Arg108 from the
topoisomerase domain and Arg293 and serines 322 and 324 from the second HhH motif
(Figure 6C). Three more phosphates are found in the groove (P3, P4, and P5) coordinated
mainly by positively charged residues, such as Arg37, Lys47, Arg108, Lys134, and Arg135
from the topoisomerase domain and also residues from the linker helix such as Tyr289 and
Arg293. In general, some of the side chains can contact more than one phosphate. The
distance between P3 and P4 and P4 and P5 is 6.8 Å and 6.5 Å respectively. A final
phosphate (P6) is located at the start of the linker helix and on the edge of the groove
(Figure 5A).
Discussion
Topoisomerase V is active at very high temperatures (122°C) and high salt concentrations.
DNA relaxation assays with various topoisomerase V fragments (Topo-44 and Topo-61)
show that a temperature above 60° C is required for optimal activity, although longer
fragments of topoisomerase V can relax DNA at lower temperatures (Taneja et al., 2007).
Topo-44 was first identified by limited proteolytic digestion of the full length topoisomerase
V protein (Belova et al., 2002) at 80°C. In contrast, Topo-61 is the shortest fragment
showing topoisomerase activity when the proteolytic reaction is performed at 37°C (Belova
et al., 2002). The N-terminal Topo-31 fragment, which contains neither HhH motifs nor the
linker helix, was identified as the smallest region spanning the topoisomerase domain from
the crystal structure of Topo-61 fragment (Taneja et al., 2006) and it was suggested that it
could represent the minimal domain capable of relaxing DNA. Relaxation experiments with
this minimal domain show that this is indeed the case, although the activity is not as robust
as with longer fragments. As expected, Topo-31 does not require magnesium for activity,
but magnesium enhances it, as is the case for type IB topoisomerases, which also uses a
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swiveling mechanism for DNA relaxation (Stewart et al., 1996). The optimal pH for activity
for the Topo-31 and the Topo-44 fragments is around 5. This pH dependence is not observed
for the Topo-78 fragment. The DNA relaxation by shorter fragments of topoisomerase V at
pH 5 could point to the involvement of some ionizable side chains in the relaxation activity.
It could also be simply due to the effects of various side chains on DNA binding. Further
experiments with different active site mutations in both longer and shorter fragments of
topoisomerase V will be required to probe the pH dependence of the relaxation reaction by
shorter topoisomerase V fragments.
Gel shift experiments show that Topo-44 and also longer fragments (Topo-78) can bind
double stranded DNA. Surprisingly, Topo-31 does not show DNA binding activity in these
assays even though it is still capable of relaxing DNA. It appears that the presence of the
(HhH)2 domains stabilizes the DNA/protein complex. One possibility is that the (HhH)2
domains could play a similar role to the cap domain present in type IB enzymes, which helps
to encircle the DNA during the swiveling reaction (Redinbo et al., 1998). In addition, both
short fragments of topoisomerase V do not bind single stranded DNA, whereas Topo-78 can
form a stable complex with single stranded DNA (data not shown). (HhH)2 domains binding
to single stranded DNA has been observed before. For instance, the N-terminal 8 kDa of
mammalian polymerase β, which contains a single HhH motif, binds to single stranded DNA
through both helices (Kumar et al., 1990; Liu et al., 1994). The exact mode of single
stranded DNA binding by Topo-78 or the possible role in relaxation or repair activities is not
yet clear.
The structure of Topo-61 showed that the topoisomerase active site of topoisomerase V is
buried by one of the (HhH)2 domains suggesting that conformational changes are essential
for the protein to bind DNA. The present structures of Topo-44 reinforce this observation
and show that the (HhH)2 domains can change their position relative to the topoisomerase
domain and that this change is mediated by the movement of the linker helix. The (HhH)2
domains act as rigid individual units, as evidenced by the fact that in different structures
they show the same structure and relative orientation of the two HhH motifs. The
topoisomerase domain also appears to be rigid showing the same structure even in the total
absence of the rest of the protein. The linker helix (residues 269-295), which is a long bent
helix, serves as a hinge for the movement of the (HhH)2 domains away from the rigid
topoisomerase domain, possibly by responding to interactions with double stranded DNA.
This movement has to be quite large. The Topo-44 structures in the absence of DNA capture
the regions that move, but do not show the full extent of the movement or indicate the way
the HhH motifs interact with DNA.
As mentioned before, topoisomerase V binds double stranded DNA and has a groove wide
enough to accommodate double strand DNA (Figure 6C). The presence of an HTH domain
normally associated with DNA binding, the positively charged nature of it, and several
phosphates bound along it suggest that this groove could be involved in DNA binding. In
addition, the active site is found in this groove and some residues form part of the HTH
domain. Previously, DNA was modeled bound to the topoisomerase domain (Taneja et al.,
2006) based on the structures of HTH domains in complex with DNA but there was no
evidence to support it. Using the phosphates present in the groove in the current structures, it
is possible to refine this model. A superposition of the B subunit of Form II and the A and B
subunits of Form III Topo-44 structures shows five different phosphate ions in the positively
charged groove which are separated by a distance of around 7 Å, consistent with the
distance of consecutive phosphates in B DNA of ~6.4 Å. A sixth phosphate ion is found
outside the groove near the linker helix. A double stranded DNA molecule was modeled into
the groove based on the positions of the phosphate ions (Figure 6). Only five out of the six
phosphates could be placed on the DNA molecule, as one of them was inconsistent with a
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double stranded molecule. Phosphate ions P3, P4, and P5 would correspond to three
adjacent phosphates in one DNA strand, while P1, located near the active site, would belong
to the opposite strand. A final phosphate (P6) is away from the groove and near the linker
helix (Figure 6A, 6B). The fit to the four inner phosphates is excellent and the DNA can be
accommodated in the groove of the Topo-31 structure without the need for any major
rearrangements of the protein backbone. The fifth phosphate (P6) does not fit as well and a
better fit would require movement of either the protein or the DNA, but the change would be
relatively modest. Several side chains would need to move, but these changes would also be
minor. The major change needed to accommodate the DNA in the structures with the
(HhH)2 domains present is the movement of the (HhH)2 domains away from the
topoisomerase domain (Figure 6B). The movement of (HhH)2 domains should be feasible as
is evident from the Topo-44 structures showing different orientations of the (HhH)2
domains. The location of the (HhH)2 domains after DNA binding is not evident, but one
possibility is that they would help enclose the DNA to form a clamp around it, similar to the
arrangement in type IB enzymes.
In the model of the topoisomerase domain in complex with DNA, the active site residues are
in close contact with the backbone of DNA. The catalytic Tyr226 is pointing towards the
phosphate of the DNA backbone; Arg131 and Arg144 are positioned to stabilize the protein-
DNA covalent complex. Surprisingly Glu215 also appears to interact directly with the DNA
phosphate backbone. The other active site residues like His200 and Lys 218 are also near the
DNA. The active site is located near the end of the groove, where it widens. At this end, the
DNA fits loosely in the groove, which is spacious to accommodate the movement of the
strands. The ‘constrained swiveling’ mechanism employed by type IB and IC enzymes
necessitates rotation of one strand about the other after forming the covalent protein-DNA
intermediate. The position of the active site at the wider end of the putative DNA binding
groove would facilitate the rotation of the DNA strand at this end, while holding the rest of
the DNA in place through extensive interactions along the groove.
Even though type IB and IC enzymes have a similar overall mechanism of action, the
structures of fragments of topoisomerase V suggest many differences. Type IB enzymes
have two domains which come together to form a C-shaped clamp around the DNA (Perry et
al., 2006; Redinbo et al., 1998; Stewart et al., 1998) The protein has an open stage where
these domains are separate and this helps in the entry and release of the DNA from the
protein active site. A wide DNA binding cavity is not observed in the topoisomerase V
structures. Instead, the structures show a positively charged groove which is always present
in the protein and does not require domain rearrangements to form. DNA can access this
groove after a conformational change involving the movement of the (HhH)2 domains
exposing the active site. The (HhH)2 domains could help enclose DNA during the swiveling
of the DNA, forming a similar enclosure to the one observed for type IB enzymes. It is not
known whether all HhH motifs contact DNA simultaneously, but this appears unlikely
without a major rearrangement of the motifs. It is likely that only some of the HhH motifs
contact DNA at any given time or that some of the motifs do not have the capacity to bind
DNA. Finally, similar to type IB enzymes (Cheng and Shuman, 1998), the putative domain
enclosing the DNA is dispensable for activity, although it enhances the relaxation activity
markedly. Thus, it is likely that type IB and IC enzymes have several overall similarities in
the way that they interact with DNA, but the atomic details are markedly different.
There are still many details of the atomic mechanism of type IC topoisomerases that need to
be understood. The present functional and structural studies provide new information about
topoisomerase V including the observations that the Topo-31 is the minimal fragment
capable of DNA relaxation, the (HhH)2 domains enhance binding of the protein to DNA, the
changes in relative orientation of the domains is mediated by the linker helix, and several
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phosphate ions bind in a positively charged groove. Furthermore, the position of the
phosphate ions in the groove helped in the placement of DNA in the topoisomerase domain
and this provides an initial model of how topoisomerase V interacts with DNA. Thus the
present study helps to establish the role of different domains more clearly, to illustrate a
mechanism to drive the conformational changes needed for activity, and to suggest a
possible manner of binding DNA. Additional work on structures of protein/DNA complexes
and intermediates in the swiveling reaction are needed to understand the way this new type
of topoisomerases interacts with DNA to perform a complex reaction.
Experimental Procedures
Protein purification
The N-terminal 31 kDa (Topo-31: residues 1-269), and 44 kDa (Topo-44: residues 1 to 380)
fragments of topoisomerase V protein were cloned into the pET15b plasmid and
transformed into Escherichia coli BL21 Rosetta (DE3) cells. The N-terminal 78 kDa
(Topo-78: residues 1 to 685) fragment of topoisomerase V protein was cloned into the
pET14b plasmid (Belova et al., 2002) and transformed into Escherichia coli BL21(DE3)
cells. For protein production, cells were grown at 37° C in LB medium containing 100 μg/ml
ampicillin and 100 μg/ml chloramphenicol for Rosetta cells and LB medium with 100 μg/ml
ampicillin for BL21(DE3) cells to an optical density (OD600) of 0.6. The cells were then
cooled down on ice, followed by the addition of isopropyl β-D-1-thiogalactopyranoside
(IPTG) to a final 0.5 mM concentration, and grown overnight at 16° C. Cells were harvested
and resuspended in 50 mM Tris pH 8, 500 mM NaCl, 0.5 mM EDTA, 1 mM DTT, flash
frozen in liquid nitrogen and stored at −80° C. After thawing the pellet, pepstatin (1μg/ml),
benzamidine (1mM), PMSF (1mM), and Brij 58 (0.1%) were added to the cells and the
protein was purified as described earlier (Taneja et al., 2006) The protein was further
purified by anion exchange and gel filtration chromatography. Pure protein was
concentrated and stored in 50 mM Tris pH 8, 250 mM NaCl, and 1 mM DTT. The seleno-
methionine substituted Topo-44 was prepared from cells grown in a minimal medium
supplemented with nutrients and salts (Doublie, 1997); protein purification followed the
same procedure as for the native protein except that 5mM DTT was used in all the
purification steps and for storage.
Relaxation assays
Relaxation assays with the different topoisomerase V fragments were carried out at pH
values ranging from 4 to 10. The pH of the buffers was adjusted at 65 °C to account for the
change in pH at higher temperature. The different buffers used were: sodium acetate for pH
4 and 5, MES for pH 6, HEPES for pH 7, TRIS for pH 8, CHES for pH 9, and CAPS for pH
10. Topoisomerase activity assays were performed by incubating varying amounts of protein
(Topo-31, Topo-44 or Topo-78) with 0.2 μg negatively supercoiled pUC19 DNA in 50 mM
of the required buffer, 30 mM NaCl, 0.2 mM or 5 mM EDTA or 1 mM MgCl2. The
reactions were carried out at 65 °C for 15 min and terminated by cooling and addition of
SDS to a final 1% concentration. The products were resolved on a 1% agarose gel and
visualized by ethidium bromide staining.
Electrophoretic Mobility Shift Assay
For Electrophoretic Mobility Shift Assay (EMSA), 4 μM of a 39mer double stranded DNA
oligonucleotide (5′ GCGACGCGAGGCTGGATGGCCTTCCCCATTATGATTCTT3′) was
incubated with different concentrations of topoisomerase V fragments in 50 mM sodium
acetate pH 5, 30 mM NaCl, 1 mM MgCl2 at 65 °C for 30 minutes. Glycerol was added to
the reaction mixture to a final concentration of 8% and the products were separated on a 4 %
acrylamide native gel. The gel was stained with ethidium bromide to detect the DNA. When
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a stable protein-DNA complex was formed, there was an upward shift in the band indicating
a higher molecular weight complex.
Crystallization
Topo-31 crystals were grown using the sitting drop vapor diffusion method equilibrated
against, 23% PEG 6000, 0.1 M Na citrate pH 5.5, at 22°C. For data collection, the Topo-31
crystals were cryo-protected by adding glycerol to the mother liquor to a final 20%
concentration. Topo-44 was crystallized by the hanging drop vapor diffusion method under
three different crystallization conditions (Forms I, II, and III). Crystal Form I grew under 0.1
M phosphate citrate pH 5, 0.2 M NaCl, 15% PEG 3350 and 8% dioxane. The crystals were
cryo-protected by increasing the PEG concentration to 30%. Form II crystals grew under 0.1
M phosphate citrate pH 5, 0.2 M NaCl, 16% PEG 8000 and 1M guanidium hydrochloride.
For cryo-protection, they were transferred to a solution with 1.5X reservoir solution and
20% 2,3 butanediol or 20% DMSO for 10 seconds and immediately flash frozen under
liquid nitrogen. Form III crystals grew under 0.1 M phosphate citrate pH 5.5, 0.15 M sodium
sulfate, 0.01 M MgCl2, 1 M guanidium hydrochloride, and 28 % PEG 3350. The crystals
were grown at 30°C and were cryo-protected by increasing the PEG concentration to 40%.
Further details of crystallization are presented in the Supplementary Information.
Data collection and structure determination
Diffraction data were collected at the Dupont Northwestern Dow and Life Science
Collaborative Access Team stations (DND and LS CAT) at the Advanced Photon Source in
Argonne National Laboratory. Data collection and refinement statistics are shown in Table I.
All data were processed and integrated using XDS (Kabsch, 1993) and scaled with SCALA
(Collaborative-Computational-Project-4, 1994). Data on the Topo-31 crystals were collected
to 2.4 Å resolution. The structure was solved by Molecular Replacement (McCoy et al.,
2007) using the topoisomerase domain from the Topo-61 structure (residues 1-266) (Taneja
et al., 2006) as the search model. It was refined with refmac5 (Murshudov et al., 1997) and
Phenix (Afonine et al., 2005) to a final Rwork of 20.0 % and Rfree of 24.8 %. Topo-44 Form I
crystals diffract to 1.8 Å. The structure of Form I crystals was solved by Molecular
Replacement (McCoy et al., 2007) using the topoisomerase domain from the Topo-61
structure as the search model. Model rebuilding was performed using coot (Emsley and
Cowtan, 2004), and refinement using refmac5 (Murshudov et al., 1997). The final Rwork and
Rfree are 17.5 % and 22.0 % respectively.
For Topo-44 Form II and Form III crystals, seleno-methionine derivatized crystals were
used for single-wavelength anomalous dispersion (SAD) phasing. AutoSharp (Vonrhein et
al., 2007) was used for locating the selenium atoms; model building was done using coot
(Emsley and Cowtan, 2004), and refinement was carried out using refmac5 (Murshudov et
al., 1997) Three phosphate ions were noticed in the Form II structure; two of which present
in the topoisomerase active site and are separated by a distance of ~7.5 Å. The structure was
refined to a final Rwork of 24.1 % and Rfree of 28.9 %. Topo-44 Form III crystals diffracted
to 1.4 Å. The final Rwork and Rfree are 16.5 % and 18.4%, respectively. An interesting
observation is the presence of both phosphate and guanidium ions in the Form III Topo-44
structure. The linker helix and part of the first HhH motif of the B monomer show alternate
conformations and were built as two separate chains with occupancy of 0.5 each. Further
details on data collection and structure determination are given in the Supplementary
Information.
All figures were made with Pymol (DeLano, 2002) and the electrostatic surfaces were
calculated with APBS (Baker et al., 2001).
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Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We acknowledge staff and instrumentation support from the Keck Biophysics Facility and the Center for Structural
Biology at Northwestern University, and DND and LS-CAT at the Advanced Photon Source (APS) at Argonne
National Laboratory. Support from the R.H. Lurie Comprehensive Cancer Center of Northwestern University to the
Structural Biology Facility is also acknowledged. DND-CAT is supported by Dupont, DOW and the NSF. LS-CAT
was supported by the Michigan Economic Development Corporation and the Michigan Technology Tri-Corridor.
Use of the APS is supported by the Department of Energy (DOE). Research was supported by NIH grant GM51350
(to AM).
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Figure 1. Organization of topoisomerase V
Topoisomerase V is a multi-domain protein consisting of 24 helix-hairpin-helix (HhH) DNA
binding motifs arranged as 12 (HhH)2 domains following the N-terminal topoisomerase
domain. A) Schematic diagram of various topoisomerase V fragments. The topoisomerase
domain is shown in red, the (HhH)2 domains are shown in alternating colors of cyan and
yellow. The (HhH)2 domains with repair activity are shown in green. All fragments shown
have topoisomerase activity, but only the full length protein and the Topo78 fragment have
repair activity. B) Crystal structure of Topo-61 fragment (Taneja et al., 2006). The coloring
scheme is the same as in Figure 1A, except that the linker helix is shown in grey.
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Figure 2. DNA relaxation activity and EMSA for Topo-31, Topo-44 and Topo-78 fragments of
topoisomerase V
A) pH profile of the DNA relaxation activity of Topo-78 and Topo-44 fragments. 0.2 μg of
pUC19 DNA were incubated with 0.1 μg of Topo-78 or 1.5 μg of Topo-44 proteins at 65°C
for 15 minutes for the relaxation reaction. The reaction buffer contained 50 mM of the
appropriate buffer, 30 mM NaCl and 0.2 mM EDTA. Topo-78 relaxes DNA at a wider pH
range (5 to 9) than Topo-44, which relaxes DNA efficiently only at pH 5. DNA relaxation
activity of Topo-31 (B) and Topo-44 (C) fragments in the absence and presence of MgCl2.
0.2 μg of pUC19 DNA were incubated with 9 μg of Topo-31 or 1.5 μg of Topo-44 proteins
at 65°C for 15 minutes for the relaxation reaction. The reaction buffer contained 50 mM of
the appropriate buffer, 30 mM NaCl and 5 mM EDTA or 1 mM MgCl2. Both Topo-31 and
Topo-44 fragments can relax DNA in the absence of MgCl2, but MgCl2 enhances the DNA
relaxation activity of the topoisomerase V fragments. The black triangle in panels A, B and
C represents increasing pH from 4 to 10 by one pH unit. D) EMSA of Topo-44 and Topo-78
fragments with a 39mer double stranded DNA. Both Topo-44 and Topo-78 form stable
complexes with DNA, although Topo-78 seems to saturate DNA binding while Topo-44
does not. In addition, Topo-44 shows some cleavage of the DNA (bottom free DNA band),
while the cleavage is not apparent in Topo-78. The numbers at the bottom represent the
molar ratio of protein to DNA.
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Figure 3. Structure of Topo-44 fragments
A) Overlay of Form I (green), Form II (B subunit: blue), and Form III (B subunit: magenta)
structures onto the Topo-61 structure (B subunit: orange). The topoisomerase domains
superimpose very well for all the structures, while the linker helix and (HhH)2 domains
show differences in orientation. B) Overlay of the linker helices of Form I, II, and III
structures with that of Topo-61. The color scheme is same for all the figures unless
mentioned otherwise. Note that the linker helices have the same orientation at the start and
they change as they move further down the helix. C) Superposition of Form I, II, and III
Topo-44 structures with that of Topo-61. Only the (HhH)2 domains are colored while the
remaining parts are shown in gray for clarity. The active site residues are shown as orange
sticks. Note that the (HhH)2 domains adopt different orientations in all the structures. D)
Orientation of the (HhH)2 domains of Form I, II and Topo-61 structures. In Form I and II
structures, the (HhH)2 domains are moved away from the topoisomerase domain. For clarity,
the (HhH)2 domains of Form III are not shown. In panels C and D, the topoisomerase
domains were superposed to emphasize the different orientation of the other domains.
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Figure 4. Phosphate ions present near the active site of the Topo-44 structure
A) Stereo view of a Form III difference electron density map calculated with a model not
including the phosphates. The electron density is contoured at 3.7σ and shows the
tetrahedral shape of the phosphate ions. The active site residues are shown in stick. B)
Stereo view of the interaction of the phosphate ions with the putative active site residues.
The B subunit of Form II structure was superimposed onto the B subunit of Form III
structure and the phosphates ions from both structures are shown together with the Form II
B subunit protein backbone. The interactions made by the phosphate ion with the active site
residues and the corresponding distances in Å are represented as black dotted lines.
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Figure 5. Representation of the unique phosphate ions from Form II and Form III Topo-44
structures
A) An overlay of the A (magenta) and B (brown) subunits of the Form III structure and B
(blue) subunit of Form II Topo-44 structures. The positions of eight unique phosphate ions
(orange spheres) are shown. Note that most phosphate ions are found along the DNA
binding groove of the topoisomerase domain. B) The phosphate ions in the DNA binding
groove are separated by distances of around 7 Å. The protein backbone is that of the B
subunit of Form III structure. The active site residues are represented as sticks and distances
in Å between adjacent phosphate ions are shown as black dotted lines.
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Figure 6. Model showing DNA bound to the topoisomerase domain
A) Model of a 17-mer double stranded DNA bound to the Topo-31 structure (teal). The
DNA is represented as green sticks, where as phosphate ions are represented as orange
sticks. DNA binds along the DNA binding groove and five of the eight phosphate ions noted
in the Topo-44 structures coincide with the DNA backbone. B) Model of Topo-44 (Form II,
B subunit: blue) binding to 17-mer double stranded DNA. Note that the linker helix and the
(HhH)2 domains interfere with DNA binding to the topoisomerase domain and are likely to
move away to allow binding. C) Electrostatic surface representation of the Topo-31
structure. The positively charged DNA binding groove is clearly visible and the phosphate
ions are bound in this groove. The orientation corresponds to a 90° rotation of the one shown
in Figure 6A in the direction of the arrow. Note that the DNA binding groove goes from one
end of the molecule to the other and it is narrower at one end (start of the linker helix) and
wider at the other end. The putative active site residues (green sticks) are located at the
wider end of the groove. Other residues lining the groove and interacting with the phosphate
ions are shown as cyan sticks. D) Electrostatic surface representation of Topo-31 with
phosphate ions (orange) and DNA (green). Three phosphate ions (P3, P4, and P5) coincide
with the phosphates of one of the DNA strands, where as P1 coincides with a phosphate of
the opposite DNA strand. The model shows that the DNA binding groove of topoisomerase
V is wide enough to bind DNA and that the movement of linker helix and (HhH)2 domains
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are required to accommodate the DNA. The electrostatic potential was calculated with a
dielectric constant of 80 for solvent and 2 for protein. The surface is colored with a blue to
red gradient from +10 to −10 KbT/ec.
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Table 1
Data collection and refinement statistics
Topo-31
Topo-44 Form I
Topo-44 Form II
Topo-44 Form III
Data Collection
Space group
C2221
C121
P41212
P212121
Cell dimensions
a=106.7 Å, b=119.4
Å, c=63.7 Å
a=104.2 Å, b=47.7 Å,
c=81.2 Å (β=112.48)
a=b=70.1 Å, c=349.6 Å
a=63.6 Å, b=80.1 Å,
c=137.2 Å
Resolution (Å)a
79.56 – 2.4 (2.53 –
2.4)
75.05 – 1.82 (1.91 –
1.82)
29.5- 2.6 (2.72-2.6)
28.9-1.4 (1.46-1.4)
Number of observed
reflections
78,729 (11,538
134,411 (13,220)
227,408 (19,917)
1,157,917 (126,319)
Number of unique reflections
16,259 (2,346)
32,998 (4,301)
28,151 (3,331)
136,662 (15,986)
Completeness (%)
99.8 (99.8)
98.3 (88.6)
99.9 (100.0)
98.8 (95.5)
Multiplicity
4.8 (4.9)
4.1 (3.1)
8.1 (6.0)
8.5 (7.9)
Rmerge (%)b
4.7 (71.1)
4.0 (16.3)
7.4 (52.2)
4.5 (37.9)
Rmeas (%)c
5.3 (79.6)
4.6 (19.4)
7.9 (57.2)
4.8 (40.5)
≪I>/σ(<I>)>d
20.5 (2.5)
23.0 (6.8)
19 (3.2)
27.5 (5.3)
Refinement
Resolution (Å)
79.56 - 2.4 (2.46 -
2.4)
28.06 -1.82 (1.87 –
1.82)
29.14 – 2.6 (2.67 – 2.6)
28.9 - 1.4 (1.44 - 1.4)
Number of reflections
working/test
15,419/821
31,317/1,673
26,710/1,438
129,802/6,859
Rwork (%)e
20.0(24.3)
17.5 (17.9)
24.1(36.6)
16.5 (19.3)
Rfree(%)f
24.8 (31.1)
22.0 (24.8)
28.9 (45.1)
18.4 (22.1)
Protein residues/atomsg
269/2,203
376/3212
727/5,970
738/7,511
Atoms in alternate
conformations
0
258 (20 protein
residues)
8 (1 protein residue)
2846 (157 protein
residues)
Water molecules
29
238
30
573
Other atoms
-
-
3 PO4
7 PO4, 3 Gmh, 3 Mg++, 2
Cl−
B-factor (Å2)
Protein atoms (chain)
68.4
22.8
A:53.8; B:58.2
A:13.4; B:14.9
Water molecules
59.1
29.3
40.0
23.7
r.m.s. deviations
bond lengths (Å)
0.015
0.006
0.01
0.009
bond angles (°)
1.42
0.920
1.2
1.2
Ramachandran ploti
Favored regions (%)
94.3
98.9
96.2
98.5
Outliers (%)
0.0
0.0
0.3
0
aNumbers in parenthesis correspond to highest resolution shell.
bRmerge= Σ|I − <I>|/ΣI, where I is the observed intensity and <I> the average intensity obtained from multiple measurements.
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cRmeas as described in Diederichs and Karplus (Diederichs and Karplus, 1997).
d≪I>/σ(<I>)> = Mean Ih over the standard deviation of the mean Ih averaged over all reflections in a resolution shell.
eRwork= Σ ||Fo| − |Fc||/Σ|Fo|, where |Fo| is the observed structure factor amplitude and |Fc| the calculated structure factor amplitude.
fRfree: Rfactor based on 5% of the data excluded from refinement.
gTotal number of protein atoms, including those in alternate conformations.
hGm: guanidinum ion.
iAs reported by Molprobity (Davis et al., 2004).
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|
3M7H
|
Crystal structure of the bacteriocin LLPA from Pseudomonas sp.
|
Structural Determinants for Activity and Specificity of
the Bacterial Toxin LlpA
Maarten G. K. Ghequire1., Abel Garcia-Pino2,3., Eline K. M. Lebbe1¤, Stijn Spaepen1, Remy Loris"2,3*,
Rene´ De Mot"1*
1 Centre of Microbial and Plant Genetics, University of Leuven, Heverlee-Leuven, Belgium, 2 Molecular Recognition Unit, Department of Structural Biology, Vlaams
Instituut voor Biotechnologie, Brussel, Belgium, 3 Structural Biology Brussels, Department of Biotechnology (DBIT), Vrije Universiteit Brussel, Brussel, Belgium
Abstract
Lectin-like bacteriotoxic proteins, identified in several plant-associated bacteria, are able to selectively kill closely related
species, including several phytopathogens, such as Pseudomonas syringae and Xanthomonas species, but so far their mode
of action remains unrevealed. The crystal structure of LlpABW, the prototype lectin-like bacteriocin from Pseudomonas
putida, reveals an architecture of two monocot mannose-binding lectin (MMBL) domains and a C-terminal b-hairpin
extension. The C-terminal MMBL domain (C-domain) adopts a fold very similar to MMBL domains from plant lectins and
contains a binding site for mannose and oligomannosides. Mutational analysis indicates that an intact sugar-binding pocket
in this domain is crucial for bactericidal activity. The N-terminal MMBL domain (N-domain) adopts the same fold but is
structurally more divergent and lacks a functional mannose-binding site. Differential activity of engineered N/C-domain
chimers derived from two LlpA homologues with different killing spectra, disclosed that the N-domain determines target
specificity. Apparently this bacteriocin is assembled from two structurally similar domains that evolved separately towards
dedicated functions in target recognition and bacteriotoxicity.
Citation: Ghequire MGK, Garcia-Pino A, Lebbe EKM, Spaepen S, Loris R, et al. (2013) Structural Determinants for Activity and Specificity of the Bacterial Toxin
LlpA. PLoS Pathog 9(2): e1003199. doi:10.1371/journal.ppat.1003199
Editor: Ambrose Cheung, Geisel School of Medicine at Dartmouth, United States of America
Received August 22, 2012; Accepted January 3, 2013; Published February 28, 2013
Copyright: 2013 Ghequire et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was financially supported by FWO Vlaanderen (Research project G.0393.09), by The Onderzoeksraad of the VUB, by VIB and by the Hercules
Foundation. The authors acknowledge support of the European Community - Research Infrastructure Action under the FP6 ‘‘Structuring the European Research
Area Program’’, contract number: RII3-CT-2004-506008. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of
the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: remy.loris@vib-vub.be (RL); rene.demot@biw.kuleuven.be (RDM)
¤ Current address: Laboratory of Toxicology, University of Leuven, Leuven, Belgium.
. These authors contributed equally to this work.
" These authors also contributed equally to this work and should be considered joint senior authors.
Introduction
In most natural settings, complex interactions occur among
microorganisms,
ranging
from
nutritional
co-operation
to
warfare among competitors. Examples of such interplay have
been reported not only between unrelated microorganisms (e.g.
fungi and bacteria [1,2]), but also between distant relatives (e.g.
members of different bacterial genera [3]), and even between
close relatives (e.g. at inter- and intra-species levels [4,5]). A
major strategy in niche colonization is the production of growth
inhibitors or toxins directed at microbial competitors [6]. While
a huge variety of secondary metabolites is used to target
phylogenetically-distant competitors, ribosome-synthesized pep-
tides or proteins are typically active against close relatives.
These protein toxins are collectively referred to as bacteriocins,
and may either be released into the environment or transferred
to the host via specialized contact-dependent delivery systems
[7–9].
Bacteriocins are structurally and mechanistically very diverse.
This is reflected in the bacteriocinogenic potential of the genus
Pseudomonas [10]. Their R- and F-type pyocins are multi-subunit
protein complexes evolutionarily related to contractile tails of
bacteriophages [11–13]. R-pyocins attach to specific lipopolysac-
charide moieties at the cell surface of susceptible cells and insert
their core structure through the cell envelope, causing depolar-
ization of the cytoplasmic membrane [14]. The S-type pyocins of
Pseudomonas aeruginosa share structural and functional features with
Escherichia coli colicins [15]. Following docking onto surface-
exposed targets such as siderophore receptors [16,17], S-pyocins
kill cells
by nucleic
acid
degradation
[10,17], cytoplasmic
membrane damage [18], or inhibition of peptidoglycan synthesis
[19,20]. Putidacin A (or LlpABW), first identified in Pseudomonas
putida BW11M1 [21], represents a class of Pseudomonas-specific
antibacterial proteins not related to any known bacteriocin.
Additional llpA-like genes encoding functional bacteriocins were
identified by genome mining in the biocontrol strain Pseudomonas
fluorescens Pf-5 [22] and in the phytopathogen Pseudomonas syringae
pv. syringae 642 [23]. Identification of this type of protein in two
Xanthomonas pathovars extended its occurrence as a genus-specific
killer protein [23]. The Xanthomonas LlpA precursor is proteolyt-
ically processed by removal of a characteristic Type II secretion
signal peptide, whereas such N-terminal sequence is lacking in
Pseudomonas homologues, indicating that secretory routes may
differ among LlpA producers.
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The amino acid sequence of LlpA suggests the presence of
two related domains belonging to the ‘monocot mannose-
binding lectin’ (MMBL) family [24]. The MMBL domain
consists of a b-prism fold containing three potential carbohy-
drate-binding pockets, each generated by a QxDxNxVxY
sequence (with x, any amino acid), but some sites may be
inactive due to degeneracy of the signature motif [25]. This
domain (Pfam domain: B_lectin - PF01453) was initially
identified in lectins of monocot plants [26,27], but a more
widespread occurrence of MMBL lectins has become evident
and includes representatives in fungi [28,29], slime molds [30],
sponges [31], and fishes [32–34]. The LlpA branch occupies a
unique position among MMBL-domain proteins, harboring
non-eukaryotic representatives and being equipped with the
capacity to kill bacterial cells with bacteriocin-like specificity, a
property not yet demonstrated for other family members [25].
Next to proteins with the LlpA-type tandem-MMBL organiza-
tion, many other predicted MMBL proteins are encoded by
bacterial genomes. Often the MMBL module is embedded in a
larger protein. For one such protein, bacteriocin-like activity
among Ruminococcus species, Gram-positive bacteria colonizing
the rumen, was demonstrated [35].
Here we report on the crystal structure of LlpABW as the
prototype of a novel family of antibacterial proteins and explore
how domain architecture and specific structural elements contrib-
ute to its activity and specificity.
Results
LlpA forms a rigid MMBL tandem
The crystal structure of LlpABW from P. putida BW11M1
(LlpABW) shows it contains two b-prism MMBL domains,
referred to as the N-domain and the C-domain following their
position in the amino acid sequence (Figure 1A,B; Figure S1).
The N-domain spans residues Arg4-Pro135 while the C-domain
encompasses residues Ala136-Gln253. Each domain exhibits
pseudo-threefold symmetry and the corresponding subdomains
will be referred to as IN, IIN, IIIN, IC, IIC and IIIC, respectively
(Figure 1A and Figure S1). Following these two domains, a b-
hairpin extension is formed by residues Pro254-His275 (the
numbering used in this paper corresponds to that of the wild-type
protein without His-tag [21]).
The two-domain architecture reflects the b-strand swapping
that is typical in dimers of single-domain mannose-binding
monocot lectins (Figure 1A,B) [36] and which apparently is
retained after the ancestral fusion or duplication of the two
domains, as is also the case in certain MMBL tandems or
heterodimers from monocots [37,38]. Thus, residues Asp126-
Pro135 from the first MMBL sequence complement the fold of
the C-domain while residues Pro245-Gln253 from the second
MMBL sequence
complement the
fold of
the N-domain.
However, in LlpABW, the relative orientation of both domains
is different compared to what is observed in a canonical MMBL
lectin dimer, such as snowdrop lectin [36], in the heterodimeric
MMBL lectin ASA I from Allium sativum [38], or in the tandem
MMBL SCAfet from Scilla campanulata [37] (Figure 1C and Figure
S2). In contrast to these plant MMBL proteins, the resulting
architecture of LlpABW does not obey pseudo-twofold symmetry
(Figure 1C).
LlpABW is a very rigid molecule. The two monomers present in
the asymmetric unit are essentially identical with a root-mean-
square deviation (RMSD) of 0.34 A˚ for 270 Ca atoms. This
RMSD value does not change significantly when the individual
domains are fitted separately (0.32 A˚ for 120 Ca’s of the N-
domain and 0.22 A˚ for 115 Ca’s of the C-domain), indicating that
the inter-domain orientation is fixed. This stems from three sets of
interactions (Figure 2). Both domains are connected by a two-
stranded anti-parallel b-sheet that is involved in the b-strand
swapping mentioned above and that links both domains. The C-
terminal b-hairpin extension makes extensive contacts, through
hydrophobic and hydrogen bonds, with both domains. Finally, the
stretch Val140-Asp145 of the C-domain makes extensive contacts
with stretch Val115-Asp118 and with the side chains of Ser15 and
Pro32 of the N-domain.
Domains of LlpABW are shaped by differential
evolutionary pressure
A superposition of the Ca-trace of the N- and C-domain of
LlpABW as well as the MMBL domain of snowdrop lectin is shown
in Figure S3. Based on 79 Ca atoms that form the common b-
sheet core of the MMBL domains, the RMSD between the N- and
C-domains of LlpABW is 1.84 A˚ . While the secondary structure
elements of the C-domain are restricted to the three four-stranded
b-sheets of the b-prism fold, the N-domain contains three
additional secondary structure elements (Figure 1A). A three-turn
a-helix (a1) is inserted in the loop between strands b9 and b10,
and sheet IIN contains two additional strands. Strand b69 is
inserted in the loop between strands b6 and b7 and provides an
anti-parallel extension to sheet II (hydrogen bonding to strand b9).
Strand b19 is a short piece of b-strand that is part of the long N-
terminus and forms a parallel extension on the opposite site of
sheet IIN (hydrogen bonding to strand b2), making this b-sheet a
mixed type six-stranded one rather than the canonical four-
stranded anti-parallel sheet.
Despite these additions to the b-prism fold, the common core
of the N-domain more closely resembles that of the well-studied
and highly conserved monocot lectins (e.g. RMSD of 1.35 A˚ with
snowdrop lectin compared to 1.82 A˚ for the C-domain). This
structural divergence is in contrast with the degree of conserva-
Author Summary
In their natural environments, microorganisms compete
for space and nutrients, and a major strategy to assist in
niche colonization is the deployment of antagonistic
compounds directed at competitors, such as secondary
metabolites (antibiotics) and antibacterial peptides or
proteins (bacteriocins). The latter selectively kill closely
related bacteria, which is also the case for members of the
LlpA family. Here, we investigate the structure-function
relationship for the prototype LlpABW from a saprophytic
plant-associated Pseudomonas whose genus-specific tar-
get spectrum includes several phytopathogenic pseudo-
monads. By determining the 3D structure of this protein,
we could assign LlpA to the so-called monocot mannose-
binding
lectin
(MMBL)
family,
representing
its
first
prokaryotic member, and also add a new type of
protective function, as the eukaryotic MMBL members
have been linked with antiviral, antifungal, nematicidal or
insecticidal activities. For the protein containing two
similarly folded domains, we constructed site-specific
mutants affected in carbohydrate binding and domain
chimers from LlpA homologues to show that mannose-
specific sugar binding mediated by one domain is required
for activity and that the other domain determines target
strain specificity. The strategy that evolved for these
bacteriocins is reminiscent of the one used by mammalian
bactericidal proteins of the RegIII family that recruited a C-
type lectin fold to kill bacteria.
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Figure 1. Overall structure of LlpABW. (A) Topology diagram of LlpABW. The N-domain is shown in red, the C-domain in blue and the C-terminal
extension in green. The different strands and subdomains are labeled. Domain swapping involves b-strand segments b11b and b22b, which together
with b-strand segments b11a and b22a link both MMBL domains. (B) Cartoon representation of LlpABW with the different domains colored as in panel
A. The bound Me-Man residue is shown as an orange stick representation. (C) Domain orientations of LlpABW compared with the heterodimeric MMBL
ASA I (Allium sativum agglutinin, PDB entry 1KJ1) and tandem MMBL SCAfet (Scilla campanulata fetuin-binding lectin, PDB entry 1DLP). In each case,
the C-domain is shown in the same orientation, highlighting the different relative orientation of the N-domain in LlpABW. Domain-swapped dimers in
homo-oligomeric plant MMBL lectins such as snowdrop lectin have their domain orientation similar to ASA I and SCAfet.
doi:10.1371/journal.ppat.1003199.g001
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tion of the carbohydrate-binding motif characteristic of the
monocot lectins (QxDxNxVxY) in each of the three subdomains.
In the N-domains of LlpA homologues, the surface-exposed
motifs III and II are not well conserved and likely lost their
function during evolution. In contrast they seem to be better
conserved in the C-domains (Figure S4). Apparently, the two
MMBL domains of LlpA experienced a differential evolutionary
pressure resulting in different degrees of global and local
(carbohydrate-binding motif) conservation, suggesting distinct
functional roles for each domain.
The C-domain of LlpABW further extends into a b-hairpin
that helps to define the relative orientations of its two MMBL
domains. This b-hairpin is highly bent due to a b-bulge inserted
into its second b-strand (Figure 1B). It is absent in all plant
representatives including tandem MMBL proteins such as
SCAfet (Figure 1C). In bacteria it represents the most divergent
part of LlpA homologues, both in primary sequence and in
length
(Figure
S5).
Most
of
these
C-terminal
extensions
terminate with a phenylalanine residue. This is reminiscent of
the conserved terminal phenylalanine of outer membrane
proteins from Gram-negative bacteria such as PhoE, required
for their translocation to the cell envelope [39]. An equivalent
extension
appears
to
be
absent
in
the
Xanthomonas
and
Arthrobacter sequences (Figure S5).
LlpA is capable of binding mannose-containing
carbohydrates
Subdomains IIC and IIICof LlpABW contain the typical sugar-
binding signature (QxDxNxVxY) of an active MMBL mannose-
binding site (Figure S1 and S4). Soaking crystals of LlpABW with
200 mM methyl-a-D-mannopyranoside (Me-Man) led to clear
electron density of a single Me-Man in site IIIC of each of the
two LlpABW monomers in the asymmetric unit (Figure S6A).
This site comprises the side chains from Gln171, Asp173,
Asn175 and Tyr179, which contribute to hydrogen bond
interactions and the side chains of residues Val177, Asn188,
Gln192 and Ala185, which contribute to van der Waals contacts
with the carbohydrate ligand (Figure 3A, Figure S7A,C). This
architecture is very similar to what is observed for mannose
bound to other MMBL-type lectins such as snowdrop and garlic
lectin (Figure S7B).
Soaks with oligomannoses revealed additional sugar-binding
subsites.
Binding
site
IIIC
accommodates
the
disaccharide
Mana(1–2)Man and the pentasaccharide GlcNAcb(1–2)Mana(1–
3)[GlcNAcb(1–2)Mana(1–6)]Man (Figure S6B,C). In the case of
Figure 2. Domain interactions within LlpABW. (A) The C-terminal hairpin extension (green cartoon) covers the interface between the N-domain
(red surface representation) and the C-domain (blue surface representation). (B) Stereo view of the interactions between loop segments Val140-
Asp145 (cyan) of the C-domain and Val115-Asp118 (yellow) and Ser31-Gln34 (orange) of the N-domain. Other structural elements are colored
according to panel A. (C) Stereo view of the two-stranded b-sheet formed by strands b11a,b and b22a,b that links the N- and the C-domains and gives
rise to domain swapping. Colors according to panel A and B.
doi:10.1371/journal.ppat.1003199.g002
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the pentasaccharide, the central reducing mannose is located in
the shallow Me-Man binding site and the two GlcNAcb(1–2)Man
moieties stretch out over the surface making only a few additional
hydrogen bonds or van der Waals contacts (Figure 3B). In the
bound disaccharide, the non-reducing mannose is located in the
Man-Me binding site while the reducing mannose faces the solvent
and does not interact directly with the protein (Figure 3C).
Site IIC of both LlpABW molecules in the asymmetric unit is
involved in crystal packing interactions and the presence of Me-
Man is therefore sterically excluded. All residues that form specific
hydrogen bonds with Me-Man are retained but substitutions occur
for three side chains that provide van der Waals contacts (Figure
S4 and S8A). In contrast, site IC lost the conserved QxDxNxVxY
motif (Figure S4) and is involved in inter-domain contacts and
therefore inaccessible to ligands (Figure S8B).
The putative carbohydrate-binding sites in the N-domain of
LlpABW are less conserved. Similar to the C-domain, site IN is
inaccessible and involved in inter-domain interactions (Figure
S9A). In the IIN subdomain, the canonical mannose-binding motif
QxDxNxVxY is essentially absent, with only the Gln residue of the
motif being conserved as Gln82 (Figure S4). All other donors or
acceptors required for hydrogen bonds with a mannose ligand are
missing. In addition, the presence of Phe86 at the equivalent
position of the expected Val sterically hinders the binding of
mannose (Figure S9B). The potential carbohydrate-binding site on
subdomain IIIN is only partially conserved (Figure S9C) and
contains two relevant substitutions from the canonical signature:
Figure 3. Carbohydrate binding in site IIIC of LlpABW. (A) Stereoview of methyl-a-D-mannopyranoside bound to subdomain IIIC. Methyl-a-D-
mannopyranoside is shown in blue and indicated by M. Residues belonging to the QxDxNxVxY motif and hydrogen bonding to the sugar as well as
Asn188 are labeled. Water molecules bridging protein and carbohydrate are shown in cyan (B) Similar view of the pentasaccharide GlcNAcb(1–
2)Mana(1–3)[GlcNAcb(1–2)Mana(1–6)]Man. The mannose residue occupying the primary binding site is shown in blue and labeled M. The additional
two mannoses (labeled +1 and 21) and two N-acetyl glucosamine residues (labeled +2 and 22) are shown in green. Other colors are as in panel A. (C)
Binding of the disaccharide Mana(1–2)Man. The non-reducing mannose residue occupying the primary binding site is shown in blue and labeled M.
The second, reducing mannose is shown in green. Other colors are as in panel A.
doi:10.1371/journal.ppat.1003199.g003
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(1) the Tyr residue of the QxDxNxVxY motif is replaced by the
shorter Gln49, thereby removing the canonical hydrogen bond
between Man O4 and Tyr OH, and (2) a threonine at position
54 which may compensate the hydrogen bond lost due to the
Tyr-to-Gln substitution in the canonical motif. The lack of
electron density at this site in our Me-Man soak nevertheless
indicates that this site does not recognize this ligand or that its
affinity is so low that recognition would only be achieved in the
context of a larger and as yet unidentified mannose-containing
ligand. Alternatively, this putative site may possess specificity for a
different monosaccharide. In order to evaluate this hypothesis, we
soaked LlpABW crystals with D-galactose, N-acetyl-D-glucosamine
and L-fucose. No electron density was observed for any of these
sugars, suggesting that the N-domain has a function distinct from
carbohydrate recognition (data not shown).
Carbohydrate-binding capacity is required for LlpA
toxicity
The LlpABW motifs IIIN, IIIC and IIC create potential
carbohydrate binding sites that may be involved in bacteriotoxicity
of the protein. We therefore examined the role of carbohydrate
binding in the bactericidal function of LlpABW. The presence of
methyl-a-D-mannopyranoside up to 500 mM in the medium did
not influence the activity of LlpABW on P. syringae GR12-2R3.
Glycan array profiling did not highlight any specific oligosaccha-
ride structure that could represent a natural ligand of LlpABW
(Table S1). This could be due to the array design that is principally
based on eukaryotic glycans and may therefore lack an appropri-
ate carbohydrate for this prokaryotic toxin. Previously, it was
observed that LlpABW from concentrated culture supernatant does
not agglutinate rabbit red blood cells, nor binds to a mannose-
agarose affinity matrix [21].
To assess whether the mannose-recognizing QxDxNxVxY
motifs in LlpABW are nevertheless relevant for bactericidal activity,
the
conserved
valine
residue
was
mutated
to
tyrosine
in
subdomains IIIN, IIIC, and IIC. These mutations sterically
preclude mannose or any other ligand to enter the binding sites
(Figure S7C). Semi-quantitative activity assays with permeabilized
E. coli cells expressing the LlpA variants in motifs IIIN, IIIC and IIC
were used to assess the relationship between carbohydrate binding
and bactericidal activity. Modification of the IIIN site, for which no
mannose binding was observed, does not affect the antibacterial
activity against P. syringae GR12-2R3 (Figure 4). In contrast, the
altered IIIC pocket strongly diminishes activity, either alone or in
pairwise combination with the other mutated sites (IIIN or IIC). A
minor negative effect of the IIC mutation is only apparent in a
double mutant, when combined with a modified IIIN motif.
Purified proteins were prepared to further quantify these effects.
Far UV CD spectra of these mutant forms are identical to that of
native protein LlpABW, indicating that the mutations do not affect
the overall structure of the protein. Isothermal titration calorim-
etry (ITC) showed that LlpABW has an affinity of 2.1 mM for the
pentasaccharide
GlcNAcb(1–2)Mana(1–3)[GlcNAcb(1–2)Ma-
na(1–6)]Man, the highest among all the tested oligo-mannosides
(See Figure 5 and Table 1 for a summary of the experimentally
validated LlpABW-carbohydrate interactions). This is in agreement
with the crystal structures of the different complexes since this
sugar is the one with the largest binding interface (Figure 3).
Titrations of LlpABW, of the mutants LlpAV177Y (a site IIIC
knockout), LlpAV208Y (a site IIC knockout) and of the double
mutant LlpAV177Y-V208Y with a-methyl mannoside clearly pin-
point site IIIC as the only responsible for the sugar binding activity.
Point mutations in both sites or IIIC (V177Y) alone, completely
abrogate sugar binding. However knocking out site IIC (V208Y)
has little effect in binding and the affinities of LlpAV208Y for a-
methyl mannoside and Mana(1–3)Man are very close to the ones
measured for the wild-type protein (See Table 1 and Figure 5B).
While the V208Y mutation in the IIC site has no observable
effect on the MIC value for P. syringae GR12-2R3, the altered IIIC
motif engenders a 5.2-fold increase in MIC (Figure 4). The mutant
protein LlpAV177Y-V208Y suffers a further reduction in activity,
yielding a 31.6-fold increased MIC compared to native LlpABW.
The biological activities of LlpA and its mutants were further
assessed by live/dead staining and subsequent flow cytometry
analysis (Figure 6, Figure S10). Proportions of dead cells after
1 hour of exposure to LlpA or LlpAV208Y were comparable (10.1%
and 9.7%, respectively). For LlpAV177Y this value was reduced to
6.1%, significantly lower than for LlpA. Killing activity was even
further reduced for LlpAV177Y-V208Y (3.7%). These results are
consistent with the MIC determination and ITC data, indicating
that an active site IIIC is required to generate a fully active LlpA
bacteriocin. The difference in bacteriotoxicity between LlpAV177Y
and LlpAV177Y-V208Y suggests that site IIC has a supporting role in
the LlpABW bacteriotoxicity.
All domains are necessary for LlpABW functionality
The site-directed mutagenesis approach revealed an important
role for the C-domain’s carbohydrate-binding capacity in LlpABW
toxicity. Considering the increased binding motif degeneration in
the N-domain and the fact that a Ruminococcus bacteriocin
composed of only a single MMBL domain fused to an unknown
domain has been identified [35], the N-domain may fulfill a
distinct function, different from that of the C-domain. In order to
scrutinize the contribution of individual domains to overall
activity, six domain deletion constructs of llpABW were engineered
to potentially encode proteins lacking the first or second MMBL
domain, a gene product devoid of the C-terminal hairpin, or a
protein retaining only an individual domain (N-domain, C-
domain, or hairpin) (Figure S11). To take the domain swapping
into account, the constructs containing only a single MMBL
domain were designed with a fusion of the swapped C-terminal b-
strands to the corresponding domain via a short linker.
None of these deletion constructs resulted in the production of
an active protein, indicating that none of the domains are
dispensable. Removal of the terminal phenylalanine residue still
allows expression of a functional bacteriocin in E. coli (Figure 7),
but a further C-terminal truncation (deletion of Trp-His-Phe tail)
resulted in a negative bacteriocin assay (data not shown). From
these data we conclude that both MMBL domains as well as the
C-terminal hairpin extension are required for activity of LlpA.
Whether the role of the C-terminal hairpin is any other than
simply stabilization of the C-domain cannot be concluded.
Target specificity of LlpA is hosted by the N-domain
In order to investigate the role of the different domains in target
specificity, we created hybrid LlpA proteins using the domains of
LlpABW from P. putida BW11M1 and LlpA1 from P. fluorescens Pf-5.
These two LlpA proteins share 45% sequence identity and differ in
their target spectra. Strains P. syringae GR12-2R3 and P. fluorescens
LMG1794 were identified as specific indicators for LlpABW [21]
and LlpA1 [22], respectively. Six constructs carrying llpABW/
llpA1chimeric genes were made with domain exchanges involving
the N-domain, C-domain, and hairpin region (Figure 7 and Figure
S11). For four of these constructs activity against one of both
indicators was detected. Only constructs retaining the original N-
domain give rise to inhibition of the cognate indicator strain. The
C-domain or the hairpin of LlpABW could be replaced with the
corresponding LlpA1 domains without changing target specificity.
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Conversely, the original specificity of LlpA1 is retained upon
replacement of its C-domain by the LlpABW equivalent.
Discussion
Structure elucidation of LlpABW from P. putida BW11M1
unequivocally assigns this bacteriocin to the MMBL lectin family,
in which it constitutes the first prokaryotic member, representative
for a group of bacterial proteins composed of two MMBL domains
[21–23]. Systematic inactivation of the three potential carbohy-
drate-binding sites present in the N-domain (IIIN) and in the
C-domain (IIIC and IIC) of LlpABW, revealed that a non-occluded
IIIC pocket is required to obtain a fully active LlpABW molecule. A
negative co-operative effect on activity resulted when the IIC site
was additionally modified. Although mannose-containing carbo-
hydrates can bind to the IIIC pocket of LlpABW, it remains unclear
Figure 4. Inhibitory activity of wild-type LlpABW and selected mutants with modified (potential) mannose-binding sites. The domain
structure (N-domain in red, C-domain in blue and C-terminal extension in green) and the position of the MMBL motifs (potentially active binding sites
in orange, inactive ones in grey) are shown. The positions of conserved valine residues converted to tyrosine residues by site-directed mutagenesis
are indicated with a black bar. Inhibitory activity of E. coli strains expressing mutant LlpABW forms was assayed against P. syringae GR12-2R3 and semi-
quantified according to the size (inner zone radius) of the growth inhibition halo relative to LlpABW (+++, native LlpABW; ++, halo size reduced; + halo
size strongly reduced; 2, no halo; NT, not tested). For wild-type LlpABW and three purified His-tagged mutant forms (LlpAV177Y, LlpAV208Y and
LlpAV177Y-V208Y) the MIC values were determined with indicator P. syringae GR12-2R3. Molar minimal inhibitory concentrations of recombinant
proteins (with standard deviations): LlpA, 2.08 nM (60.58 nM); LlpAV177Y, 10.9 nM (60.66 nM); LlpAV208Y, 1.98 nM (60.066 nM); 65.72 nM
(62.80 nM).
doi:10.1371/journal.ppat.1003199.g004
Table 1. Binding affinities and thermodynamic parameters obtained from ITC titrations.
Type of protein-carbohydrate
interaction
Kd (mM)
DG6(kcal mol21)
DH6 (kcal mol21)
2TDS6(kcal mol21)
LlpABW Me-a-D-Man
45.9
21.8
25.4
3.6
LlpABW Mana(1–2)Man
42.4
21.9
23.6
1.7
LlpABW Mana(1–3)Man
18.2
22.4
25.9
3.5
LlpABW Mana(1–6)Man
17.2
22.4
25.5
3.1
LlpABW Mana(1–3)[Mana(1–6)]Man
10.1
22.6
26.4
3.8
LlpABW GlcNAcb(1–2)Mana(1–3)[GlcNAcb(1–
2)Mana(1–6)]Man
2.1
23.7
21.6
22.1
LlpAV208Y Me-a-D-Man
58.8
21.7
23.3
1.6
LlpAV208Y Mana(1–3)Man
23.0
22.2
25.1
2.9
The reported values for Kd, DGu, DHu and 2TDSu were determined from fitting a single site interaction model (n = 1) to the experimental ITC data. The interaction of the
mutants LlpAV177Y and LlpAV177-V208Y with the different sugars is negligible and no heat effect was observed. Therefore they are not included in this table.
doi:10.1371/journal.ppat.1003199.t001
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if these are part of or mimic the natural ligand required for
biological activity since bacteriocin activity is not impaired in the
presence of excess mannose.
A mutated IIIN site did not provoke a negative effect on
antibacterial activity. However, the N-domain appears to play a
major role in target selection. This was demonstrated by
assessing the differential activity of domain chimers against
two target strains, diagnostic for the LlpABW- and LlpA1-
specific killing.
The b-hairpin does not appear to be a specificity determi-
nant, although it constitutes the most variable region among
LlpA-like bacteriocins. Possibly, it is required for thermody-
namic stability since it needs to be intact in LlpABW. An
equivalent C-terminal stretch is absent from the Xanthomonas citri
LlpA-like
bacteriocin
[23].
From
our
results
relying
on
heterologous expression in E. coli and a bacteriocin assay with
permeabilized cells, it cannot be excluded that this structural
element may play a role in the way an LlpA protein is exported
by its native producer cells.
Figure 6. Killing activity of LlpABW and mutant proteins. Percentages of dead cells after live/dead staining as quantified by flow cytometry
analysis (Figure S10). P. syringae GR12-2R3 was used as indicator strain and treated at a final concentration of 50 mg/ml for 1 h. Average values (with
standard deviations; indicated by error bars): LlpA, 10.1 (61.04); LlpAV177Y, 6.1 (60.44); LlpAV208Y, 9.7 (61.39); LlpAV177Y-V208Y, 3.7 (60.90); buffer
(control), 1.0 (60.11).Values are significantly different for (a) and (b), (b) and (c) (p,0.01).
doi:10.1371/journal.ppat.1003199.g006
Figure 5. ITC analysis of carbohydrate binding to LlpABW and mutants. (A) Binding of LlpABW to the pentasaccharide GlcNAcb(1–2)Mana(1–
3)[GlcNAcb(1–2)Mana(1–6)]Man. (B) Binding of LlpABW (blue circles, wild type) and the mutants LlpAV177Y (green circles, site IIIC knockout), LlpAV208Y
(red circles, site IIC knockout) and LlpAV177Y-V208Y (black circles, site IIC and IIIC knockout) to a-methyl mannoside. There is no heat exchanged in the
titration of the double mutant or the site IIIC knockout LlpAV177Y, whereas the site IIC knockout LlpAV208Y, binds the monosaccharide in a ‘‘wildtype’’-
like fashion, showing that only site IIIC is involved in sugar binding.
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In general, a defensive role has been proposed for the
(oligo)mannose-binding MMBL lectins based on insecticidal,
nematicidal, antifungal, or even antiviral activities demonstrated
for several of these proteins that are abundantly found in monocot
plants [40–46]. Some of these plant lectins trigger apoptosis in
cancer cells [47]. Also their identification in fish mucus and
epithelial cells is in line with a general protective (antimicrobial)
function for MMBL domains [32]. LlpABW as a bactericidal
protein fits within this picture of MMBL domains being involved
in general defense mechanisms. Since no antibacterial activity has
been assigned to the eukaryotic MMBL proteins, it is challenging
to identify structural features that confer the intragenus-specific
bacteriocin activity of LlpA, as shown for proteins from P. putida
[21], P. fluorescens [22], P. syringae [23], and Xanthomonas citri [23].
Their
target
spectra
are
narrower
than
reported
for
the
mammalian antibacterial C-type lectins of the RegIII family, such
as mouse RegIIIc and its human homolog HIP/PAP that bind to
the surface-exposed peptidoglycan layer of Gram-positive bacteria
[48], and RegIIIb that also binds to the lipid A moiety of
lipopolysaccharides
on
the
cell
envelope
of
Gram-negative
bacteria [49].
The absence of any known secretory signal sequence in
LlpABW and its homologues in other Pseudomonas species is
intriguing in view of their extracellular location [21]. The
translocation of the outer membrane-associated mannose/
fucose-specific lectin LecB of P. aeruginosa, that also lacks such
signal sequence [50], is dependent on its glycosylation [51].
Contrary to LlpA that is exported to the culture supernatant to
exert its antagonistic activity, LecB remains associated with the
cell
envelope
through
interaction
with
the
major
outer
membrane protein OprF [52], in line with its role in biofilm
formation.
Materials and Methods
Strains and culture conditions
Bacterial strains and plasmids used in this study are listed in
Table S2. Escherichia coli was routinely grown in shaken Luria-
Bertani (LB, MP Biomedicals) broth at 37uC. Pseudomonas strains
were grown in Tryptic Soy Broth (BD Biosciences) at 30uC with
shaking. Media were solidified with 1.5% agar (Invitrogen) and
supplemented with filter-sterilized antibiotics as required at
following concentrations: ampicillin (Sigma-Aldrich), 100 mg/ml
or kanamycin (Sigma-Aldrich), 50 mg/ml. Isopropyl b-D-thioga-
lactoside (IPTG 40 mg/ml, ForMedium) and 5-bromo-4-chloro-3-
indolyl-b-D-galactopyranoside (X-Gal 40 mg/ml, ForMedium)
were added for blue/white screening of pUC18-derived plasmids
in E. coli.
Plasmids used for antibacterial testing and sequencing were
propagated in E. coli TOP10F9 (Invitrogen). E. coli BL21(DE3)
(Novagen) was used as a host for plasmids driving recombinant
protein expression. Genomic DNA from P. putida BW11M1 was
isolated using the Puregene Yeast/Bact. Kit B (Qiagen). Plasmid
DNA was extracted using the QIAprep Spin Miniprep Kit
(Qiagen). Stocks were stored at 280uC in the appropriate medium
in 25% (v/v) glycerol.
Figure 7. Differential inhibitory activity of wild-type LlpABW and LlpA/LlpA1 domain chimers. The domain structures of LlpABW (as in
Figure 4) and of LlpA1 (inferred by pairwise alignment; N-domain in orange, C-domain in purple and C-terminal extension in grey) are depicted, along
with those of chimeric forms (in dashed box). The LlpA variant lacking the terminal phenylalanine residue is marked with a yellow hexagon. Inhibitory
activity of the respective E. coli recombinants was tested with diagnostic indicators for LlpABW (P. syringae GR12-2R3) and LlpA1 (P. fluorescens LMG
1794). Halo sizes are semi-quantified according to size of the growth inhibition halo (+++, native halo size of LlpABW and LlpA1; ++, halo size reduced;
C, local clearing confined to producer colony spot; 2, no halo or clearing; NT, not tested). Additional chimeric and domain deletion constructs not
conferring bacteriocin activity against one of the indicator strains are specified in Figure S11.
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Recombinant DNA methods
Standard methods were used for preparation of competent E.
coli cells, heat shock transformation of E. coli and DNA
electrophoresis [53]. Restriction enzymes were used according to
the supplier’s specifications (Roche Diagnostics and BIOKE´).
DNA ligation was performed using T4 DNA ligase (Invitrogen).
Plasmid sequencing was performed by GATC Biotech (Constance,
Germany). Constructs that were generated are listed in Table S2
and primers are listed in Table S3.
A 921-bp fragment containing llpABW was amplified by PCR
with Platinum Pfx DNA polymerase (Invitrogen), using a C1000
Thermal Cycler (Bio-Rad). P. putida BW11M1 genomic DNA
was taken as a template, and combined with primers PGPRB-
3155 and PGPRB-3156. The amplicon was purified using the
QIAquick PCR Purification Kit (Qiagen), digested with KpnI
and BamHI, ligated in pUC18, and transformed to E. coli
TOP10F9. Transformants were verified for the presence of the
insert by PCR using Taq Polymerase (BIOKE´ ) with primers
PGPRB-2545
and
PGPRB-2546.
The
cloned
construct
(pCMPG6129)
was
purified
and
its
insert
confirmed
by
sequencing.
DpnI-mediated site-directed mutagenesis was performed to
construct valine-to-tyrosine mutant forms of llpABW in pUC18
(pCMPG6129) and N-terminal His-tagged llpABW in pET28a
(pCMPG6056 [54]). PCR conditions were: 2 min initial denatur-
ation, followed by 16 cycles of denaturation (1 min), annealing
(1 min, primer-dependent temperature) and elongation at 68uC
(1 min./kb). Final elongation was for 8 min at 68uC. After PCR,
samples were immediately treated with DpnI at 37uC for 2 h and
transformed into E. coli TOP10F9 and selected on the appropriate
medium. Plasmid inserts of selected transformants were verified by
sequence
analysis. Double
mutants
were
constructed
using
plasmids with a single point mutation as a template.
Domain
deletants
of
llpABW
were
constructed
using
pCMPG6129 as a template (llpABW from P. putida BW11M1).
Chimeric constructs were obtained using pCMPG6129 and
pCMPG6053 (llpA1 from P. fluorescens Pf-5 [22]) as templates.
Artificial ligation of gene fragments, generated with the PCR
primers specified in Table S3, was performed by using splicing by
overlap extension (SOE). The resulting recombinant amino acid
sequences are listed in Table S4.
Recombinant protein expression and purification
Protein isolation and purification of N-terminal His6-tagged
LlpABW, LlpAV177Y, LlpAV208Y, and LlpAV177Y-V208Y from E. coli
BL21(DE3),
carrying
expression
constructs
pCMPG6056,
pCMPG6149, pCMPG6150 and pCMPG6151 respectively, were
performed as described by Parret and collaborators [54]. The
presence of His-tagged protein was observed via immunodetection
by Western blot, using monoclonal anti-His6 (IgG1 from mouse;
Roche Diagnostics) as primary antibody. Fractions free of other
proteins, as verified by SDS-PAGE and subsequent Coomassie
Blue staining, were dialyzed against bis-TRIS propane buffer
(20 mM, 200 mM NaCl, pH 7.0). Concentrations of purified
proteins were determined by absorbance measurement at 280 nm
using molar extinction coefficients of 62910 M21 cm21 for
LlpABW, 64400 M21 cm21 for LlpAV177Y and LlpAV208Y, and
65890 M21 cm21 for LlpAV177Y-V208Y. Extinction coefficients
were calculated according to Pace and collaborators [55].
Antibacterial assays
Bacteriocin production by E. coli cells carrying pUC18-derived
constructs was assayed as follows: 2-ml drops of an overnight
stationary-phase culture were spotted onto LB agar plates and
incubated for 8 h at 37uC. Next, plates were exposed to
chloroform vapor (30 min), aerated and subsequently overlaid
with 5 ml of soft agar (0.5%), seeded with 200 ml of a cell culture of
an indicator strain (,108 CFU/ml), followed by overnight
incubation at 30uC. Next day, plates were scored for the presence
of halos in the confluently grown overlay.
Antibacterial activity assays with purified recombinant His6-
tagged proteins were performed as described by Ghequire and
collaborators [23]. To assess the influence of added sugar, the
same assay was carried out on agar medium supplemented with D-
mannose (Sigma-Aldrich) or methyl-a-D-mannopyranoside (Sig-
ma-Aldrich) to a final concentration of 0.01 M to 0.5 M.
A Bioscreen C apparatus (Oy Growth Curves Ab Ltd, Finland)
was used to determine the minimum inhibitory concentration
(MIC). An overnight culture (16 h) of the indicator strain was
diluted to 104–105 CFU/ml and incubated at 30uC, with a two-
fold dilution series of recombinant His6-tagged LlpABW or mutant
LlpABW. Bis-TRIS propane buffer was used as control. The MIC
value was determined as the minimum concentration of protein at
which no growth of the indicator strain (OD600,0.2) occurred
after 24 h. At least three independent repeats, each with three
replicates, were carried out.
Glycan array
His6-tagged LlpABW was lyophilized and verified for antibac-
terial activity. After re-dissolving in MilliQ water, recombinant
LlpABW was diluted to 200 mg/ml with binding buffer (20 mM
TRIS-HCl pH 7.4, 150 mM NaCl, 2 mM CaCl2, 2 mM MgCl2,
0.05% Tween 20, 1% BSA), and used to probe the printed glycan
arrays [56] following the standard procedures of Core H of the
Consortium
for
Functional
Glycomics
(http://www.
functionalglycomics.org/).
Monoclonal
anti-His6
antibodies
(Roche Diagnostics) were used as primary antibodies, and
fluorescently labeled anti-mouse IgG as secondary antibodies.
Circular dichroism
CD spectra were acquired on a Jasco J-715 spectropolarimeter.
Curves were averaged over 8 scans, taken at 25uC using a 1 mm
cuvette. Samples were dialyzed against bis-TRIS propane buffer
(20 mM, NaCl 200 mM, pH 7.0), filtered and degassed prior to
data acquisition. All proteins were assayed at 0.4 mg/ml.
Isothermal titration calorimetry
ITC titrations were carried out on an ITC200 apparatus
(MicroCal). Prior to the measurement, LlpABW, LlpAV177Y,
LlpAV208Y and LlpAV177Y-V208Y was dialyzed to bis-TRIS
propane buffer. Sugars were directly dissolved into the same
buffer. The samples were filtered and degassed for 10 min at
25uC before being examined in the calorimeter. The titrations
were carried out at 25uC, injecting the sugars (methyl-a-D-
mannoside, Mana(1–2)Man, Mana(1–3)Man, Mana(1–6)Man,
Mana(1–3)[Mana(1–6)]Man
and
GlcNAcb(1–2)Mana(1–
3)[GlcNAcb(1–2)Mana(1–6)]Man) into a protein solution (pro-
tein concentrations ranged from 2 mM to 4 mM depending on
protein availability). All data were analyzed using the MicroCal
Origin ITC 7.0 software. Binding affinities and thermodynamic
parameters from all ITC titrations are reported in Table 1.
X-ray data collection and structure determination
Expression, purification and crystallization of recombinant His-
tagged LlpABW have been described [54]. X-ray data for native and
derivative crystals were collected on EMBL beamline BW7A of the
DESY synchrotron (Hamburg, Germany). For each potential
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derivative, the wavelength was chosen to be at the high-energy side
of the absorption edge in order to ensure a usable anomalous signal.
All data were scaled and merged with the HKL package of
programs. Data collection statistics are given in Table S5.
The crystal structure of free LlpABW was solved combining
single
isomorphous
replacement
with
anomalous
scattering
(SIRAS strategy) from a p-chloromercurybenzoate derivative.
The heavy-atom substructure was determined with SHELXD
[57] using a resolution cutoff of 4.0 A˚ . Heavy-atom refinement
and phasing were performed with SHARP [58]. Phase improve-
ment by solvent flattening was performed with SOLOMON [59].
Non-crystallographic symmetry averaging with density modifica-
tion [60] further improved the electron density. A partial model
(94% of the residues comprising the asymmetric unit) was
automatically built with ARP/wARP [61] and the remainder
was built manually over several cycles of model building with Coot
[62], alternated with refinement using phenix.refine [63,64]. Phasing
and refinement statistics are shown in Table S5.
Carbohydrate soaks
Crystals of LlpABW were transferred to artificial mother liquor
(0.1 M imidazole pH 6.5, 1.3 M sodium acetate) enriched with
either 200 mM methyl-a-D-mannopyranoside (Me-Man), Ma-
na(1–2)Man,
GlcNAcb(1–2)Mana(1–3)[GlcNAcb(1–2)Mana(1–
6)]Man (M592), D-galactose, L-fucose or N-acetyl-D-glucosamine
and allowed to equilibrate overnight (all carbohydrates obtained
from Dextra Laboratories, Reading, U.K.). Data were collected at
room temperature on EMBL beamline X13, except for the N-
acetyl-D-glucosamine soak collected at the PROXIMA-1 beam-
line of the SOLEIL synchrotron (Gif-sur-Yvette, France) and the
D-galactose soak collected at ESRF beam line ID14-1 (Grenoble,
France). All data were scaled and merged using the HKL package.
Refinement was started from the coordinates of the ligand-free
structure using phenix.refine. Manual rebuilding, including the
introduction of the carbohydrate ligand if present, was done with
Coot [62]. Crystal structures of LlpABW from P. putida BW11M1
(PDB entry 3M7H) and LlpABW in complex with methyl-a-D-
mannoside (PDB entry 3M7J), with Mana(1–2)Man (PDB entry
4GC1), and with GlcNAcb(1–2)Mana(1–3)[GlcNAcb(1–2)Ma-
na(1–6)]Man (PDB entry 4GC2) have been deposited at the PDB.
Flow cytometry
Overnight cultures of P. syringae GR12-2R3 (16 h) were diluted
to OD600 0.5 and washed twice with phosphate-buffered saline
(PBS). Cells were treated with LlpA, mutant proteins or buffer (bis-
TRIS propane buffer, negative control), at a final concentration of
50 mg/ml for 1 h, at 20uC. Next, PBS-washed bacteria were
stained using the Live/Dead BacLight bacterial viability kit
(Invitrogen), incubated for 15 minutes, and analyzed on a BD
Influx (BD Biosciences). Excitation of the dyes was done at
488 nm, and fluorescence measured at 530 nm for SYTO 9 and at
610 nm for propidium iodide. Results were processed with FlowJo
10.0.4 software (Figure S10). Measurements were done indepen-
dently and based on six biological repeats. Results are expressed as
percentages of dead cells [dead/(live+dead) * 100)].
Supporting Information
Figure
S1
Amino acid sequence of LlpABW colored
according to its domain structure. The N-domain is shown
in red, the C-domain in blue and the C-terminal extension in green.
Residues belonging to sequences equivalent to the mannose binding
site signature motif QxDxNxVxY are in bold and underlined.
(JPG)
Figure S2
Quaternary structures and domain organi-
zation of various MMBL family members. Individual
domains or protomers are shown in different colours. The
domain or protomer colored green (which in the tandem MMBLs
of LlpA, ASA I and SCAfet corresponds to the N-terminal
domain) is always shown in the same orientation. Bound
carbohydrates are shown in black stick representation. For LlpA
a single pentasaccharide is bound to site IIIC. In the case of
Galanthus nivalis (snowdrop) lectin (PDB entry 1JPC), twelve
trimannosides are bound to all QxDxNxVxY motifs (three on
each monomer of the homotetrameric protein). The snowdrop
lectin tetramer consists of the association of two domain-swapped
dimers (green-blue and pink-yellow). In the case of Allium sativum
(garlic lectin ASA I - PDB entry 1KJ1), again each QxDxNxVxY
motif has a dimannose bound while an additional sugar (shown in
red) is bound to a non-canonical site. The protein is synthesized
as a single chain precursor and post-translationally cleaved into
two MMBL domains that adopt the same domain-swapped dimer
as found in snowdrop lectin. Gastrodianin is a monomeric
MMBL family member from the orchid Gastrodia elata (PDB entry
1XD5). The location(s) of its carbohydrate-binding site(s) is (are)
not known. The fetuin-binding tandem-MMBL SCAfet from
Scilla campanulata (PDB entry 1DLP) consists of two covalently
attached MMBL domains, whereas in LlpA the swap of the C-
terminal b-strands is retained. The relative orientation in the two
domains is as in ASA I. This lectin binds fetuin rather than
oligomannosides, but the locations of the binding sites are not
known.
(JPG)
Figure S3
Stereo view of the superpositions (Ca repre-
sentations) of the N-domain of LlpABW (red), C-domain
of LlpABW (blue) and Galanthus nivalis lectin (PDB entry
1MSA, black). The superposition is shown in two orientations
rotated by 90u.
(JPG)
Figure S4
Sequence alignment of potential mannose-
binding motifs in prokaryotic tandem MMBL proteins.
The
sequences
corresponding
to
the
consensus
motif
QxDxNxVxY, extracted from the N-domain and the C-domain
of P. putida LlpABW and its homologues, are aligned per domain.
Sequence conservation is visualized by differential shading. The
sequence logo representation visualizes the degree of consensus for
each residue. LlpA proteins with proven bacteriotoxic activity are
labeled with an asterisk. Accession numbers: Arthrobacter sp. FB24
(YP_829274), Burkholderia ambifaria MEX-5 (ZP_02905572), Burk-
holderia cenocepacia AU 1054 ([1], ABF75998; [2], ABF75999),
Pseudomonas
chlororaphis
subsp.
aureofaciens
30–84
(EJL08681),
Pseudomonas putida BW11M1 (AAM95702), Pseudomonas fluorescens
Pf-5 (LlpA1 [1], YP_258360; LlpA2 [2], YP_259234), Pseudomonas
sp. GM80 ([1], ZP_10606046; [2], ZP_10606131), Pseudomonas
syringae pv. aptata DSM 50252 (EGH77666), Pseudomonas syringae pv.
syringae 642 (ZP_07263221), Xanthomonas axonopodis pv. citri str. 306
(AAM35756).
(TIF)
Figure S5
Sequence alignment of the carboxy-terminal
sequences of LlpA-like proteins. The P. putida LlpABW
sequence adopting a b-hairpin fold is delineated in Figure S1. The
preceding conserved tryptophan residue is located C-terminally to
IC (Figure S1). The sequence logo representation visualizes the
degree of consensus for each residue. Accession numbers are listed
in Figure S4.
(TIF)
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Figure S6
Electron density for (A) Methyl-a-D-Man, (B)
Mana(1–2)Man
and
(C)
GlcNAcb(1–2)Mana(1–
3)[GlcNAcb(1–2)Mana(1–6)]Man. Difference electron-density
maps are calculated by removing the sugar residues from the final
coordinates and applying one round of slow-cool simulated
annealing refinement to remove potential bias. The atomic model
is superimposed in each case.
(JPG)
Figure S7
Mannose binding to LlpABW and garlic lectin.
(A) Cartoon representation of subdomain IIIC of LlpABW (green)
with residues implicated in carbohydrate binding showing in ball-
and-stick representation and labeled (carbon green, oxygen red,
nitrogen blue). The Me-Man residue is shown in red. Selected
hydrogen bonds are shown as black dotted lines. (B) Stereoview of
the superposition of subdomain IIIC of LlpABW (green) on the
equivalent subdomain of garlic lectin (blue). The Me-Man residue
bound to LlpABW is shown in red, the mannose bound to garlic
lectin in blue. (C) Stereoview of the superposition of subdomain
IIIC of LlpABW (green) identical as in panel A, but emphasizing the
location of Val177 (shown as black sticks). The modeled
Val177Tyr mutation is shown as orange sticks. Tyr177 makes a
steric clash with the bound mannose (red) and is therefore
expected to prevent binding, in agreement with our ITC
experiments.
(JPG)
Figure S8
Sites II and I of the LlpABW C-domain. (A)
Stereoview of site IIC of the C-domain (colored according to
atom type) superimposed on site IIIC of the C-domain (dark
gray). The Me-Man bound in site IIIC is shown in red. This site is
very similar to site IIIC but in the crystal it is inaccessible due to
crystal lattice interactions. Residue labels correspond to residues
of site IIC. (B) Similar view showing site IC of the C-domain
(colored according to atom type) superimposed on site IIIC of the
C-domain (dark gray). The Me-Man bound in site IIIC is shown
in red. The stretch of Ile137-Leu139 that provides a steric conflict
preventing Me-Man binding in site IC, is highlighted with carbon
atoms drawn in green. Residue labels correspond to residues of
site IC.
(JPG)
Figure S9
Sites of the LlpABW N-terminal domain. (A)
Stereoview of site IN of the N-domain (colored according to atom
type) superimposed on site IIIC of the C-domain (dark gray). The
Me-Man bound in site IIIC is shown in red. The stretch of Ile271-
Trp274 that provides a steric conflict preventing Me-Man
binding in site IN is highlighted with carbon atoms drawn in
green. Residue numbering corresponds to residues of site IN. (B)
Similar superposition for site IIN of the N-domain. Phe86 that
prevents Me-Man binding to this site through a steric conflict is
highlighted in green. Other residues belonging to site IIN are
labeled in teal. Three residues of site IIIC for which site IIN has
no structural equivalent are labeled in black. (C) Similar
superposition for site IIIN of the N-domain. Residues belonging
to site IIIN are labeled in teal. One residue of site IIIC for which
site IIIN has no structural equivalent, is labeled in black. For this
site there are no obvious steric conflicts that would prevent
positioning of a Me-Man residue although none is observed
experimentally.
(JPG)
Figure S10
Quantification of live and dead cells by flow
cytometry. P. syringae GR12-2R3 cells were treated with LlpA
(A), LlpAV177Y (B), LlpAV208Y (C), LlpAV177Y-V208Y (D), or buffer
(E) at a final concentration of 50 mg/ml for 1 h at 20uC. After
live/dead staining, cells were analysed by flow cytometry. Data
processing allowed to distinguish populations of dead (left) and live
(right) cells. Spot densities ranging from high to low are
differentiated by a color gradient from red, yellow, green, teal to
blue. Representative samples for LlpA, mutant proteins and buffer
control are shown in panels A–E.
(TIF)
Figure S11
Overview of inactive LlpABW deletants and
inactive LlpABW/LlpA1 chimers. The equivalent domains of
LlpA1 are delineated based on pairwise sequence alignment with
LlpABW: N-domain (orange), C-domain (purple), C-terminal
extension (grey). No bacteriocin activity was conferred by these
constructs upon recombinant E. coli cells tested against P. syringae
GR12-2R3 (indicator strain for native LlpABW) and P. fluorescens
LMG 1794 (indicator strain for native LlpA1). The small black
rectangle represents an artificial linker sequence (DASRS).
(TIF)
Table S1
Glycan array profile of LlpABW as measured
by fluorescence intensity. Results including a comprehensive
list of oligosaccharides (array version PA_v5) are available from
the
Consortium
of
Functional
Glycomics
(CFG,
www.
functionalglycomics.org).
(XLS)
Table S2
Bacterial strains and plasmids used in this
study.
(DOC)
Table S3
PCR primers used in this study.
(DOCX)
Table S4
Protein sequences of LlpABW deletants and
LlpABW/LlpA1 chimers.
(DOCX)
Table S5
Structure determination and refinement.
(DOCX)
Acknowledgments
The authors acknowledge the Consortium for Functional Glycomics for
performing glycan array tests on LlpABW, and the use of the
macromolecular crystallography beamlines at EMBL/DESY (Hamburg,
Germany), ESRF (Grenoble, France) and SOLEIL (Gif-sur-Yvette, France)
for X-ray data collection.
Author Contributions
Conceived and designed the experiments: MGKG AGP RL RDM.
Performed the experiments: MGKG AGP EKML SS. Analyzed the data:
MGKG AGP EKML SS RL RDM. Contributed reagents/materials/
analysis tools: MGKG AGP EKML SS RL RDM. Wrote the paper:
MGKG AGP RL RDM.
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|
3M7J
|
Crystal structure of the bacteriocin LLPA from pseudomonas sp. in complex with Met-mannose
|
Structural Determinants for Activity and Specificity of
the Bacterial Toxin LlpA
Maarten G. K. Ghequire1., Abel Garcia-Pino2,3., Eline K. M. Lebbe1¤, Stijn Spaepen1, Remy Loris"2,3*,
Rene´ De Mot"1*
1 Centre of Microbial and Plant Genetics, University of Leuven, Heverlee-Leuven, Belgium, 2 Molecular Recognition Unit, Department of Structural Biology, Vlaams
Instituut voor Biotechnologie, Brussel, Belgium, 3 Structural Biology Brussels, Department of Biotechnology (DBIT), Vrije Universiteit Brussel, Brussel, Belgium
Abstract
Lectin-like bacteriotoxic proteins, identified in several plant-associated bacteria, are able to selectively kill closely related
species, including several phytopathogens, such as Pseudomonas syringae and Xanthomonas species, but so far their mode
of action remains unrevealed. The crystal structure of LlpABW, the prototype lectin-like bacteriocin from Pseudomonas
putida, reveals an architecture of two monocot mannose-binding lectin (MMBL) domains and a C-terminal b-hairpin
extension. The C-terminal MMBL domain (C-domain) adopts a fold very similar to MMBL domains from plant lectins and
contains a binding site for mannose and oligomannosides. Mutational analysis indicates that an intact sugar-binding pocket
in this domain is crucial for bactericidal activity. The N-terminal MMBL domain (N-domain) adopts the same fold but is
structurally more divergent and lacks a functional mannose-binding site. Differential activity of engineered N/C-domain
chimers derived from two LlpA homologues with different killing spectra, disclosed that the N-domain determines target
specificity. Apparently this bacteriocin is assembled from two structurally similar domains that evolved separately towards
dedicated functions in target recognition and bacteriotoxicity.
Citation: Ghequire MGK, Garcia-Pino A, Lebbe EKM, Spaepen S, Loris R, et al. (2013) Structural Determinants for Activity and Specificity of the Bacterial Toxin
LlpA. PLoS Pathog 9(2): e1003199. doi:10.1371/journal.ppat.1003199
Editor: Ambrose Cheung, Geisel School of Medicine at Dartmouth, United States of America
Received August 22, 2012; Accepted January 3, 2013; Published February 28, 2013
Copyright: 2013 Ghequire et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was financially supported by FWO Vlaanderen (Research project G.0393.09), by The Onderzoeksraad of the VUB, by VIB and by the Hercules
Foundation. The authors acknowledge support of the European Community - Research Infrastructure Action under the FP6 ‘‘Structuring the European Research
Area Program’’, contract number: RII3-CT-2004-506008. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of
the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: remy.loris@vib-vub.be (RL); rene.demot@biw.kuleuven.be (RDM)
¤ Current address: Laboratory of Toxicology, University of Leuven, Leuven, Belgium.
. These authors contributed equally to this work.
" These authors also contributed equally to this work and should be considered joint senior authors.
Introduction
In most natural settings, complex interactions occur among
microorganisms,
ranging
from
nutritional
co-operation
to
warfare among competitors. Examples of such interplay have
been reported not only between unrelated microorganisms (e.g.
fungi and bacteria [1,2]), but also between distant relatives (e.g.
members of different bacterial genera [3]), and even between
close relatives (e.g. at inter- and intra-species levels [4,5]). A
major strategy in niche colonization is the production of growth
inhibitors or toxins directed at microbial competitors [6]. While
a huge variety of secondary metabolites is used to target
phylogenetically-distant competitors, ribosome-synthesized pep-
tides or proteins are typically active against close relatives.
These protein toxins are collectively referred to as bacteriocins,
and may either be released into the environment or transferred
to the host via specialized contact-dependent delivery systems
[7–9].
Bacteriocins are structurally and mechanistically very diverse.
This is reflected in the bacteriocinogenic potential of the genus
Pseudomonas [10]. Their R- and F-type pyocins are multi-subunit
protein complexes evolutionarily related to contractile tails of
bacteriophages [11–13]. R-pyocins attach to specific lipopolysac-
charide moieties at the cell surface of susceptible cells and insert
their core structure through the cell envelope, causing depolar-
ization of the cytoplasmic membrane [14]. The S-type pyocins of
Pseudomonas aeruginosa share structural and functional features with
Escherichia coli colicins [15]. Following docking onto surface-
exposed targets such as siderophore receptors [16,17], S-pyocins
kill cells
by nucleic
acid
degradation
[10,17], cytoplasmic
membrane damage [18], or inhibition of peptidoglycan synthesis
[19,20]. Putidacin A (or LlpABW), first identified in Pseudomonas
putida BW11M1 [21], represents a class of Pseudomonas-specific
antibacterial proteins not related to any known bacteriocin.
Additional llpA-like genes encoding functional bacteriocins were
identified by genome mining in the biocontrol strain Pseudomonas
fluorescens Pf-5 [22] and in the phytopathogen Pseudomonas syringae
pv. syringae 642 [23]. Identification of this type of protein in two
Xanthomonas pathovars extended its occurrence as a genus-specific
killer protein [23]. The Xanthomonas LlpA precursor is proteolyt-
ically processed by removal of a characteristic Type II secretion
signal peptide, whereas such N-terminal sequence is lacking in
Pseudomonas homologues, indicating that secretory routes may
differ among LlpA producers.
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The amino acid sequence of LlpA suggests the presence of
two related domains belonging to the ‘monocot mannose-
binding lectin’ (MMBL) family [24]. The MMBL domain
consists of a b-prism fold containing three potential carbohy-
drate-binding pockets, each generated by a QxDxNxVxY
sequence (with x, any amino acid), but some sites may be
inactive due to degeneracy of the signature motif [25]. This
domain (Pfam domain: B_lectin - PF01453) was initially
identified in lectins of monocot plants [26,27], but a more
widespread occurrence of MMBL lectins has become evident
and includes representatives in fungi [28,29], slime molds [30],
sponges [31], and fishes [32–34]. The LlpA branch occupies a
unique position among MMBL-domain proteins, harboring
non-eukaryotic representatives and being equipped with the
capacity to kill bacterial cells with bacteriocin-like specificity, a
property not yet demonstrated for other family members [25].
Next to proteins with the LlpA-type tandem-MMBL organiza-
tion, many other predicted MMBL proteins are encoded by
bacterial genomes. Often the MMBL module is embedded in a
larger protein. For one such protein, bacteriocin-like activity
among Ruminococcus species, Gram-positive bacteria colonizing
the rumen, was demonstrated [35].
Here we report on the crystal structure of LlpABW as the
prototype of a novel family of antibacterial proteins and explore
how domain architecture and specific structural elements contrib-
ute to its activity and specificity.
Results
LlpA forms a rigid MMBL tandem
The crystal structure of LlpABW from P. putida BW11M1
(LlpABW) shows it contains two b-prism MMBL domains,
referred to as the N-domain and the C-domain following their
position in the amino acid sequence (Figure 1A,B; Figure S1).
The N-domain spans residues Arg4-Pro135 while the C-domain
encompasses residues Ala136-Gln253. Each domain exhibits
pseudo-threefold symmetry and the corresponding subdomains
will be referred to as IN, IIN, IIIN, IC, IIC and IIIC, respectively
(Figure 1A and Figure S1). Following these two domains, a b-
hairpin extension is formed by residues Pro254-His275 (the
numbering used in this paper corresponds to that of the wild-type
protein without His-tag [21]).
The two-domain architecture reflects the b-strand swapping
that is typical in dimers of single-domain mannose-binding
monocot lectins (Figure 1A,B) [36] and which apparently is
retained after the ancestral fusion or duplication of the two
domains, as is also the case in certain MMBL tandems or
heterodimers from monocots [37,38]. Thus, residues Asp126-
Pro135 from the first MMBL sequence complement the fold of
the C-domain while residues Pro245-Gln253 from the second
MMBL sequence
complement the
fold of
the N-domain.
However, in LlpABW, the relative orientation of both domains
is different compared to what is observed in a canonical MMBL
lectin dimer, such as snowdrop lectin [36], in the heterodimeric
MMBL lectin ASA I from Allium sativum [38], or in the tandem
MMBL SCAfet from Scilla campanulata [37] (Figure 1C and Figure
S2). In contrast to these plant MMBL proteins, the resulting
architecture of LlpABW does not obey pseudo-twofold symmetry
(Figure 1C).
LlpABW is a very rigid molecule. The two monomers present in
the asymmetric unit are essentially identical with a root-mean-
square deviation (RMSD) of 0.34 A˚ for 270 Ca atoms. This
RMSD value does not change significantly when the individual
domains are fitted separately (0.32 A˚ for 120 Ca’s of the N-
domain and 0.22 A˚ for 115 Ca’s of the C-domain), indicating that
the inter-domain orientation is fixed. This stems from three sets of
interactions (Figure 2). Both domains are connected by a two-
stranded anti-parallel b-sheet that is involved in the b-strand
swapping mentioned above and that links both domains. The C-
terminal b-hairpin extension makes extensive contacts, through
hydrophobic and hydrogen bonds, with both domains. Finally, the
stretch Val140-Asp145 of the C-domain makes extensive contacts
with stretch Val115-Asp118 and with the side chains of Ser15 and
Pro32 of the N-domain.
Domains of LlpABW are shaped by differential
evolutionary pressure
A superposition of the Ca-trace of the N- and C-domain of
LlpABW as well as the MMBL domain of snowdrop lectin is shown
in Figure S3. Based on 79 Ca atoms that form the common b-
sheet core of the MMBL domains, the RMSD between the N- and
C-domains of LlpABW is 1.84 A˚ . While the secondary structure
elements of the C-domain are restricted to the three four-stranded
b-sheets of the b-prism fold, the N-domain contains three
additional secondary structure elements (Figure 1A). A three-turn
a-helix (a1) is inserted in the loop between strands b9 and b10,
and sheet IIN contains two additional strands. Strand b69 is
inserted in the loop between strands b6 and b7 and provides an
anti-parallel extension to sheet II (hydrogen bonding to strand b9).
Strand b19 is a short piece of b-strand that is part of the long N-
terminus and forms a parallel extension on the opposite site of
sheet IIN (hydrogen bonding to strand b2), making this b-sheet a
mixed type six-stranded one rather than the canonical four-
stranded anti-parallel sheet.
Despite these additions to the b-prism fold, the common core
of the N-domain more closely resembles that of the well-studied
and highly conserved monocot lectins (e.g. RMSD of 1.35 A˚ with
snowdrop lectin compared to 1.82 A˚ for the C-domain). This
structural divergence is in contrast with the degree of conserva-
Author Summary
In their natural environments, microorganisms compete
for space and nutrients, and a major strategy to assist in
niche colonization is the deployment of antagonistic
compounds directed at competitors, such as secondary
metabolites (antibiotics) and antibacterial peptides or
proteins (bacteriocins). The latter selectively kill closely
related bacteria, which is also the case for members of the
LlpA family. Here, we investigate the structure-function
relationship for the prototype LlpABW from a saprophytic
plant-associated Pseudomonas whose genus-specific tar-
get spectrum includes several phytopathogenic pseudo-
monads. By determining the 3D structure of this protein,
we could assign LlpA to the so-called monocot mannose-
binding
lectin
(MMBL)
family,
representing
its
first
prokaryotic member, and also add a new type of
protective function, as the eukaryotic MMBL members
have been linked with antiviral, antifungal, nematicidal or
insecticidal activities. For the protein containing two
similarly folded domains, we constructed site-specific
mutants affected in carbohydrate binding and domain
chimers from LlpA homologues to show that mannose-
specific sugar binding mediated by one domain is required
for activity and that the other domain determines target
strain specificity. The strategy that evolved for these
bacteriocins is reminiscent of the one used by mammalian
bactericidal proteins of the RegIII family that recruited a C-
type lectin fold to kill bacteria.
Bactericidal Lectin Structure
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Figure 1. Overall structure of LlpABW. (A) Topology diagram of LlpABW. The N-domain is shown in red, the C-domain in blue and the C-terminal
extension in green. The different strands and subdomains are labeled. Domain swapping involves b-strand segments b11b and b22b, which together
with b-strand segments b11a and b22a link both MMBL domains. (B) Cartoon representation of LlpABW with the different domains colored as in panel
A. The bound Me-Man residue is shown as an orange stick representation. (C) Domain orientations of LlpABW compared with the heterodimeric MMBL
ASA I (Allium sativum agglutinin, PDB entry 1KJ1) and tandem MMBL SCAfet (Scilla campanulata fetuin-binding lectin, PDB entry 1DLP). In each case,
the C-domain is shown in the same orientation, highlighting the different relative orientation of the N-domain in LlpABW. Domain-swapped dimers in
homo-oligomeric plant MMBL lectins such as snowdrop lectin have their domain orientation similar to ASA I and SCAfet.
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tion of the carbohydrate-binding motif characteristic of the
monocot lectins (QxDxNxVxY) in each of the three subdomains.
In the N-domains of LlpA homologues, the surface-exposed
motifs III and II are not well conserved and likely lost their
function during evolution. In contrast they seem to be better
conserved in the C-domains (Figure S4). Apparently, the two
MMBL domains of LlpA experienced a differential evolutionary
pressure resulting in different degrees of global and local
(carbohydrate-binding motif) conservation, suggesting distinct
functional roles for each domain.
The C-domain of LlpABW further extends into a b-hairpin
that helps to define the relative orientations of its two MMBL
domains. This b-hairpin is highly bent due to a b-bulge inserted
into its second b-strand (Figure 1B). It is absent in all plant
representatives including tandem MMBL proteins such as
SCAfet (Figure 1C). In bacteria it represents the most divergent
part of LlpA homologues, both in primary sequence and in
length
(Figure
S5).
Most
of
these
C-terminal
extensions
terminate with a phenylalanine residue. This is reminiscent of
the conserved terminal phenylalanine of outer membrane
proteins from Gram-negative bacteria such as PhoE, required
for their translocation to the cell envelope [39]. An equivalent
extension
appears
to
be
absent
in
the
Xanthomonas
and
Arthrobacter sequences (Figure S5).
LlpA is capable of binding mannose-containing
carbohydrates
Subdomains IIC and IIICof LlpABW contain the typical sugar-
binding signature (QxDxNxVxY) of an active MMBL mannose-
binding site (Figure S1 and S4). Soaking crystals of LlpABW with
200 mM methyl-a-D-mannopyranoside (Me-Man) led to clear
electron density of a single Me-Man in site IIIC of each of the
two LlpABW monomers in the asymmetric unit (Figure S6A).
This site comprises the side chains from Gln171, Asp173,
Asn175 and Tyr179, which contribute to hydrogen bond
interactions and the side chains of residues Val177, Asn188,
Gln192 and Ala185, which contribute to van der Waals contacts
with the carbohydrate ligand (Figure 3A, Figure S7A,C). This
architecture is very similar to what is observed for mannose
bound to other MMBL-type lectins such as snowdrop and garlic
lectin (Figure S7B).
Soaks with oligomannoses revealed additional sugar-binding
subsites.
Binding
site
IIIC
accommodates
the
disaccharide
Mana(1–2)Man and the pentasaccharide GlcNAcb(1–2)Mana(1–
3)[GlcNAcb(1–2)Mana(1–6)]Man (Figure S6B,C). In the case of
Figure 2. Domain interactions within LlpABW. (A) The C-terminal hairpin extension (green cartoon) covers the interface between the N-domain
(red surface representation) and the C-domain (blue surface representation). (B) Stereo view of the interactions between loop segments Val140-
Asp145 (cyan) of the C-domain and Val115-Asp118 (yellow) and Ser31-Gln34 (orange) of the N-domain. Other structural elements are colored
according to panel A. (C) Stereo view of the two-stranded b-sheet formed by strands b11a,b and b22a,b that links the N- and the C-domains and gives
rise to domain swapping. Colors according to panel A and B.
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the pentasaccharide, the central reducing mannose is located in
the shallow Me-Man binding site and the two GlcNAcb(1–2)Man
moieties stretch out over the surface making only a few additional
hydrogen bonds or van der Waals contacts (Figure 3B). In the
bound disaccharide, the non-reducing mannose is located in the
Man-Me binding site while the reducing mannose faces the solvent
and does not interact directly with the protein (Figure 3C).
Site IIC of both LlpABW molecules in the asymmetric unit is
involved in crystal packing interactions and the presence of Me-
Man is therefore sterically excluded. All residues that form specific
hydrogen bonds with Me-Man are retained but substitutions occur
for three side chains that provide van der Waals contacts (Figure
S4 and S8A). In contrast, site IC lost the conserved QxDxNxVxY
motif (Figure S4) and is involved in inter-domain contacts and
therefore inaccessible to ligands (Figure S8B).
The putative carbohydrate-binding sites in the N-domain of
LlpABW are less conserved. Similar to the C-domain, site IN is
inaccessible and involved in inter-domain interactions (Figure
S9A). In the IIN subdomain, the canonical mannose-binding motif
QxDxNxVxY is essentially absent, with only the Gln residue of the
motif being conserved as Gln82 (Figure S4). All other donors or
acceptors required for hydrogen bonds with a mannose ligand are
missing. In addition, the presence of Phe86 at the equivalent
position of the expected Val sterically hinders the binding of
mannose (Figure S9B). The potential carbohydrate-binding site on
subdomain IIIN is only partially conserved (Figure S9C) and
contains two relevant substitutions from the canonical signature:
Figure 3. Carbohydrate binding in site IIIC of LlpABW. (A) Stereoview of methyl-a-D-mannopyranoside bound to subdomain IIIC. Methyl-a-D-
mannopyranoside is shown in blue and indicated by M. Residues belonging to the QxDxNxVxY motif and hydrogen bonding to the sugar as well as
Asn188 are labeled. Water molecules bridging protein and carbohydrate are shown in cyan (B) Similar view of the pentasaccharide GlcNAcb(1–
2)Mana(1–3)[GlcNAcb(1–2)Mana(1–6)]Man. The mannose residue occupying the primary binding site is shown in blue and labeled M. The additional
two mannoses (labeled +1 and 21) and two N-acetyl glucosamine residues (labeled +2 and 22) are shown in green. Other colors are as in panel A. (C)
Binding of the disaccharide Mana(1–2)Man. The non-reducing mannose residue occupying the primary binding site is shown in blue and labeled M.
The second, reducing mannose is shown in green. Other colors are as in panel A.
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(1) the Tyr residue of the QxDxNxVxY motif is replaced by the
shorter Gln49, thereby removing the canonical hydrogen bond
between Man O4 and Tyr OH, and (2) a threonine at position
54 which may compensate the hydrogen bond lost due to the
Tyr-to-Gln substitution in the canonical motif. The lack of
electron density at this site in our Me-Man soak nevertheless
indicates that this site does not recognize this ligand or that its
affinity is so low that recognition would only be achieved in the
context of a larger and as yet unidentified mannose-containing
ligand. Alternatively, this putative site may possess specificity for a
different monosaccharide. In order to evaluate this hypothesis, we
soaked LlpABW crystals with D-galactose, N-acetyl-D-glucosamine
and L-fucose. No electron density was observed for any of these
sugars, suggesting that the N-domain has a function distinct from
carbohydrate recognition (data not shown).
Carbohydrate-binding capacity is required for LlpA
toxicity
The LlpABW motifs IIIN, IIIC and IIC create potential
carbohydrate binding sites that may be involved in bacteriotoxicity
of the protein. We therefore examined the role of carbohydrate
binding in the bactericidal function of LlpABW. The presence of
methyl-a-D-mannopyranoside up to 500 mM in the medium did
not influence the activity of LlpABW on P. syringae GR12-2R3.
Glycan array profiling did not highlight any specific oligosaccha-
ride structure that could represent a natural ligand of LlpABW
(Table S1). This could be due to the array design that is principally
based on eukaryotic glycans and may therefore lack an appropri-
ate carbohydrate for this prokaryotic toxin. Previously, it was
observed that LlpABW from concentrated culture supernatant does
not agglutinate rabbit red blood cells, nor binds to a mannose-
agarose affinity matrix [21].
To assess whether the mannose-recognizing QxDxNxVxY
motifs in LlpABW are nevertheless relevant for bactericidal activity,
the
conserved
valine
residue
was
mutated
to
tyrosine
in
subdomains IIIN, IIIC, and IIC. These mutations sterically
preclude mannose or any other ligand to enter the binding sites
(Figure S7C). Semi-quantitative activity assays with permeabilized
E. coli cells expressing the LlpA variants in motifs IIIN, IIIC and IIC
were used to assess the relationship between carbohydrate binding
and bactericidal activity. Modification of the IIIN site, for which no
mannose binding was observed, does not affect the antibacterial
activity against P. syringae GR12-2R3 (Figure 4). In contrast, the
altered IIIC pocket strongly diminishes activity, either alone or in
pairwise combination with the other mutated sites (IIIN or IIC). A
minor negative effect of the IIC mutation is only apparent in a
double mutant, when combined with a modified IIIN motif.
Purified proteins were prepared to further quantify these effects.
Far UV CD spectra of these mutant forms are identical to that of
native protein LlpABW, indicating that the mutations do not affect
the overall structure of the protein. Isothermal titration calorim-
etry (ITC) showed that LlpABW has an affinity of 2.1 mM for the
pentasaccharide
GlcNAcb(1–2)Mana(1–3)[GlcNAcb(1–2)Ma-
na(1–6)]Man, the highest among all the tested oligo-mannosides
(See Figure 5 and Table 1 for a summary of the experimentally
validated LlpABW-carbohydrate interactions). This is in agreement
with the crystal structures of the different complexes since this
sugar is the one with the largest binding interface (Figure 3).
Titrations of LlpABW, of the mutants LlpAV177Y (a site IIIC
knockout), LlpAV208Y (a site IIC knockout) and of the double
mutant LlpAV177Y-V208Y with a-methyl mannoside clearly pin-
point site IIIC as the only responsible for the sugar binding activity.
Point mutations in both sites or IIIC (V177Y) alone, completely
abrogate sugar binding. However knocking out site IIC (V208Y)
has little effect in binding and the affinities of LlpAV208Y for a-
methyl mannoside and Mana(1–3)Man are very close to the ones
measured for the wild-type protein (See Table 1 and Figure 5B).
While the V208Y mutation in the IIC site has no observable
effect on the MIC value for P. syringae GR12-2R3, the altered IIIC
motif engenders a 5.2-fold increase in MIC (Figure 4). The mutant
protein LlpAV177Y-V208Y suffers a further reduction in activity,
yielding a 31.6-fold increased MIC compared to native LlpABW.
The biological activities of LlpA and its mutants were further
assessed by live/dead staining and subsequent flow cytometry
analysis (Figure 6, Figure S10). Proportions of dead cells after
1 hour of exposure to LlpA or LlpAV208Y were comparable (10.1%
and 9.7%, respectively). For LlpAV177Y this value was reduced to
6.1%, significantly lower than for LlpA. Killing activity was even
further reduced for LlpAV177Y-V208Y (3.7%). These results are
consistent with the MIC determination and ITC data, indicating
that an active site IIIC is required to generate a fully active LlpA
bacteriocin. The difference in bacteriotoxicity between LlpAV177Y
and LlpAV177Y-V208Y suggests that site IIC has a supporting role in
the LlpABW bacteriotoxicity.
All domains are necessary for LlpABW functionality
The site-directed mutagenesis approach revealed an important
role for the C-domain’s carbohydrate-binding capacity in LlpABW
toxicity. Considering the increased binding motif degeneration in
the N-domain and the fact that a Ruminococcus bacteriocin
composed of only a single MMBL domain fused to an unknown
domain has been identified [35], the N-domain may fulfill a
distinct function, different from that of the C-domain. In order to
scrutinize the contribution of individual domains to overall
activity, six domain deletion constructs of llpABW were engineered
to potentially encode proteins lacking the first or second MMBL
domain, a gene product devoid of the C-terminal hairpin, or a
protein retaining only an individual domain (N-domain, C-
domain, or hairpin) (Figure S11). To take the domain swapping
into account, the constructs containing only a single MMBL
domain were designed with a fusion of the swapped C-terminal b-
strands to the corresponding domain via a short linker.
None of these deletion constructs resulted in the production of
an active protein, indicating that none of the domains are
dispensable. Removal of the terminal phenylalanine residue still
allows expression of a functional bacteriocin in E. coli (Figure 7),
but a further C-terminal truncation (deletion of Trp-His-Phe tail)
resulted in a negative bacteriocin assay (data not shown). From
these data we conclude that both MMBL domains as well as the
C-terminal hairpin extension are required for activity of LlpA.
Whether the role of the C-terminal hairpin is any other than
simply stabilization of the C-domain cannot be concluded.
Target specificity of LlpA is hosted by the N-domain
In order to investigate the role of the different domains in target
specificity, we created hybrid LlpA proteins using the domains of
LlpABW from P. putida BW11M1 and LlpA1 from P. fluorescens Pf-5.
These two LlpA proteins share 45% sequence identity and differ in
their target spectra. Strains P. syringae GR12-2R3 and P. fluorescens
LMG1794 were identified as specific indicators for LlpABW [21]
and LlpA1 [22], respectively. Six constructs carrying llpABW/
llpA1chimeric genes were made with domain exchanges involving
the N-domain, C-domain, and hairpin region (Figure 7 and Figure
S11). For four of these constructs activity against one of both
indicators was detected. Only constructs retaining the original N-
domain give rise to inhibition of the cognate indicator strain. The
C-domain or the hairpin of LlpABW could be replaced with the
corresponding LlpA1 domains without changing target specificity.
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Conversely, the original specificity of LlpA1 is retained upon
replacement of its C-domain by the LlpABW equivalent.
Discussion
Structure elucidation of LlpABW from P. putida BW11M1
unequivocally assigns this bacteriocin to the MMBL lectin family,
in which it constitutes the first prokaryotic member, representative
for a group of bacterial proteins composed of two MMBL domains
[21–23]. Systematic inactivation of the three potential carbohy-
drate-binding sites present in the N-domain (IIIN) and in the
C-domain (IIIC and IIC) of LlpABW, revealed that a non-occluded
IIIC pocket is required to obtain a fully active LlpABW molecule. A
negative co-operative effect on activity resulted when the IIC site
was additionally modified. Although mannose-containing carbo-
hydrates can bind to the IIIC pocket of LlpABW, it remains unclear
Figure 4. Inhibitory activity of wild-type LlpABW and selected mutants with modified (potential) mannose-binding sites. The domain
structure (N-domain in red, C-domain in blue and C-terminal extension in green) and the position of the MMBL motifs (potentially active binding sites
in orange, inactive ones in grey) are shown. The positions of conserved valine residues converted to tyrosine residues by site-directed mutagenesis
are indicated with a black bar. Inhibitory activity of E. coli strains expressing mutant LlpABW forms was assayed against P. syringae GR12-2R3 and semi-
quantified according to the size (inner zone radius) of the growth inhibition halo relative to LlpABW (+++, native LlpABW; ++, halo size reduced; + halo
size strongly reduced; 2, no halo; NT, not tested). For wild-type LlpABW and three purified His-tagged mutant forms (LlpAV177Y, LlpAV208Y and
LlpAV177Y-V208Y) the MIC values were determined with indicator P. syringae GR12-2R3. Molar minimal inhibitory concentrations of recombinant
proteins (with standard deviations): LlpA, 2.08 nM (60.58 nM); LlpAV177Y, 10.9 nM (60.66 nM); LlpAV208Y, 1.98 nM (60.066 nM); 65.72 nM
(62.80 nM).
doi:10.1371/journal.ppat.1003199.g004
Table 1. Binding affinities and thermodynamic parameters obtained from ITC titrations.
Type of protein-carbohydrate
interaction
Kd (mM)
DG6(kcal mol21)
DH6 (kcal mol21)
2TDS6(kcal mol21)
LlpABW Me-a-D-Man
45.9
21.8
25.4
3.6
LlpABW Mana(1–2)Man
42.4
21.9
23.6
1.7
LlpABW Mana(1–3)Man
18.2
22.4
25.9
3.5
LlpABW Mana(1–6)Man
17.2
22.4
25.5
3.1
LlpABW Mana(1–3)[Mana(1–6)]Man
10.1
22.6
26.4
3.8
LlpABW GlcNAcb(1–2)Mana(1–3)[GlcNAcb(1–
2)Mana(1–6)]Man
2.1
23.7
21.6
22.1
LlpAV208Y Me-a-D-Man
58.8
21.7
23.3
1.6
LlpAV208Y Mana(1–3)Man
23.0
22.2
25.1
2.9
The reported values for Kd, DGu, DHu and 2TDSu were determined from fitting a single site interaction model (n = 1) to the experimental ITC data. The interaction of the
mutants LlpAV177Y and LlpAV177-V208Y with the different sugars is negligible and no heat effect was observed. Therefore they are not included in this table.
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if these are part of or mimic the natural ligand required for
biological activity since bacteriocin activity is not impaired in the
presence of excess mannose.
A mutated IIIN site did not provoke a negative effect on
antibacterial activity. However, the N-domain appears to play a
major role in target selection. This was demonstrated by
assessing the differential activity of domain chimers against
two target strains, diagnostic for the LlpABW- and LlpA1-
specific killing.
The b-hairpin does not appear to be a specificity determi-
nant, although it constitutes the most variable region among
LlpA-like bacteriocins. Possibly, it is required for thermody-
namic stability since it needs to be intact in LlpABW. An
equivalent C-terminal stretch is absent from the Xanthomonas citri
LlpA-like
bacteriocin
[23].
From
our
results
relying
on
heterologous expression in E. coli and a bacteriocin assay with
permeabilized cells, it cannot be excluded that this structural
element may play a role in the way an LlpA protein is exported
by its native producer cells.
Figure 6. Killing activity of LlpABW and mutant proteins. Percentages of dead cells after live/dead staining as quantified by flow cytometry
analysis (Figure S10). P. syringae GR12-2R3 was used as indicator strain and treated at a final concentration of 50 mg/ml for 1 h. Average values (with
standard deviations; indicated by error bars): LlpA, 10.1 (61.04); LlpAV177Y, 6.1 (60.44); LlpAV208Y, 9.7 (61.39); LlpAV177Y-V208Y, 3.7 (60.90); buffer
(control), 1.0 (60.11).Values are significantly different for (a) and (b), (b) and (c) (p,0.01).
doi:10.1371/journal.ppat.1003199.g006
Figure 5. ITC analysis of carbohydrate binding to LlpABW and mutants. (A) Binding of LlpABW to the pentasaccharide GlcNAcb(1–2)Mana(1–
3)[GlcNAcb(1–2)Mana(1–6)]Man. (B) Binding of LlpABW (blue circles, wild type) and the mutants LlpAV177Y (green circles, site IIIC knockout), LlpAV208Y
(red circles, site IIC knockout) and LlpAV177Y-V208Y (black circles, site IIC and IIIC knockout) to a-methyl mannoside. There is no heat exchanged in the
titration of the double mutant or the site IIIC knockout LlpAV177Y, whereas the site IIC knockout LlpAV208Y, binds the monosaccharide in a ‘‘wildtype’’-
like fashion, showing that only site IIIC is involved in sugar binding.
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In general, a defensive role has been proposed for the
(oligo)mannose-binding MMBL lectins based on insecticidal,
nematicidal, antifungal, or even antiviral activities demonstrated
for several of these proteins that are abundantly found in monocot
plants [40–46]. Some of these plant lectins trigger apoptosis in
cancer cells [47]. Also their identification in fish mucus and
epithelial cells is in line with a general protective (antimicrobial)
function for MMBL domains [32]. LlpABW as a bactericidal
protein fits within this picture of MMBL domains being involved
in general defense mechanisms. Since no antibacterial activity has
been assigned to the eukaryotic MMBL proteins, it is challenging
to identify structural features that confer the intragenus-specific
bacteriocin activity of LlpA, as shown for proteins from P. putida
[21], P. fluorescens [22], P. syringae [23], and Xanthomonas citri [23].
Their
target
spectra
are
narrower
than
reported
for
the
mammalian antibacterial C-type lectins of the RegIII family, such
as mouse RegIIIc and its human homolog HIP/PAP that bind to
the surface-exposed peptidoglycan layer of Gram-positive bacteria
[48], and RegIIIb that also binds to the lipid A moiety of
lipopolysaccharides
on
the
cell
envelope
of
Gram-negative
bacteria [49].
The absence of any known secretory signal sequence in
LlpABW and its homologues in other Pseudomonas species is
intriguing in view of their extracellular location [21]. The
translocation of the outer membrane-associated mannose/
fucose-specific lectin LecB of P. aeruginosa, that also lacks such
signal sequence [50], is dependent on its glycosylation [51].
Contrary to LlpA that is exported to the culture supernatant to
exert its antagonistic activity, LecB remains associated with the
cell
envelope
through
interaction
with
the
major
outer
membrane protein OprF [52], in line with its role in biofilm
formation.
Materials and Methods
Strains and culture conditions
Bacterial strains and plasmids used in this study are listed in
Table S2. Escherichia coli was routinely grown in shaken Luria-
Bertani (LB, MP Biomedicals) broth at 37uC. Pseudomonas strains
were grown in Tryptic Soy Broth (BD Biosciences) at 30uC with
shaking. Media were solidified with 1.5% agar (Invitrogen) and
supplemented with filter-sterilized antibiotics as required at
following concentrations: ampicillin (Sigma-Aldrich), 100 mg/ml
or kanamycin (Sigma-Aldrich), 50 mg/ml. Isopropyl b-D-thioga-
lactoside (IPTG 40 mg/ml, ForMedium) and 5-bromo-4-chloro-3-
indolyl-b-D-galactopyranoside (X-Gal 40 mg/ml, ForMedium)
were added for blue/white screening of pUC18-derived plasmids
in E. coli.
Plasmids used for antibacterial testing and sequencing were
propagated in E. coli TOP10F9 (Invitrogen). E. coli BL21(DE3)
(Novagen) was used as a host for plasmids driving recombinant
protein expression. Genomic DNA from P. putida BW11M1 was
isolated using the Puregene Yeast/Bact. Kit B (Qiagen). Plasmid
DNA was extracted using the QIAprep Spin Miniprep Kit
(Qiagen). Stocks were stored at 280uC in the appropriate medium
in 25% (v/v) glycerol.
Figure 7. Differential inhibitory activity of wild-type LlpABW and LlpA/LlpA1 domain chimers. The domain structures of LlpABW (as in
Figure 4) and of LlpA1 (inferred by pairwise alignment; N-domain in orange, C-domain in purple and C-terminal extension in grey) are depicted, along
with those of chimeric forms (in dashed box). The LlpA variant lacking the terminal phenylalanine residue is marked with a yellow hexagon. Inhibitory
activity of the respective E. coli recombinants was tested with diagnostic indicators for LlpABW (P. syringae GR12-2R3) and LlpA1 (P. fluorescens LMG
1794). Halo sizes are semi-quantified according to size of the growth inhibition halo (+++, native halo size of LlpABW and LlpA1; ++, halo size reduced;
C, local clearing confined to producer colony spot; 2, no halo or clearing; NT, not tested). Additional chimeric and domain deletion constructs not
conferring bacteriocin activity against one of the indicator strains are specified in Figure S11.
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Recombinant DNA methods
Standard methods were used for preparation of competent E.
coli cells, heat shock transformation of E. coli and DNA
electrophoresis [53]. Restriction enzymes were used according to
the supplier’s specifications (Roche Diagnostics and BIOKE´).
DNA ligation was performed using T4 DNA ligase (Invitrogen).
Plasmid sequencing was performed by GATC Biotech (Constance,
Germany). Constructs that were generated are listed in Table S2
and primers are listed in Table S3.
A 921-bp fragment containing llpABW was amplified by PCR
with Platinum Pfx DNA polymerase (Invitrogen), using a C1000
Thermal Cycler (Bio-Rad). P. putida BW11M1 genomic DNA
was taken as a template, and combined with primers PGPRB-
3155 and PGPRB-3156. The amplicon was purified using the
QIAquick PCR Purification Kit (Qiagen), digested with KpnI
and BamHI, ligated in pUC18, and transformed to E. coli
TOP10F9. Transformants were verified for the presence of the
insert by PCR using Taq Polymerase (BIOKE´ ) with primers
PGPRB-2545
and
PGPRB-2546.
The
cloned
construct
(pCMPG6129)
was
purified
and
its
insert
confirmed
by
sequencing.
DpnI-mediated site-directed mutagenesis was performed to
construct valine-to-tyrosine mutant forms of llpABW in pUC18
(pCMPG6129) and N-terminal His-tagged llpABW in pET28a
(pCMPG6056 [54]). PCR conditions were: 2 min initial denatur-
ation, followed by 16 cycles of denaturation (1 min), annealing
(1 min, primer-dependent temperature) and elongation at 68uC
(1 min./kb). Final elongation was for 8 min at 68uC. After PCR,
samples were immediately treated with DpnI at 37uC for 2 h and
transformed into E. coli TOP10F9 and selected on the appropriate
medium. Plasmid inserts of selected transformants were verified by
sequence
analysis. Double
mutants
were
constructed
using
plasmids with a single point mutation as a template.
Domain
deletants
of
llpABW
were
constructed
using
pCMPG6129 as a template (llpABW from P. putida BW11M1).
Chimeric constructs were obtained using pCMPG6129 and
pCMPG6053 (llpA1 from P. fluorescens Pf-5 [22]) as templates.
Artificial ligation of gene fragments, generated with the PCR
primers specified in Table S3, was performed by using splicing by
overlap extension (SOE). The resulting recombinant amino acid
sequences are listed in Table S4.
Recombinant protein expression and purification
Protein isolation and purification of N-terminal His6-tagged
LlpABW, LlpAV177Y, LlpAV208Y, and LlpAV177Y-V208Y from E. coli
BL21(DE3),
carrying
expression
constructs
pCMPG6056,
pCMPG6149, pCMPG6150 and pCMPG6151 respectively, were
performed as described by Parret and collaborators [54]. The
presence of His-tagged protein was observed via immunodetection
by Western blot, using monoclonal anti-His6 (IgG1 from mouse;
Roche Diagnostics) as primary antibody. Fractions free of other
proteins, as verified by SDS-PAGE and subsequent Coomassie
Blue staining, were dialyzed against bis-TRIS propane buffer
(20 mM, 200 mM NaCl, pH 7.0). Concentrations of purified
proteins were determined by absorbance measurement at 280 nm
using molar extinction coefficients of 62910 M21 cm21 for
LlpABW, 64400 M21 cm21 for LlpAV177Y and LlpAV208Y, and
65890 M21 cm21 for LlpAV177Y-V208Y. Extinction coefficients
were calculated according to Pace and collaborators [55].
Antibacterial assays
Bacteriocin production by E. coli cells carrying pUC18-derived
constructs was assayed as follows: 2-ml drops of an overnight
stationary-phase culture were spotted onto LB agar plates and
incubated for 8 h at 37uC. Next, plates were exposed to
chloroform vapor (30 min), aerated and subsequently overlaid
with 5 ml of soft agar (0.5%), seeded with 200 ml of a cell culture of
an indicator strain (,108 CFU/ml), followed by overnight
incubation at 30uC. Next day, plates were scored for the presence
of halos in the confluently grown overlay.
Antibacterial activity assays with purified recombinant His6-
tagged proteins were performed as described by Ghequire and
collaborators [23]. To assess the influence of added sugar, the
same assay was carried out on agar medium supplemented with D-
mannose (Sigma-Aldrich) or methyl-a-D-mannopyranoside (Sig-
ma-Aldrich) to a final concentration of 0.01 M to 0.5 M.
A Bioscreen C apparatus (Oy Growth Curves Ab Ltd, Finland)
was used to determine the minimum inhibitory concentration
(MIC). An overnight culture (16 h) of the indicator strain was
diluted to 104–105 CFU/ml and incubated at 30uC, with a two-
fold dilution series of recombinant His6-tagged LlpABW or mutant
LlpABW. Bis-TRIS propane buffer was used as control. The MIC
value was determined as the minimum concentration of protein at
which no growth of the indicator strain (OD600,0.2) occurred
after 24 h. At least three independent repeats, each with three
replicates, were carried out.
Glycan array
His6-tagged LlpABW was lyophilized and verified for antibac-
terial activity. After re-dissolving in MilliQ water, recombinant
LlpABW was diluted to 200 mg/ml with binding buffer (20 mM
TRIS-HCl pH 7.4, 150 mM NaCl, 2 mM CaCl2, 2 mM MgCl2,
0.05% Tween 20, 1% BSA), and used to probe the printed glycan
arrays [56] following the standard procedures of Core H of the
Consortium
for
Functional
Glycomics
(http://www.
functionalglycomics.org/).
Monoclonal
anti-His6
antibodies
(Roche Diagnostics) were used as primary antibodies, and
fluorescently labeled anti-mouse IgG as secondary antibodies.
Circular dichroism
CD spectra were acquired on a Jasco J-715 spectropolarimeter.
Curves were averaged over 8 scans, taken at 25uC using a 1 mm
cuvette. Samples were dialyzed against bis-TRIS propane buffer
(20 mM, NaCl 200 mM, pH 7.0), filtered and degassed prior to
data acquisition. All proteins were assayed at 0.4 mg/ml.
Isothermal titration calorimetry
ITC titrations were carried out on an ITC200 apparatus
(MicroCal). Prior to the measurement, LlpABW, LlpAV177Y,
LlpAV208Y and LlpAV177Y-V208Y was dialyzed to bis-TRIS
propane buffer. Sugars were directly dissolved into the same
buffer. The samples were filtered and degassed for 10 min at
25uC before being examined in the calorimeter. The titrations
were carried out at 25uC, injecting the sugars (methyl-a-D-
mannoside, Mana(1–2)Man, Mana(1–3)Man, Mana(1–6)Man,
Mana(1–3)[Mana(1–6)]Man
and
GlcNAcb(1–2)Mana(1–
3)[GlcNAcb(1–2)Mana(1–6)]Man) into a protein solution (pro-
tein concentrations ranged from 2 mM to 4 mM depending on
protein availability). All data were analyzed using the MicroCal
Origin ITC 7.0 software. Binding affinities and thermodynamic
parameters from all ITC titrations are reported in Table 1.
X-ray data collection and structure determination
Expression, purification and crystallization of recombinant His-
tagged LlpABW have been described [54]. X-ray data for native and
derivative crystals were collected on EMBL beamline BW7A of the
DESY synchrotron (Hamburg, Germany). For each potential
Bactericidal Lectin Structure
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derivative, the wavelength was chosen to be at the high-energy side
of the absorption edge in order to ensure a usable anomalous signal.
All data were scaled and merged with the HKL package of
programs. Data collection statistics are given in Table S5.
The crystal structure of free LlpABW was solved combining
single
isomorphous
replacement
with
anomalous
scattering
(SIRAS strategy) from a p-chloromercurybenzoate derivative.
The heavy-atom substructure was determined with SHELXD
[57] using a resolution cutoff of 4.0 A˚ . Heavy-atom refinement
and phasing were performed with SHARP [58]. Phase improve-
ment by solvent flattening was performed with SOLOMON [59].
Non-crystallographic symmetry averaging with density modifica-
tion [60] further improved the electron density. A partial model
(94% of the residues comprising the asymmetric unit) was
automatically built with ARP/wARP [61] and the remainder
was built manually over several cycles of model building with Coot
[62], alternated with refinement using phenix.refine [63,64]. Phasing
and refinement statistics are shown in Table S5.
Carbohydrate soaks
Crystals of LlpABW were transferred to artificial mother liquor
(0.1 M imidazole pH 6.5, 1.3 M sodium acetate) enriched with
either 200 mM methyl-a-D-mannopyranoside (Me-Man), Ma-
na(1–2)Man,
GlcNAcb(1–2)Mana(1–3)[GlcNAcb(1–2)Mana(1–
6)]Man (M592), D-galactose, L-fucose or N-acetyl-D-glucosamine
and allowed to equilibrate overnight (all carbohydrates obtained
from Dextra Laboratories, Reading, U.K.). Data were collected at
room temperature on EMBL beamline X13, except for the N-
acetyl-D-glucosamine soak collected at the PROXIMA-1 beam-
line of the SOLEIL synchrotron (Gif-sur-Yvette, France) and the
D-galactose soak collected at ESRF beam line ID14-1 (Grenoble,
France). All data were scaled and merged using the HKL package.
Refinement was started from the coordinates of the ligand-free
structure using phenix.refine. Manual rebuilding, including the
introduction of the carbohydrate ligand if present, was done with
Coot [62]. Crystal structures of LlpABW from P. putida BW11M1
(PDB entry 3M7H) and LlpABW in complex with methyl-a-D-
mannoside (PDB entry 3M7J), with Mana(1–2)Man (PDB entry
4GC1), and with GlcNAcb(1–2)Mana(1–3)[GlcNAcb(1–2)Ma-
na(1–6)]Man (PDB entry 4GC2) have been deposited at the PDB.
Flow cytometry
Overnight cultures of P. syringae GR12-2R3 (16 h) were diluted
to OD600 0.5 and washed twice with phosphate-buffered saline
(PBS). Cells were treated with LlpA, mutant proteins or buffer (bis-
TRIS propane buffer, negative control), at a final concentration of
50 mg/ml for 1 h, at 20uC. Next, PBS-washed bacteria were
stained using the Live/Dead BacLight bacterial viability kit
(Invitrogen), incubated for 15 minutes, and analyzed on a BD
Influx (BD Biosciences). Excitation of the dyes was done at
488 nm, and fluorescence measured at 530 nm for SYTO 9 and at
610 nm for propidium iodide. Results were processed with FlowJo
10.0.4 software (Figure S10). Measurements were done indepen-
dently and based on six biological repeats. Results are expressed as
percentages of dead cells [dead/(live+dead) * 100)].
Supporting Information
Figure
S1
Amino acid sequence of LlpABW colored
according to its domain structure. The N-domain is shown
in red, the C-domain in blue and the C-terminal extension in green.
Residues belonging to sequences equivalent to the mannose binding
site signature motif QxDxNxVxY are in bold and underlined.
(JPG)
Figure S2
Quaternary structures and domain organi-
zation of various MMBL family members. Individual
domains or protomers are shown in different colours. The
domain or protomer colored green (which in the tandem MMBLs
of LlpA, ASA I and SCAfet corresponds to the N-terminal
domain) is always shown in the same orientation. Bound
carbohydrates are shown in black stick representation. For LlpA
a single pentasaccharide is bound to site IIIC. In the case of
Galanthus nivalis (snowdrop) lectin (PDB entry 1JPC), twelve
trimannosides are bound to all QxDxNxVxY motifs (three on
each monomer of the homotetrameric protein). The snowdrop
lectin tetramer consists of the association of two domain-swapped
dimers (green-blue and pink-yellow). In the case of Allium sativum
(garlic lectin ASA I - PDB entry 1KJ1), again each QxDxNxVxY
motif has a dimannose bound while an additional sugar (shown in
red) is bound to a non-canonical site. The protein is synthesized
as a single chain precursor and post-translationally cleaved into
two MMBL domains that adopt the same domain-swapped dimer
as found in snowdrop lectin. Gastrodianin is a monomeric
MMBL family member from the orchid Gastrodia elata (PDB entry
1XD5). The location(s) of its carbohydrate-binding site(s) is (are)
not known. The fetuin-binding tandem-MMBL SCAfet from
Scilla campanulata (PDB entry 1DLP) consists of two covalently
attached MMBL domains, whereas in LlpA the swap of the C-
terminal b-strands is retained. The relative orientation in the two
domains is as in ASA I. This lectin binds fetuin rather than
oligomannosides, but the locations of the binding sites are not
known.
(JPG)
Figure S3
Stereo view of the superpositions (Ca repre-
sentations) of the N-domain of LlpABW (red), C-domain
of LlpABW (blue) and Galanthus nivalis lectin (PDB entry
1MSA, black). The superposition is shown in two orientations
rotated by 90u.
(JPG)
Figure S4
Sequence alignment of potential mannose-
binding motifs in prokaryotic tandem MMBL proteins.
The
sequences
corresponding
to
the
consensus
motif
QxDxNxVxY, extracted from the N-domain and the C-domain
of P. putida LlpABW and its homologues, are aligned per domain.
Sequence conservation is visualized by differential shading. The
sequence logo representation visualizes the degree of consensus for
each residue. LlpA proteins with proven bacteriotoxic activity are
labeled with an asterisk. Accession numbers: Arthrobacter sp. FB24
(YP_829274), Burkholderia ambifaria MEX-5 (ZP_02905572), Burk-
holderia cenocepacia AU 1054 ([1], ABF75998; [2], ABF75999),
Pseudomonas
chlororaphis
subsp.
aureofaciens
30–84
(EJL08681),
Pseudomonas putida BW11M1 (AAM95702), Pseudomonas fluorescens
Pf-5 (LlpA1 [1], YP_258360; LlpA2 [2], YP_259234), Pseudomonas
sp. GM80 ([1], ZP_10606046; [2], ZP_10606131), Pseudomonas
syringae pv. aptata DSM 50252 (EGH77666), Pseudomonas syringae pv.
syringae 642 (ZP_07263221), Xanthomonas axonopodis pv. citri str. 306
(AAM35756).
(TIF)
Figure S5
Sequence alignment of the carboxy-terminal
sequences of LlpA-like proteins. The P. putida LlpABW
sequence adopting a b-hairpin fold is delineated in Figure S1. The
preceding conserved tryptophan residue is located C-terminally to
IC (Figure S1). The sequence logo representation visualizes the
degree of consensus for each residue. Accession numbers are listed
in Figure S4.
(TIF)
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February 2013 | Volume 9 | Issue 2 | e1003199
Figure S6
Electron density for (A) Methyl-a-D-Man, (B)
Mana(1–2)Man
and
(C)
GlcNAcb(1–2)Mana(1–
3)[GlcNAcb(1–2)Mana(1–6)]Man. Difference electron-density
maps are calculated by removing the sugar residues from the final
coordinates and applying one round of slow-cool simulated
annealing refinement to remove potential bias. The atomic model
is superimposed in each case.
(JPG)
Figure S7
Mannose binding to LlpABW and garlic lectin.
(A) Cartoon representation of subdomain IIIC of LlpABW (green)
with residues implicated in carbohydrate binding showing in ball-
and-stick representation and labeled (carbon green, oxygen red,
nitrogen blue). The Me-Man residue is shown in red. Selected
hydrogen bonds are shown as black dotted lines. (B) Stereoview of
the superposition of subdomain IIIC of LlpABW (green) on the
equivalent subdomain of garlic lectin (blue). The Me-Man residue
bound to LlpABW is shown in red, the mannose bound to garlic
lectin in blue. (C) Stereoview of the superposition of subdomain
IIIC of LlpABW (green) identical as in panel A, but emphasizing the
location of Val177 (shown as black sticks). The modeled
Val177Tyr mutation is shown as orange sticks. Tyr177 makes a
steric clash with the bound mannose (red) and is therefore
expected to prevent binding, in agreement with our ITC
experiments.
(JPG)
Figure S8
Sites II and I of the LlpABW C-domain. (A)
Stereoview of site IIC of the C-domain (colored according to
atom type) superimposed on site IIIC of the C-domain (dark
gray). The Me-Man bound in site IIIC is shown in red. This site is
very similar to site IIIC but in the crystal it is inaccessible due to
crystal lattice interactions. Residue labels correspond to residues
of site IIC. (B) Similar view showing site IC of the C-domain
(colored according to atom type) superimposed on site IIIC of the
C-domain (dark gray). The Me-Man bound in site IIIC is shown
in red. The stretch of Ile137-Leu139 that provides a steric conflict
preventing Me-Man binding in site IC, is highlighted with carbon
atoms drawn in green. Residue labels correspond to residues of
site IC.
(JPG)
Figure S9
Sites of the LlpABW N-terminal domain. (A)
Stereoview of site IN of the N-domain (colored according to atom
type) superimposed on site IIIC of the C-domain (dark gray). The
Me-Man bound in site IIIC is shown in red. The stretch of Ile271-
Trp274 that provides a steric conflict preventing Me-Man
binding in site IN is highlighted with carbon atoms drawn in
green. Residue numbering corresponds to residues of site IN. (B)
Similar superposition for site IIN of the N-domain. Phe86 that
prevents Me-Man binding to this site through a steric conflict is
highlighted in green. Other residues belonging to site IIN are
labeled in teal. Three residues of site IIIC for which site IIN has
no structural equivalent are labeled in black. (C) Similar
superposition for site IIIN of the N-domain. Residues belonging
to site IIIN are labeled in teal. One residue of site IIIC for which
site IIIN has no structural equivalent, is labeled in black. For this
site there are no obvious steric conflicts that would prevent
positioning of a Me-Man residue although none is observed
experimentally.
(JPG)
Figure S10
Quantification of live and dead cells by flow
cytometry. P. syringae GR12-2R3 cells were treated with LlpA
(A), LlpAV177Y (B), LlpAV208Y (C), LlpAV177Y-V208Y (D), or buffer
(E) at a final concentration of 50 mg/ml for 1 h at 20uC. After
live/dead staining, cells were analysed by flow cytometry. Data
processing allowed to distinguish populations of dead (left) and live
(right) cells. Spot densities ranging from high to low are
differentiated by a color gradient from red, yellow, green, teal to
blue. Representative samples for LlpA, mutant proteins and buffer
control are shown in panels A–E.
(TIF)
Figure S11
Overview of inactive LlpABW deletants and
inactive LlpABW/LlpA1 chimers. The equivalent domains of
LlpA1 are delineated based on pairwise sequence alignment with
LlpABW: N-domain (orange), C-domain (purple), C-terminal
extension (grey). No bacteriocin activity was conferred by these
constructs upon recombinant E. coli cells tested against P. syringae
GR12-2R3 (indicator strain for native LlpABW) and P. fluorescens
LMG 1794 (indicator strain for native LlpA1). The small black
rectangle represents an artificial linker sequence (DASRS).
(TIF)
Table S1
Glycan array profile of LlpABW as measured
by fluorescence intensity. Results including a comprehensive
list of oligosaccharides (array version PA_v5) are available from
the
Consortium
of
Functional
Glycomics
(CFG,
www.
functionalglycomics.org).
(XLS)
Table S2
Bacterial strains and plasmids used in this
study.
(DOC)
Table S3
PCR primers used in this study.
(DOCX)
Table S4
Protein sequences of LlpABW deletants and
LlpABW/LlpA1 chimers.
(DOCX)
Table S5
Structure determination and refinement.
(DOCX)
Acknowledgments
The authors acknowledge the Consortium for Functional Glycomics for
performing glycan array tests on LlpABW, and the use of the
macromolecular crystallography beamlines at EMBL/DESY (Hamburg,
Germany), ESRF (Grenoble, France) and SOLEIL (Gif-sur-Yvette, France)
for X-ray data collection.
Author Contributions
Conceived and designed the experiments: MGKG AGP RL RDM.
Performed the experiments: MGKG AGP EKML SS. Analyzed the data:
MGKG AGP EKML SS RL RDM. Contributed reagents/materials/
analysis tools: MGKG AGP EKML SS RL RDM. Wrote the paper:
MGKG AGP RL RDM.
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3M7K
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Crystal structure of PacI-DNA Enzyme product complex
|
Unusual target site disruption by the rare-cutting HNH restriction
endonuclease PacI
Betty Shen1, Daniel F. Heiter2, Siu-Hong Chan2, Hua Wang2, Shuang-Yong Xu2, Richard D.
Morgan2, Geoffrey G. Wilson2, and Barry L. Stoddard1,3
1Division of Basic Sciences, Fred Hutchinson Cancer Research Center, 1100 Fairview Ave. N.
A3-025, Seattle, WA 98109, USA
2New England Biolabs, Inc., 240 County Road, Ipswich, MA 01938, USA
Abstract
The crystal structure of the rare-cutting HNH restriction endonuclease PacI in complex with its eight
base pair target recognition sequence 5'-TTAATTAA-3' has been determined to 1.9 Å resolution.
The enzyme forms an extended homodimer, with each subunit containing two zinc-bound motifs
surrounding a ββα-metal catalytic site. The latter is unusual in that a tyrosine residue likely initiates
strand-cleavage. PacI dramatically distorts its target sequence from Watson-Crick duplex DNA
basepairing, with every base separated from its original partner. Two bases on each strand are
unpaired, four are engaged in non-canonical A:A and T:T base pairs, and the remaining two bases
are matched with new Watson-Crick partners. This represents a highly unusual DNA binding
mechanism for a restriction endonuclease, and implies that initial recognition of the target site might
involve significantly different contacts from those visualized in the DNA-bound cocrystal structures.
Restriction endonucleases (REases) occur in all free-living bacteria and archaea and are
believed to function to defend their hosts against invasion by foreign DNA, particularly from
bacteriophage (Pingoud et al., 2005). REases vary in sequence, structure, oligomeric
composition, substrate-specificity, and enzymatic behavior (Bujnicki, 2003). They range from
compact monomers that act independently, to elaborate multifunctional protein assemblages,
and typically recognize target sequences in duplex DNA ranging from four to eight specific
base pairs in length. (Pingoud et al., 2005). These sequences can be symmetric or asymmetric,
as well as continuous or discontinuous, depending upon the enzyme architecture.
Several distinct catalytic site motifs and mechanisms have been identified among restriction
endonucleases, suggesting this enzymatic and biological function has evolved independently
several times. The most common catalytic motif, that of the 'PD…(D/E)xK' nuclease
superfamily, is the the most wide-spread and best understood (Kosinski et al., 2005).
Alternative catalytic motifs, associated with quite different core protein folds, have been
identified in many additional restriction endonucleases, including the ‘HNH’ (Cymerman et
al., 2006; Jakubauskas et al., 2007; Saravanan et al., 2004) and the ‘GIY-YIG’ (Ibryashkina et
al., 2007) motifs (both of which are more commonly associated with mobile homing
endonucleases from bacteriophage) (Stoddard, 2005). All three of these catalytic lineages are
© 2010 Elsevier Inc. All rights reserved.
3Corresponding author, bstoddar@fhcrc.org 1-206-667-4031 (office) -4066 (lab) -3331 (fax).
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also found in a much wider variety of enzymes involved in DNA metabolism and modification,
including those responsible for DNA repair, recombination and fidelity (Cymerman et al.,
2006; Dunin-Horkawicz et al., 2006; Kosinski et al., 2005). As well, isolated examples of two
additional structural motifs (containing the phospholipase D and 'half-pipe' folds) have also
observed for R.BfiI and R.PabI, respectively (Grazulis et al., 2005; Miyazono et al., 2007).
The conserved structural core surrounding the HNH motif is termed the 'ββα-metal' fold. This
protein topology consists of two anti-parallel β-strands connected by a loop of variable length,
flanked by an α-helix (Mehta et al., 2004),(Kuhlmann et al., 1999). A binding site for a single
catalytic metal ion—typically magnesium—is embedded within this catalytic fold. In some
instances, significant insertions of additional structural elements are observed within this motif
(Eastberg et al., 2007; Stoddard, 2005). The ββα-metal fold can exist as an independently folded
catalytic domain (as observed in colicins) or it can be fused to additional protein domains that
dictate DNA binding specificity and cleavage activity.
The PD…(D/E)×K motif can be very well-suited for recognition of short DNA sequences with
high fidelity, because the catalytic center is surrounded by a densely packed array of side chains
that can contact neighboring base pairs in the major groove in a sequence-specific manner
(Orlowski and Bujnicki, 2008). In contrast, the HNH motif and its associated ββα-metal fold
appears less well-suited for this task. In order to target the scissile phosphate, the catalytic core
motifs of these enzymes primarily interact with the DNA backbone where they contribute little
to sequence-specificity and fidelity (Eastberg et al., 2007). Sequence-recognition by these
enzymes is therefore usually carried out by additional protein domains that are tethered to the
ββα-metal region, necessitating significant repackaging and augmentation of this catalytic
motif.
Recently, the structure of the ββα-metal restriction endonuclease Hpy99I was determined in
complex with its DNA substrate, 5' - CGWCG - 3', at 1.5 Å resolution (Sokolowska et al.,
2009) (W=A or T). Hpy99I binds as a homodimer and forms a ring-like structure that encircles
the DNA. The protein contacts all four C:G base pairs within both the minor and major groove,
and contacts the central base pair (A:T or T:A) in only the minor groove. The DNA is slightly
bent in the complex. All nucleotides in the target site are found in canonical Watson-Crick
basepair interactions.
In contrast, PacI is a 'rare-cutting' homodimeric HNH restriction endonuclease found in the
bacterium Pseudomonas alcaligenes. It recognizes the symmetric eight base pair duplex DNA
sequence 5' – TTAAT/TAA - 3' and cleaves each strand between the internal thymine residues
(as the position indicated by "/") to generate product fragments containing 2-base, 3’-overhangs
(Roberts et al., 2010). PacI is one of the smallest REases known, comprising only 142 amino
acids per subunit, eight of which are cysteines (Figure 1). Its gene resides within a super-
integron, a chromosomal array that contains multiple gene cassettes each flanked by a large
direct repeat sequence and mobilized by a common site-specific integrase (Vaisvila et al.,
2001). Unlike the vast majority of REases, which are accompanied by DNA-methyltransferases
that protect the cell’s own DNA from REase auto-digestion, PacI appears to be a solitary
enzyme with no companion methyltransferase (see Supplementary Material). Host protection
in this rare instance seems likely to depend not on the methylation of recognition sequences,
but rather on the absence of such sequences in the P.alcaligenes genome.
The length of the PacI recognition sequence (eight basepairs) places this enzyme in the
company of NotI and SfiI, two other 'rare-cutting' endonucleases the co-crystal structures of
which have been solved (Qiang and Schildkraut, 1987). NotI and SfiI belong to the PD…(D/
E)×K catalytic site superfamily, and in contrast to PacI, recognize sequences composed entirely
of G:C base pairs.
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Bioinformatics analysis of PacI (Orlowski and Bujnicki, 2008), and independent analyses with
online protein fold prediction servers such as PHYRE (Bennett-Lovsey et al., 2008) suggest
the presence of an HNH-related catalytic site. The likely presence of this motif, combined with
the opportunity to compare a ‘rare A:T-cutter’ to two ‘rare G:C-cutters’, led us to determine
the structure of PacI bound to DNA. PacI displays little resemblance to either NotI or SfiI, and
while it contains structural elements similar to those in Hpy99I, the arrangements of these
elements and the overall folds of the two proteins are strikingly different. PacI binding induces
an unusual distortion of its DNA target sequence that completely disrupts and reorganizes it
normal Watson-Crick duplex structure.
Results
Overall protein structure and catalytic site
The structure of PacI was determined in complex with its eight base pair cognate target site,
within the context of an 18 base pair synthetic DNA duplex. The structure was determined both
in the presence of calcium (yielding a co-crystal structure containing uncleaved DNA that
extended to 2.0 Å resolution) and in the presence of magnesium (resulting in a bound product
complex that was visualized at 1.9 Å resolution). The two structures are virtually identical,
with the exception of the presence of free 5' phosphate and 3' hydroxyl DNA product ends in
the endonuclease catalytic sites in the presence of magnesium. Data collection and refinement
statistics are provided in Table 1, and a detailed description of materials and methods is
provided in Supplementary Information. Examples of the experimental electron density,
calculated using phases derived by a combination of the multiple isomorphous replacement
(MIR) and single anomalous dispersion (SAD) methods, are shown in Supplementary Figure
S1.
The overall structure of the endonuclease homodimer bound to its DNA target is shown in
Figure 1a; two separate views of a single enzyme subunit are shown in Figure 1b. The overall
core topology of the PacI subunit corresponds to "β1–β2–α2–α3–β4–α4–α5", with the β3–β4–
α4 secondary structure elements comprising the ββα-metal catalytic site motif. This core
topology is further extended by very short β-hairpin motifs on the protein surface that are
involved in DNA contacts.
The PacI subunits display an extended structure containing a pair of bound zinc ions, each of
which is coordinated by four cysteine residues. The first zinc ion is entirely sequestered within
an N-terminal region (containing cysteines 4, 7, 24 and 27) that appears to be a unique feature
of PacI: the only three homologues of PacI currently in Genbank (all from strains of the
bacterium Campylobacter) display little sequence similarity to this region (Figure 1c). The
second zinc ion is buried in the enzyme core, and is also coordinated by four cysteine residues
(Cys 63, 66, 109 and 112). This zinc ion is located near the endonuclease catalytic site. Two
of the cysteine residues involved in its coordination (Cys 109 and 112) extend from the α4
helix from the ββα-metal motif.
The overall structural organization of the PacI enzyme resembles, at a superficial level, the
organization of the homodimeric HNH restriction endonuclease Hpy99I (Sokolowska et al.,
2009), and more distantly resembles the homodimeric HNH homing endonuclease I-PpoI
(Flick et al., 1998). All three proteins contain a catalytic ββα-metal motif and contain two
structural zinc ions embedded within each protein subunit, and all three position their active
sites across the minor groove to produce 3' overhangs. However, the extended architecture and
DNA binding modes of these enzymes are very different from one another (Figure 2 and
supplementary Figure S2), indicating that they appear to have independently acquired and then
optimized similar structural strategies for stabilization and catalysis, presumably after their
divergence from a common ancestral endonuclease.
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The backbone conformation and metal coordination exhibited by the 'ββα-metal catalytic core
of PacI is similar to those observed in other HNH endonucleases (Kuhlmann et al., 1999)
(Figure 3a and Supplementary Figures S2 and S3). Structure-based alignments of this region
with five separate ββα-metal endonucleases (E9 colicin, endonuclease VII, Hpy99I, I-HmuI
and I-PpoI) gives RMSD values for all backbone atoms of 1.5 to 1.8 Å, with corresponding
sequence identities ranging from as low at 6.7% (I-PpoI) to as high as 24% (I-HmuI and
EndoVII). A single divalent cation is bound within the PacI catalytic motif, where it is
coordinated by aspartate 92 and by asparagine 113, and also interacts with the 3' oxygen and
a nonbridging oxygen of the scissile phosphate. The distance from the metal to each DNA atom
is approximately 2.5 Å.
In spite of the structural similarity of the ββα-metal motif described above, the PacI catalytic
site displays a significant departure from the typical HNH motif. The position within the ββα-
metal motif that is normally the site of a histidine general base that activates the water
nucleophile (corresponding to His 149 in Hpy99I and His 98 in I-PpoI; Figure 3b and 3c) is
instead occupied by an arginine residue (Arg 93), that interacts with the 3' leaving group . In
place of the usual histidine, a neighboring tyrosine residue (Tyr 100) is instead positioned to
either assist in activation of a nucleophilic water (which is not observed), or perhaps to act
directly as a nucleophile itself. The distance from the tyrosine hydroxyl group to the phosphorus
atom is 4.5 angstroms in the uncleaved calcium-bound complex, and is 3.3 angstroms in the
cleaved magnesium-bound complex (the distance is reduced in the cleaved complex due to
rotation of the 5' phosphate group after cleavage). The nearest histidine residue (His 42) is
located approximately over 8 Å from the scissile phosphate, and like Arg 93, is located closer
to the leaving group than to the site of the nucleophilic attack. Thus, the PacI endonuclease
displays a dramatic alteration and rearrangement of the usual side chains found in a ββα-metal
catalytic motif, and perhaps a change in the actual cleavage mechanism.
To assess the relevance of Tyr 100, Arg 93 and His 42 in the PacI catalytic site, each was
mutated by PCR and the mutant proteins were expressed in vitro and assayed for activity (Table
2). To the best of our ability to measure, the amounts of each protein construct generated in
vitro, and then used in individual digest experiments, were comparable. Y100F was found to
be inactive (less than 10−4 WT activity) indicating that the phenolic oxygen of this amino acid
appears to be essential for catalysis . R93A and M were also inactive, but R93K displayed
reduced activity, suggesting that a positively charged group in this position in the catalytic site
is also essential. H42A displayed reduced activity (~10−2 WT activity). The putative metal-
binding residues Asp 92 and Asn 113 were also mutated and assayed. D92L and N113L were
inactive, whereas the D92A and N113A mutants displayed a low level of activity, indicating
that the metal-binding residues are critical components of the catalytic site.
The two bound zinc ions in PacI (each coordinated in a Cys4-Zn tetrahedral cluster) represent
a widely distributed conserved structural motif, distinct from the conventional trinucleotide-
specific Cys2-His2 ‘Zinc-finger’ domains of eukaryotic transcription factors (Supplementary
Figure S3). The Cys4-Zn motif comprises a pair of CxxC sequences. The first two cysteines
flank a loop, while the second two initiate an alpha helix (in some related sequences, the third
Cys or the fourth Cys is replaced by His instead). The region between each CxxC pair varies
in length and function, as does the helix. In many instances, this region includes catalytic
residues that contribute to the HNH catalytic site. Approximately 200 HNH-like domains are
aligned in pfam01844, and over one-third of these are embedded in Cys4-Zn motifs, indicating
that this structural architecture is often associated with an HNH catalytic site. In contrast, this
same region in the GATA family of transcription factors includes residues responsible for DNA
sequence recognition (Bates et al., 2008) (Supplementary Figure S3, panel L).
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DNA binding and recognition
The mode of DNA binding displayed by PacI is very unusual. The ββα-metal catalytic sites
from each protein subunit straddle the minor groove at the center of the DNA target, resulting
in an overall bend angle of approximately 90 degrees (Figures 1a and 4a). This results in a
dramatic widening of the minor groove (to approximately 18 Å) and a corresponding reduction
in the width of the opposing major groove. This bend is accompanied by a radical alteration
of the DNA duplex: every base throughout the target site is unpaired from its original Watson-
Crick partner (Figure 4b). Within the eight base pair target site, two bases on each strand are
completely unpaired, four are engaged in non-canonical A:A and T:T base pairs, and the
remaining two bases are matched with new Watson-Crick partners. This disruption of the DNA
duplex is entirely localized to the PacI target site; the base pairs immediately outside the 5' -
TTAATTAA - 3' sequence still display canonical B-form interactions. It does not appear that
crystal contacts play a role in these features of the protein-DNA complex: the solvent content
of the crystals is not unusual (about 55%) and the regions of the protein and DNA involved in
contacts and recognition are not located near symmetry mates in the crystal lattice.
The bound conformation of the DNA target was analyzed using the online program 3DNA
(Zheng et al., 2009) (Supplementary Figure S4). The perturbation of the DNA structure results
from significant distortion of the individual ribose moieties and the corresponding glycosidic
bonds between sugar C1' carbons and the corresponding nucleotide bases. Only three ribose
sugars on each strand (corresponding to −4T, −2A and +3A) are found in their original C2'-
endo pucker, while the remaining sugars are predominantly flipped into a C1'-exo
conformation. The chi angles linking the ribose C1' carbons to the N1 nitrogen of the thymines,
or to the N9 nitrogen of the adenines, deviate from their nominal B-form values by as much
as +/− 40°, leading to a rotation of individual bases that allows non-canonical A:A or T:T base
pairing. These base pairs still exhibit two intra-strand hydrogen bonds (Figure 4c), linking the
thymine-thymine pairs via the O2-N3 and N3-O4 atoms of their pyrimidine rings, and the
adenine-adenine pairs via the N6-N1 and N7-N6 atoms of their purine rings. These base pair
interactions, while rarely observed in DNA duplexes, are often found in folded RNA structures
(Olson et al., 2009).
The deformation of the DNA in PacI is accompanied by a significant unwinding of the duplex
at base step −2A in each DNA half-site (Supplementary Figure S4). That base, which is engaged
in an A:A base pair with its −1A partner, exhibits a −40° tilt and unstacking from its neighboring
(symmetry-related) A:A base pair. The local unwinding of each DNA half-site at these A:A
base pairs is complemented by local over winding of the adjacent base steps and base pairs,
allowing the rearranged DNA to maintain an overall duplex architecture. Although the DNA
backbone and its base pairing interactions exhibits a dramatic rearrangement in the bound
protein complex, all the individual base pairs (both Watson-Crick and non-Watson-Crick)
exhibit near normal values of propeller twist and buckle angles.
The PacI-DNA complex is further notable for the paucity of direct contacts between the protein
and the nucleotides. The two unpaired bases in each half site (+1T and +4A) are in direct contact
with amino acid side chains: +4A interacts in the major groove with Asn 32, and +1T interacts
in the minor groove with Arg 114 (Figures 4b and 4c). The adenines in the reorganized A:T
base pairs ( involving +3A in each DNA half-site) interact in the major groove with Asn 36.
One adenine in each A:A base pair makes a nonspecific contact to Ser 117 in the minor groove,
and the O4 groups of both thymines in each T:T base pair contact Lys 39 in the major groove.
To assess the importance of the major groove contacts, Asn 32, Asn 36, and K 39 were changed
by PCR to various other amino acids and the mutant proteins expressed in vitro and assayed
(Supplementary Table S1). Mutation of Asn 32 or Asn36 abolished activity (<10−4 WT
activity) indicating that these two amino acids are essential. Mutation of Lys 39 had little effect
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indicating that this amino acid is unimportant in spite of the hydrogen bonds it forms with the
T:T base pairs.
Thus, across the eight nucleotides in each DNA half-site, PacI makes only eight direct hydrogen
bond contacts: six in the major groove (N32 and N36 to adenine bases and K39 to thymine
bases) and two more in the minor groove. This represents a radical departure from the usual
strategy of restriction endonucleases which, under strong selective pressure for absolute
cleavage fidelity, usually make more direct contacts than are strictly necessary for high fidelity
sequence recognition.
Discussion
Diversity of site-specific HNH endonuclease scaffolds
The overall organization of PacI is superficially similar to the HNH restriction endonuclease
Hpy99I (Sokolowska et al., 2009), and more distantly related to the HNH (His-Cys box)
homing endonuclease I-PpoI (Flick et al., 1998). All are homodimers containing one ββα-metal
motif and two bound zinc ions per subunit. However, close examination of these three enzymes,
which recognize target sites ranging from 5 base pairs to 14 base pairs in length, indicates that
their folded structures, as well as their DNA binding modes and recognition mechanisms, differ
significantly (Figure 2). Whereas the core of the Hpy99I protein forms a structure that encircles
and binds almost orthogonally across and around its target site (with the helices from the
catalytic site ββα-metal motif aligned almost perpendicular with the DNA duplex axis), PacI
displays an elongated fold that associates with one face of the DNA target, with the two subunits
and the ββα-metal motif aligned nearly parallel to the DNA duplex. The structure of the I-PpoI
homing endonuclease is even more divergent: that protein relies upon extended β-sheet
structures for the completion of the core protein fold and for formation of its DNA binding
surface. Based on these observations, it seems likely that these site-specific HNH
endonucleases are distantly related, but probably all descended from a common ββα-metal
ancestor. That predecessor protein may have consisted of a nonspecific endonuclease folded
around the common catalytic motif (perhaps resembling modern colicin nucleases).
The details of the active site organization of PacI also indicate a significant divergence from
the usual architecture and mechanism that is observed for an HNH active site (Figure 3). The
presence of a tyrosine side chain at the position usually occupied by an imidazole base and
nucleophilic water, combined with the requirement of the tyrosine phenolic oxygen for
catalysis, indicates that this side chain might act as a direct nucleophile in DNA strand cleavage
(although a covalently trapped phosphotyrosyl intermediate has not been observed in either of
the structures determined in this study, and a role as a general base in more traditional
mechanism involving water-mediated hydrolysis cannot be ruled out). While such a
mechanism has not been observed previously for a restriction endonuclease, the BfiI enzyme
is a member of the phospholipase D family of nucleases, that includes many enzymes that
proceed via a phosphotyrosyl covalent intermediate, and is known to form a phospho-histidyl
covalent intermediate during strand cleavage (Sasnauskas et al., 2010).
DNA binding and perturbation
The appropriate balance of specificity, fidelity and affinity for protein-DNA interactions is one
of the most fundamental of biological requirements. Restriction endonucleases reside at one
end of the spectrum of possible DNA recognition behaviors: they cleave relatively short DNA
sequences that usually occur frequently within both the host genome and invasive DNA
sequences , and display extremely high fidelity that spares the host from off-target cleavage
(Pingoud et al., 2005).
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Protein-DNA recognition specificity is thought to depend upon a combination of direct readout
of the nucleotide bases through contacts between the protein and the DNA that can be direct
and/or water-mediated, and the additional 'indirect' exploitation of DNA conformational
preferences by inducing a DNA structural perturbation or bend that is favored by a limited
number of possible DNA sequences (Jones et al., 1999; Luscombe et al., 2001; vonHippel,
2007). Direct readout of DNA sequences is most effective within the major groove which is
physically accessible and also provides chemically distinct combinations of hydrogen-bond
partners from the four possible base pairs.
However, many specific DNA binding proteins augment these contacts with additional
interactions made within the minor groove (completely encircling the DNA), and in some cases
can achieve specificity entirely via interactions within and across the minor groove (Bewley
et al., 1998; Rohs et al., 2009). DNA-binding proteins that contact minor groove structural
elements may rely on a combination of DNA bending and surface complementarity (for
example, as displayed by the TATA binding protein) (Kim et al., 1993) and may also read out
local sequence-dependent shape and charge characteristics of the minor groove (as observed
for a variety of DNA-binding proteins, including the nucleosome core particle and the
Drosophila Hox protein SCR) (Rohs et al., 2009). In these examples, the DNA is dramatically
deformed, but the canonical Watson-Crick base pairing of the complementary strands is still
preserved.
Because of extreme pressure to maintain high fidelity of recognition, restriction endonucleases
are notable for their propensity to fully exploit multiple avenues of DNA readout and specificity
(a behavior that can be termed 'recognition overkill'). For example, the MunI restriction
endonuclease (a PD‥(D/E)×K enzyme which recognizes the six base pair sequence 5' -
GTTAAC - 3') establishes 16 direct hydrogen bonds to these bases in the major groove, 10
direct contacts to phosphates, and it induces a significant distortion between the central base
pairs in the sequence (Deibert et al., 1999). That protein displays approximately four direct
contacts per base pair--an accomplishment that is facilitated by its core fold, in which the
catalytic sites are surrounded by a densely packed array of polar side chains that can fully read
out the DNA target's sequence and its shape.
In contrast, PacI is one of the smallest known restriction endonucleases, yet it recognizes a
longer (eight basepair) target site while being folded around nonspecific catalytic motif that
primarily interacts with the phosphate backbone. Evidently, PacI achieves a similarly high
level of recognition-fidelity while forming far fewer hydrogen bonds to the nucleotides. Such
a minimal protein-DNA interface—in which less than 50% of potential hydrogen bond partners
within the target's major groove are engaged in direct contacts with the protein— would
typically be expected to correspond to greatly reduced fidelity of recognition (Chevalier et al.,
2003).
The rearrangement of the DNA conformation and its interstrand base pair contacts may
represent a mechanism that significantly increases specificity of recognition by PacI, without
requiring an investment by the enzyme in a large number of base-specific contacts. It is known
that the act of unstacking and/or unpairing consecutive base pairs can result in unfavorable
increases in free energy of binding. Computational and direct biophysical analyses indicate
that this energetic cost can differ by several kcal/mol per base step, depending on the sequence
context of the bases involved (Delcourt and Blake, 1991; Hobza and Sponer, 2002). The
sequestration of individual bases into unpaired conformations in the PacI complex and the
partial unstacking of flanking base pairs, may therefore greatly favor the correct target sequence
for binding and cleavage over closely related DNA sequences
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While no sequence-specific DNA binding proteins or endonucleases have displayed the
extreme basepair deformation and reorganization displayed by PacI, some restriction
endonudleases have been observed to unpair and 'flip out' individual bases, while also greatly
distorting the DNA backbone conformation. For example, the Ecl18kI enzyme flips the adenine
and thymine bases from each strand of its cognate 5'-CCNGG-3' target site and sequesters both
into protein binding pockets, as part of a mechanism that dramatically kinks the DNA and
greatly reduces the value for the rise between the flanking inner C:G basepairs, thus decreasing
the distance between scissile phosphates by several angstroms (Bochtler et al., 2006). A similar
deformation is seen for the PspGI restriction endonuclease (Szczepanowski et al., 2008), and
is presumably a common feature of many such REases that utilize base-flipping as part of their
recognition mechanism.
While the mechanism of nucleic acid recognition displayed by the PacI endonuclease appears
very extreme as compared to most sequence-specific DNA-binding proteins , the distortion of
the substrate and the contacts formed by the protein are in fact quite similar to the pattern of
RNA recognition exhibited by archaeosine tRNA-guanine transglycosylase, which modifies a
guanine base in the 'D arm' of its tRNA substrate. That enzyme disrupts all of the normal
basepair and tertiary interactions in the tRNA D arm, leading to reorganization of the tRNA
helical strucure and association of the G15 base with the enzyme active site (Ishitani et al.,
2003).
Initial cognate site recognition
Finally, the observation of such a dramatic reorganization of the PacI target site, involving
removal of each base from its complementary partner, begs the question of how the initial
moment of cognate site recognition is related to the subsequent formation of the catalytic
enzyme-substrate (ES) complex that is visualized in typical enzyme-DNA co crystal structures.
A long history of biophysical studies of protein-DNA recognition (recently revisited and
reviewed in (Halford, 2009)) indicates that DNA-binding proteins sample potential DNA
binding sites by rapidly associating and dissociating from non-cognate DNA sequences (a
process greatly accelerated by non-specific orientation and interaction between the oppositely
charged molecules), while also sliding back and forth across regions covering approximately
50 base pairs around each initial 'landing site' in a limited 1-dimensional search of nearby DNA
sequences.
It is generally assumed that the contacts made within the initial encounter complex between a
specific DNA-binding protein and its correct cognate target site are similar to those found in
the enzyme-substrate complex, with additional conformational changes driven by the binding
energy derived in the initial encounter with the cognate site. In this model, additional specificity
of recognition, beyond that which is engendered by the contacts made between protein and
DNA bases, can be derived by sequence-specific conformational preferences of the DNA. This
model also allows for the possibility that the unbound sequence of a cognate DNA target site
might be predisposed to physically sample a conformation that is similar to its final bound
state, which would also enhance recognition and high affinity binding.
However, the structure of the PacI endonuclease in complex with its cognate target site
indicates that for this enzyme, and perhaps for other highly specific DNA binding proteins, the
structure of the initial specific encounter complex might differ significantly from the
subsequent biologically or catalytically active complex. Its seems unlikely that the 5' -
TTAATTAA- 3' sequence recognized by PacI is predisposed to sample a conformation, in the
absence of bound protein, in which several bases are completely unpaired from their Watson-
Crick partners and the flanking base pairs are significantly unstacked. When examining the
current collection of crystallographic structures of protein-DNA complexes, many examples
can be found where the number of observable contacts between protein and DNA bases do not
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obviously correspond to the actual specificity of the binding interaction. While such
observations may be explained at least in part by the contribution of indirect readout to affinity
and specificity, it may be that some specific DNA-protein binding events may be driven by the
initial, transient formation of atomic contacts in the cognate complex that are difficult to
visualize using traditional crystallographic methods, and are then significantly rearranged to
produce the final catalytically or biologically active state.
Experimental Procedures
A detailed description of materials and methods is provided in Supplementary Information.
Briefly, the gene encoding PacI was isolated from Pseudomonas alcaligenes chromosomal
DNA and re-introduced into P.alcaligenes on a plasmid vector, resulting in a 48-fold increase
in endonuclease expression. A 100-L culture was grown of this over expressing strain, from
which 57 mg of homogeneous PacI was purified by FPLC column chromatography. The
specific activity, monitored by conventional DNA-digestion and agarose gel electrophoresis,
was approximately 6×105 units per mg. Crystals of the protein-DNA complex, using a synthetic
18 base pair DNA duplex corresponding to sequence 5' - GAGGCTTAATTAAGCCGC - 3'
and a complementary bottom strand were grown by hanging drop geometry against a
crystallization buffer containing 18 to 22% polyethylene glycol 3000 (PEG3K) 100 mM
sodium citrate, pH 5.5 and either10 mM MgCl2 or 2 mM CaCl2. The structure of the complex
in the presence of magnesium was determined using a combination of the multiple isomorphous
replacement (MIR) and single anomalous dispersion (SAD) methods, using five independently
generated heavy atom derivatives (two separate PtCl4 soaks, and one each of HgCN2, PIP and
WO4). In-house wild-type and heavy atom MIR datasets, using a rotating anode generator,
extended to approximately 2.6 Å resolution. In addition, a single SAD dataset (extending to
1.9 Å resolution) from a platinum-soaked was collected at the Advanced Light Source using
beamline 5.0.2. The combination of in-house and synchrotron data was used to determine and
refine the structure of the magnesium-bound, cleaved product complex. Subsequently, a second
2.0 Å resolution dataset of an unsoaked, wild-type crystal in the presence of calcium was also
collected and refined, yielding a corresponding model of the uncleaved protein-DNA complex.
Data and refinement statistics are provided in Table 1.
Coordinates and Data Deposition
The X-ray structure factor amplitudes and corresponding refined coordinates for the PacI/DNA
complex, in the form of calcium-bound uncleaved DNA and magnesium-bound cleaved DNA
structures, have been deposited in the RCSB database for immediate release (PDB ID code
3LDY and 3M7K). Requests for the PacI-overexpression clone should be direct to New
England Biolabs (xus@neb.com).
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
X-ray data was collected at the Advanced Light Source (ALS) synchrotron facility at the Lawrence Berkeley National
Laboratory (University of California) on beamline 5.0.2 with the assistance of ALS staff. We thank members of the
laboratories of Roland Strong and Adrian Ferre-D'Amare for advice and assistance during structure determination.
This work was supported by funding from the NIH to BLS (R01 GM49857) and by funding from the Fred Hutchinson
Cancer Center to the Program in Structural Biology.
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Figure 1. The structure of the PacI restriction endonuclease
The protein is colored according to individual structural domains and motifs; the same color
scheme is used in all panels. All figures were generated using the molecular graphics program
PYMOL (DeLano, 2002). Panel a: The PacI homodimer is bound to ts palindromic DNA target
sequence. The two protein subunits are colored green and cyan; the two bound zinc ions in
each protein subunit are labeled and colored dark green, and the single bound divalent cation
observed in each catalytic site is labeled 'Mg' and colored blue. The corresponding scissile
phosphates, that yield 2 base, 3' cohesive overhangs when cleaved, are colored red. Panel b:
A single subunit of PacI is shown, in two separate orientations. The left panel shows the same
orientation as the cyan-colored subunit in panel a. The right panel shows the same subunit,
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rotated by 90° around the horizontal axis. In both panels, the ββα-metal motif is colored dark
green, and the unique N-terminal region (which binds zinc number 1) is colored blue. On the
left, the eight cysteine ligands to the two zinc ions are labeled. On the right, the single bound
divalent cation (magnesium in the product complex) and the two residues involved in its
coordination (D92 and N113) are labeled. Panel c: Sequence homology between PacI, and its
three recognizable homologues (hypothetical protein sequences from Campylobacter
concisus, cowae and lari, respectively). The ββα-metal motif is indicated by the box. Residues
that directly contact DNA are indicated with asterisks arrows; catalytic residues of the HNH
endonuclease motif are indicated with blue font and asterisks; zinc-binding cysteine residues
are indicated with red font and asterisks. The N-terminal regions, that harbors the first bound
zinc ion, is indicated with light blue font, corresponding to the coloring of the same region in
panel b. See also supplemental Figure S1.
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Figure 2. ββα-metal (HNH) endonucleases that contain two bound zinc ions per subunit
For each endonuclease structure, the left panels show comparable orientations of the bound
DNA targets, while the right panels show comparable orientations of the protein subunits (with
the dyad symmetry axis running vertically and the protein subunits oriented to the left and right
side of that axis). For all three structures, the ββα-metal catalytic motif is colored blue, and the
bound zinc ions are shown as teal spheres. Panel a: The PacI restriction endonuclease. While
the protein's overall organization of secondary structure elements, relative to the catalytic sites
and bound zinc ions, is similar to Hpy99I, the overall architecture of the individual ββα-metal
repeats, as well as the mode of DNA binding, is significantly different. Panel b: The core of
the Hpy99I restriction endonuclease, which cleaves a five base pair CGWCG target, generating
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a 5-base, 3' overhang (Sokolowska et al., 2009). An N-terminal β-barrel domain (residues 1 to
53) that is not involved in DNA binding is removed for clarity. The remaining protein
homodimer structure includes all catalytic and zinc-binding regions that correspond to those
observed in PacI. Note that the orientation of the protein is roughly orthogonal to the major
axis of the DNA duplex, which is relatively unbent. Panel c: The I-PpoI restriction
endonuclease (a fourteen-base cutting homing endonuclease found in the eukaryotic amoeboid
Physarum polycephalum) (Flick et al., 1998). The overall bend of the DNA target is similar to
that displayed by PacI; however all base pairs are maintained in their original Watson-Crick
base pairing arrangement. The protein fold, beyond the core ββα-metal motif, is completely
different from that displayed by Hpy99I and PacI, indicating that divergence of these protein
lineages may have occurred at an early stage, from a simple ββα-metal scaffold prior to
subsequent elaboration and specialization. See also supplemental Figure S2.
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Figure 3. ββα-metal motifs and catalytic sites for HNH restriction and homing endonucleases
The core catalytic motif, consisting of a two-stranded antiparallel beta-sheet and single alpha-
helix, is shown in approximately the same orientation for (Panel a) PacI; (Panel b) Hpy99I
and (Panel c) I-PpoI . The scissile phosphate is shown in red, flanked by its 5' and 3' nucleosides
in gray. The histidine general base is colored and labeled with red, and the corresponding water
nucleophile is a small light blue sphere. In all three catalytic sites, a single bound divalent metal
ion (dark blue larger sphere) is coordinated by two asparagine/aspartate residues. An additional
polar residue (participating in cleavage as a Lewis acid, whereby it stabilizes the phosphoanion
transition state) is present in each catalytic site that is positioned to help satisfy the charge on
the phosphate during cleavage (R93 for PacI; H151 for Hpy99I, and R61 for I-PpoI). Note that
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PacI displays a structural inversion in the positions of the histidine general base and the Lewis
acid: His 42 is presented from a loop outside the ββα-metal motif, while R93 is found in the
position that is usually occupied by an HNH histidine base. Finally, peripheral cysteine residues
in all three ββα-metal motifs are involved in coordination of a structural zinc ion (C109 and
C112 in PacI). However, the location of the bound zinc in the restriction endonucleases (panels
a and b) differ from the homing endonuclease (panel c), indicating that these enzyme lineages
may have diverged from a common ancestor prior to development of these metal binding sites.
See also supplemental figure S3.
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Figure 4. DNA recognition and binding by PacI
Panel a: Conformation of the bound DNA target site. The individual bases of the 'TTAATTAA'
target are colored both by their position in the original DNA half-site (left half-site = green;
right = yellow) and by their identity (dark bases are adenine; light bases are thymine). The
bound DNA is shown in a stick representation on the left, and in a cartoon representation,
generated by the program 3DNA (Zheng et al., 2009), on the right. Those bases that lie outside
the eight base pair target site are colored grey. Panel b: Cartoon representation of the base
pairing interactions between the two target site strands and the contacts between the protein,
DNA and solvent molecules. The individual bases are colored as shown in panel (a) above.
The numbering of the bases corresponds to the position in the original unbound target sequence,
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with bases −4T to −1A corresponding to the 5' half (left half) of the target site along one DNA
strand, and bases +1T through +4A corresponding to the 3' (right) half of the target site along
the same strand. In the bound complex, all bases in the target are removed from their original
Watson-Crick partners, so that some bases are unpaired (+1T and +4A on each strand), some
are found in noncanonical A:A and T:T base pairs (+2T, −1A, −2A and −3T on each strand)
and some are found in new Watson-Crick base pairs (−4T with +3A from each strand). The
scissile phosphates on each strand are red; well-ordered water molecules are blue. Protein
residues (from only one of the two protein subunits, for clarity) that are involved in direct or
water-mediated contacts to the DNA are indicated; those that form direct contacts to DNA
bases are boxed. Panel c: Structural interactions between each unique DNA base or base pair
in a single half-site to solvent and protein residues. The numbering of bases is consistent with
panels a and b above. See also supplemental figure S4.
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Table 1
Data collection and refinement statistics
Data set Id
Native-1
Native-2
PtCl4-1
PtCl4-2
HgCN2
PIP
WO4
Wavelength (Å)
1.5418
0.97741
1.0719
1.5418
1.5418
1.5418
1.5418
Data collection
Space group
C2221
C2221
C2221
C2221
C2221
C2221
C2221
a (Å)
36.86
36.86
37.09
37.32
36.93
36.89
37.83
b (Å)
115.75
115.75
115.16
114.08
116.08
117.79
116.05
c (Å)
114.37
114.37
114.83
114.32
114.19
113.89
114.14
Resolution (Å)
50-2.64
38-1.97
50-1.92
25-3.2
50-3.0
50-2.6
50-3.07
Unique reflections
7568
17385
19094
4343
5193
7947
5014
Redundancy*
6.6(4.7)
10.7(4.2)
13.5(11.1)
6.9(7.1)
13.4(8.9)
7.1(7.0)
11.0(7.8)
Completeness (%)*
99.8(98.7)
97.2(80.1)
98.6(92.4)
100(100)
99.8(99.6)
99.1(93.5)
99.4(94.4)
I/σ*
20.4(5.0)
27.8(3.3)
44.2(3.4)
20.3(5.7)
28.7(7.1)
37.1(11.8)
32.9(13.9)
Rmergea (%)*
9.2(31.2)
6.8(22.9)
6.2(22.6)
9.7(39.3)
8.6(31.9)
4.5(13.9)
6.0(12.5)
B(iso)(Å2)
47.4
26.25
29.04
60.4
61.1
58.7
53.5
Refinement
Protein atoms#
1108
1108
1108
DNA atoms#
366
366
367
Heavy atoms
2 Zn+2
2 Zn+2
2 Zn+2, Pt+2
Catalytic Metal ions
Ca+2
Ca+2
Mg+2
Cations
---
---
SO4−2
Solvent molecules
76
94
114
R-factorb (%)*
0.208(0.293)
0.184(0.227)
0.172(0.217)
R-freeb (%)*
0.278(0.319)
0.217(0.361)
0.201(0.285)
Rmsd
Bond length (Å)
0.012
0.013
0.011
Angles (°)
1.667
1.541
1.305
Ramachandran (%)
Core region
97.83
96.38
98.43
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Data set Id
Native-1
Native-2
PtCl4-1
PtCl4-2
HgCN2
PIP
WO4
Wavelength (Å)
1.5418
0.97741
1.0719
1.5418
1.5418
1.5418
1.5418
Allowed region
1.45
2.90
1.57
Outliers
0.72
0.72
0.00
*Highest resolution shell values in parenthesis.
aRmerge = Σ|Ihi - <Ih> |/ΣIh, where Ihi is the ith measurement of reflection h, and <Ih> is the average measured intensity of reflection h.
bR-factor/R-free = Σh|Fh(o) - Fh(c)|/Σh|Fh(o)|. Where R-free was calculated with 5% of the data excluded from refinement.
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Table 2
Activity of PacI site-directed mutants
PacI variant
Endonuclease
activity
Units per
25 µl in vitro
reaction
Wild-type PacI
+++
~200
Catalytic:
H42A
+
~1
R93K
+/−
~0.2
R93A,M
−
<0.01
Y100F
−
<0.01
Mg2+-binding:
D92A,L
+/−
<0.1
N113A,L
+/−
<0.1
Specificity:
N32A,T,L,D
−
<0.01
N36A,T,L,D
−
<0.01
K39A,M
++
>10
Endonuclease activities of wild-type PacI and of mutant derivatives. Mutants were constructed by two-step PCR, expressed in vitro using the
PURExpress™ transcription/translation system, and assayed by DNA-digestion and gel electrophoresis, as described in the Supplementary Material.
The standard 25 µl PURExpress™ reaction produced approximately 200 units of endonuclease activity from the wild-type PacI gene template (1 unit
completely digests 1 µg of substrate DNA to completion in 1 h at 37°C). The limit of endonuclease activity detectable in this assay corresponded to
10−4-fold less than wild-type, or approximately 0.01 units. In most cases, several different mutants were constructed for each amino acid targeted for
alteration. Mutants yielding the same result are grouped together on a single line in the table; thus, ‘N36A,T,L,D’, for example, signifies that Asn 36
was individually changed to Ala, Thr, Leu, and Asp, and all four mutant enzymes behaved similarly—in this case displaying no detectable endonuclease
activity. Plus and minus symbols in column 2 indicate the relative levels of endonuclease activity observed across several independent experiments.
These levels are quantified approximately with respect to wild-type in column 3.
Structure. Author manuscript; available in PMC 2011 June 9.
|
3M7L
|
Crystal Structure of Plant SLAC1 homolog TehA
|
Homolog Structure of the SLAC1 Anion Channel for Closing
Stomata in Leaves
Yuhang Chen1,6, Lei Hu2, Marco Punta6,7, Renato Bruni6, Brandan Hillerich6, Brian Kloss6,
Burkhard Rost1,6,1, James Love6, Steven A. Siegelbaum2,3,5, and Wayne A.
Hendrickson1,4,5,6
1Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY
10032, USA
2Department of Neuroscience, Columbia University, New York, NY 10032, USA
3Department of Pharmacology, Columbia University, New York, NY 10032, USA
4Department of Physiology and Cellular Biophysics, Columbia University, New York, NY 10032,
USA
5Howard Hughes Medical Institute, Columbia University, New York, NY 10032, USA
6NYCOMPS, New York Structural Biology Center 89 Convent Avenue, New York, NY 10027 USA
7Department of Computer Science and Institute for Advanced Study Technical University of
Munich D-85748 Munich, Germany
Summary
The plant SLAC1 anion channel controls turgor pressure in the aperture-defining guard cells of
plant stomata, thereby regulating exchange of water vapor and photosynthetic gases in response to
environmental signals such as drought or high levels of carbon dioxide. We determined the crystal
structure of a bacterial homolog of SLAC1 at 1.20Å resolution, and we have used structure-
inspired mutagenesis to analyze the conductance properties of SLAC1 channels. SLAC1 is a
symmetric trimer composed from quasi-symmetric subunits, each having ten transmembrane
helices arranged from helical hairpin pairs to form a central five-helix transmembrane pore that is
gated by an extremely conserved phenylalanine residue. Conformational features suggest a
mechanism for control of gating by kinase activation, and electrostatic features of the pore coupled
with electrophysiological characteristics suggest that selectivity among different anions is largely
a function of the energetic cost of ion dehydration.
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Correspondence and request for materials should be addressed to WAH (wayne@convex.hhmi.columbia.edu)..
Supplementary Information is linked to the online version of the paper at www.nature.com/nature.
Author Contributions YC, LH, MP, RB, BH, BK and JL performed experiments; YC, LH, BR, SAS, and WAH analyzed data; YC,
LH, SAS, and WAH prepared the manuscript.
Author Information Atomic coordinates and diffraction data are deposited in the Protein Data Bank with accession codes 3M71,
3M73, 3M74, 3M75, 3M76, and 3M7L. See Table S4 for identifications. Reprints and permissions information is available at
www.Nature.com/reprints.
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Published in final edited form as:
Nature. 2010 October 28; 467(7319): 1074–1080. doi:10.1038/nature09487.
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Stomatal pores in the leaves of plants permit the influx of atmospheric carbon dioxide in
exchange for transpirational evaporation of water1,2. A pair of kidney-shaped guard cells
define each pore aperture, and turgor pressure variation in these cells determines the degree
of stomatal pore openness. Depending on diverse environmental factors, the stomata close to
prevent H2O losses and open to admit CO2 for photosynthesis. Environmental stimuli that
lead to stomatal closure include darkness, high CO2 levels, ozone, low air humidity and
drought. The plant hormone abscisic acid (ABA) is critical for signal transduction from
these stimuli. Mutational screens in Arabidopsis thaliana for CO2 and ozone sensitivity
identified a protein with ten predicted transmembrane (TM) helices, now called slow anion
channel 1 (SLAC1), as having a central role in the control of stomatal closure3-5. Recent
studies proved that SLAC1 is indeed an anion channel6-7, with characteristics like those of
slow anion channels found in guard cells8, and that it is activated by phosphorylation from
the OST1 kinase9. OST1 activity is negatively regulated by the ABI1 phosphatase10,11,
which is in turn inhibited by the stomatal ABA receptors PYR/RCAR12,13 when in the
ternary hormone-receptor-phosphatase complex14-18. Thereby, ABA stimulates SLAC1
channel activity. Resulting Cl− efflux through SLAC1 causes membrane depolarization,
which activates outward-rectifying K+ channels, leading to KCl and water efflux to further
reduce turgor and cause stomatal closure.
SLAC1 expression in Arabidopsis is confined to the guard cells of leaves, but other
Arabidopsis tissues do have SLAC1 homologs3, named SLAH1-SLAH4. The identifying
mutations slac1-14 and slac1-23 are, respectively, in predicted transmembrane segments 9
(S456F) and 1 (G194D) of a protein that includes substantial N- and C-terminal extensions
outside a 10-helix transmembrane (TM) domain. SLAH1, which is absent from leaves and
lacks the terminal extensions of SLAC1, fully complements the mutant phenotype in slac1-2
guard cell protoplasts3. SLAC1 and homologs are also present in other plant genomes,
including nine in rice (Oryza sativa) and five in grape vines (Vitis vinifera). SLAC1
relatives, some quite remote, also occur in bacteria, archaea and fungi. Known prokaryotic
homologs contain only the predicted transmembrane domain of SLAC1, but some fungal
homologs do have N- and C-terminal extensions. One homolog, Mae1 from the yeast
Schizosaccharomyces pombe, functions as a malate uptake transporter19; another, Ssu1 from
Saccharomyces cerevisiae and other fungi including Aspergillus fumigatus, is characterized
as a sulfite efflux pump20,21; and TehA from Escherichia coli is identified as a tellurite
resistance protein by virtue of its association in the tehA/tehB operon22,23. Despite a lack of
further biochemical characterization, many homologs are annotated as tellurite resistance/
dicarboxylate transporter (TDT) proteins.
We have undertaken structural and functional characterizations of the SLAC1 anion
channel. We first solved an atomic-resolution crystal structure of the TehA homolog from
Haemophilus influenzae, and we then developed a homology model for Arabidopsis SLAC1.
This model allowed us to conduct mutagenesis for functional testing of structure-inspired
hypotheses on gating and selectivity. We expressed Haemophilus TehA and Arabidopsis
SLAC1 in Xenopus oocytes to characterize channel properties of these proteins and mutant
variants. We also determined crystal structures for several mutant variants, including the
homolog of slac1-2.
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Structure of SLAC1 bacterial homolog TehA
We performed a bioinformatic analysis of SLAC1-related proteins, first clustering nearly
900 non-redundant sequences into a superfamily at the PSI-BLAST level E≤10−3, then into
three distinct families at an initial threshold of E≤10−30, and finally into subfamilies at a
typical initial threshold of E≤10−55. Since previous annotation is not well founded in
experiment and SLAC1 is now the best characterized member, we adopt a nomenclature
defining a SLAC superfamily divided into families identified as SF1-SF3 and subfamilies
SF1A, SF1B etc. Family SF1 comprises the plant SLAC proteins and close bacterial
homologs, family SF2 comprises a distinct set of bacterial proteins often annotated as
exfoliative toxins, and family SF3 comprises the fungal Mae1 and Ssu1 proteins and their
archaeal or bacterial homologs, respectively. SLAC family SF1 has three large subfamilies:
the plant SLAC and SLAH proteins are in subfamily SF1A, closest bacterial homologs are
in SF1B, and the TehA homologs are in SF1C (Fig. 1a). The other families also divide into
subfamilies as detailed in Table S1, and family SF1 is divided into sub-subfamilies (Table
S2). Two pertinent SF1 sequences are aligned in Fig. 1b.
We used a structural genomics approach to obtain structural information, testing expression
and purification for 43 bacterial and archaeal likely homologs, assaying for detergent choice
and stability on 8 of these, finding two with appropriate profiles by size exclusion
chromatography, and obtaining suitable crystals for one. This protein, TehA from H.
influenzae (HiTehA), was found to be trimeric both by size-exclusion multi-angle light
scattering (SEC-MALS) measurements and by chemical cross-linking. When solubilized in
β-octylglucoside, HiTehA crystallized in space group R3 with a=b=96.01Å and c=136.27Å.
Each asymmetric unit contains one subunit and 65% solvent. The structure was solved by
selenomethionyl (SeMet) SAD phasing, ultimately at 1.50Å resolution (Table S3, Fig. S1),
and then refined at 1.20Å resolution (Fig. 2a) to R/Rfree values of 14.1%/15.7% for a model
that includes ordered residues 6-313, 213 water molecules and four detergent molecules
(Table S4).
The crystal structure has TehA trimers aligned with three-fold axes of the lattice (Fig. 2b).
Subunits are tightly associated, burying 8947 Å2 of total surface area within trimer
interfaces. The electrostatic potential surface is largely negative on the extracellular surface
(Fig. 2c) and largely positive on the cytoplasmic surface (Fig. 2d). The membrane
orientation is specified experimentally from GFP tagging of E. coli TehA24. Each TehA
protomer has ten TM helices, as predicted; however, the fold is novel. Tandemly repeated
helical hairpins are arranged with quasi-five-fold symmetry (Figs. 2e,S2). Extracellular
inter-helix loops are short (2-5 residues) whereas intracellular inter-helix connections are
longer, including a nine-residue helix H2,3 between TM2 and TM3 (Fig. 2f). An inner pentad
of outwardly directed, TModd, helices creates an apparent pore through each protomer
perpendicular to the putative membrane plane. TMeven helices from the five hairpins
surround the inner pore and make an outer layer.
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Homology model for plant SLAC1
Arabidopsis SLAC1 (AtSLAC1) is substantially similar to bacterial homologs, notably
HiTehA (Fig. 1b). All HiTehA TM helices are fully aligned to predicted SLAC1 TM
helices, but there are short inter-helical gaps (1-5 residues) in all five extracellular loops and
in two of the intracellular loops. The TM domain of AtSLAC1 (residues 188-504) aligns to
HiTehA with 19% sequence identity and with a PSI-BLAST E-value of 3×10−22. For
comparison, and in keeping with the family tree (Fig. 1a), the TM domain of AtSLAC1
shares sequence identities of 76% with rice SLAC1, 41% with Arabidopsis SLAH1, 25%
with an SL1B homolog from Halorhodospira halophila, 11% with S. cerevisiae Ssu1, and
9% with S. pombe Mae1. A conceptual model with the AtSLAC1 sequence transposed onto
the HiTehA helices sufficed to guide most of our mutational tests, but a detailed AtSLAC1
homology model helped to refine our ideas. Surface variability and electrostatic potential are
plotted onto the surface of this model (Fig. 2g,2h).
The most remarkable feature of the TehA structure and corresponding SLAC1 model is the
central pore through each protomer. As for acetylcholine receptors25, the SLAC1 pore is
formed by five helices; but the SLAC1 helices come from one protein molecule rather than
five. The SLAC1 pore has a relatively uniform diameter of approximately 5Å across nearly
five helical turns (Fig. S3), except for a pronounced constriction in the middle of the
membrane (Fig. 3a) where the pore is occluded by the side chain of Phe 450 (Phe 262 in
HiTehA). This residue is the only absolutely conserved amino acid residue of the SLAC1
family. The pore is lined with highly conserved (86% identity among five SLAC1 orthologs;
32% identity between AtSLAC1 and HiTehA) and generally hydrophobic residues (Figs. 1b,
3b,3c,S4). Despite this hydrophobicity, the electrostatic potential on the pore surface is
polarized (Fig. 3a), presumably due to an invaginated shape adjacent to charged residues
outside the membrane. The generally electropositive character of the cytoplasmic surface
likely contributes to anion efflux.
Kinks in the pore helices contribute to formation of a relatively constant pore diameter
across the membrane. Four of the five HiTehA inner helices have centrally located proline
residues, which necessarily generate kinks, and TM9 is kinked at a backbone-coordinated
water molecule (Fig. 3d). Proline replaces Gly263 in all SF1A and SF1B relatives, including
Pro451 AtSLAC1. This water-displacing change is isostructural26. Centrally located prolines
also prevail in TM3, TM5 and TM7 across the SF1 proteins. The outer helices are longer and
straighter, but more inclined. The only outer-helix proline kink of HiTehA is in TM6 at the
trimer three-fold axis.
Two of the gene-identifying mutations in Arabadopsis SLAC1 are selected point mutations,
others are disruptive transfer DNA (T-DNA) insertions4. In the AtSLAC1 homology model,
the slac1-2 (G194D) mutation3 points into the pore from TM1 and can be accommodated
structurally whereas the slac1-1 (S456F) mutation4 points away from the pore six residues
after pore-blocking Phe450 on TM9 and would be expected to disruptive. Ser456 interacts
with outer helix TM10 in the homology model, and the phenyl bulk from S456F would not
fit. Position 456 has alanine in HiTehA, 66% of all 204 SF1 homologs and another 27%
have threonine or serine (as in all SLAC1 channels); phenylalanine never occurs among all
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814 SLAC superfamily members. Position 194 has glycine in 58% of SF1 homologs and
alanine in another 22%. Asp194, which never occurs naturally, would block the pore and be
expected to repel anions.
Mutational tests of channel function
Mutational studies corroborate the hypothesis that a TehA-based SLAC1 model is
appropriate. First, as discussed above, the slac1-1 mutant (S456F) is expected to be
structurally disruptive, and indeed it is inactive in guard cells4, and the slac1-2 mutation
(G194D) is expected to block the pore, and we show below that this variant is also inactive.
We have also shown that the introduction of SLAC1-conserved proline residues into
HiTehA (A208P/G263P) is accommodated isomorphously26. Moreover, as shown below,
channel conductance properties of several mutants are similar for AtSLAC1 and HiTehA.
To examine characteristics of the SLAC1 channel in light of the structural model, we
performed electrophysiological tests of membrane currents from voltage-clamped Xenopus
oocytes following injection of wild-type or mutant AtSLAC1 or HiTehA cRNAs. We
observed modest-sized chloride currents with wild-type AtSLAC1 cRNA, as found
previously6,7, but did not detect any chloride current following injection of wild-type
HiTehA cRNA. We found that SLAC1 Cl− conductance was enhanced when the OST1
kinase cRNA was co-injected with SLAC1 cRNA, but only to the levels found by Lee et al.6
and not to the much higher levels found by Geiger et al.7 with OST1 physically connected to
SLAC1 by split YFP linkage. In keeping with structural evidence that Phe262 blocks the
HiTehA pore, removal of the phenyl group in HiTehA F262A or in the homologous
AtSLAC1 F450A resulted in very large Cl− currents relative to wild-type levels, and the
SLAC1 currents were now less enhanced by presence of OST1 (Fig. 4a). The tempting
interpretation of an opened gate will require validation with appropriately analyzed single-
channel recordings27. In keeping with the slac1-2 phenotype3, neither the functionally
impaired AtSLAC1 G194D nor its HiTehA G15D homolog showed any substantial
conductance; moreover, further consistent with pore blockage by Asp194 in slac1-2, the
large conductances of HiTehA F262A and AtSLAC1 F450A were stopped in the double
mutants AtSLAC1 G194D/F450A and HiTehA G15D/F262A (Fig. 4a). Here again, the
effects in SLAC1 were independent of OST1.
We also tested conductance characteristics for a series of AtSLAC1 F450X substitution
mutants – F450A,G,T,V,L– and for the corresponding HiTehA F262X series –
F262A,G,T,V,L (Figs. 4b,S5, Table S5). Findings from the two series are roughly parallel;
in particular, the alanine and glycine substitutions lead to large currents for both and in
comparison to the others. There are distinctions, of course, including generally higher
conductances for AtSLAC1 over HiTehA and less conductance of F262T TehA compared to
F450T SLAC1. It is also noteworthy that OST1 activation is very muted for the F450A,G,L
mutants, which is consistent with SLAC1 gating at Phe450.
Crystal structures were also determined for several of the HiTehA mutant variants (Table
S4). The structures of F262A (1.15Å), F262V (1.60Å), F262L (1.65Å) and G15D (1.50Å)
are all essentially isomorphous with the wild-type TehA structure with changes localized
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primarily at the sites of mutation; the same is true for the double mutations of F262A/G15D,
F262G/G15D and A208P/G263P26. The F262A structure has a wide open pore (Fig. 5a)
with a relatively uniform pore diameter of ~5Å through ~30Å across the membrane (Fig.
S3) whereas G15D has a doubly occluded pore (Fig. 5b). The pores of other mutant variants
are consistent with the sizes of constrictive residues and with the observed conductances.
Gating and activation
The crystal structures of TehA and its mutant variants when taken together with the
functional studies in Xenopus oocytes point to a crucial role for Phe450 in gating of the
SLAC1 anion channel. Conservation of this residue across the SLAC1 family implies
functional importance. The occlusion of the pore by the presence of F262 in the structure of
wild-type TehA and the openness of the pore upon its substitution by alanine in the structure
of the F262A mutant provides physical evidence for a gating role of this residue. This
interpretation is supported by the correlated conductance characteristics from variants of the
AtSLAC1 and HiTehA channels (Figs. 4b,S5). While these observations may suffice for
placing the gate within the channel pore, they do not by themselves suggest a mechanism for
gating in response to physiological stimuli. Some insight does come from conformational
details defined at high resolution.
One important structural clue is that the side chain of Phe262 is in a high energy
conformation in the HiTehA structure, with χ1/χ2 at −160°/-4°. Although χ1 is in a preferred
trans conformation, the phenyl ring is restricted by contacts with Val210 and Leu18 to a χ2
value near 0° rather than near to the preferred 90° orientation (Fig. 5c). Further evidence that
Phe262 is restrained from local equilibrium comes from shifts observed in crystal structures
of the F262A, F262V, F262L and F262G/G15D variants, which all show consistent
backbone movements that displace Cβ(262) by 0.47-0.78Å (Fig. 5d). By contrast, Leu262 in
F262L is in a preferred trans/gauche+ conformation at χ1/χ2 = 177°/63° as is Val262 in
F262V at χ1 = −176°. What might control activation of HiTehA is unclear, but for AtSLAC1
activation is by OST1 phosphorylation6,7. The molecular consequences of OST1
phosphorylation of SLAC1 remain unknown, but it is plausible that associated shifts in pore-
helix orientations would unlatch Phe450 in SLAC1 from a TehA-like restrained orientation.
By analogy with Leu262 in F262L, we expect a preferred rotameric state for Leu450 in
AtSLAC1 F450L. Thus, the lack of appreciable OST1 activation of conductance in
AtSLAC1 F450L (Fig. 5e) might be explained by the lack of a restraining latch, whereby the
channel remains closed despite OST1 activation. Puzzles certainly remain since OST1 does
substantially activate AtSLAC1 F450T or F450V, which like HiTehA F262V should also be
unrestrained; presumably activating adjustments widen the pore enough for ion permeation
past threonine and valine but not leucine.
Phosphorylation sites have been discovered in the N- and C-terminal tails of AtSLAC16,7,28
(179 and 51 residues long, respectively), but these alone cannot explain OST1 activation of
SLAC1. First SLAH1, which fully complements the slac1-1 mutation, does not have these
cytoplasmic tails. Secondly, although OST1 phosphorylation of Ser120 in the N-terminal tail
is necessary for SLAC1 activation, it is not sufficient7. Thus, we surmise that direct
phosphorylation of the SLAC1 TM domain must be critical, and SLAC1 has four conserved
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Ser/Thr candidates in its cytoplasmic loops. Moreover, SLAC1 proteins have proline-
mediated kinks at the putative Phe450 gate in helix TM9, and also in adjacent helix TM7;
these features may play a role in phosphorylation-driven unlatching of the Phe450 gate in
SLAC1.
Ion selectivity and discrimination
Our studies of SLAC1 channel relative ion permeabilities, based on measurements of current
reversal potential, are consistent with earlier work demonstrating that AtSLAC1 conducts
anions but not cations and is selective among anions, with greater permeability for nitrate
than for chloride (as in Vicia faba guard cell protoplasts27) and much reduced permeability
for malate, bicarbonate or sulfate6,7. We also find that SLAC1 has little permeability for
sulfite. Additionally, we find that wild-type SLAC1, F450A and F450T all have similar
relative permeabilities to chloride, sulfite and malate, despite having widely different
conductance levels, but the gating mutants do show small but significant decreases in nitrate
permeability (Fig. 4c, Table S6).
The relative insensitivity of anion permeability to gating residue changes suggests that
selectivity for these anions may occur away from the central constriction at the channel gate.
To some extent, ionic discrimination must depend on pore geometry; thus, an organic anion
such as malate may be simply too large to pass through the 5-Å wide pore. Although the
SLAC1 pore is lined largely with hydrophobic sidechains (Fig. 3c), it is sprinkled with
hydroxyl groups from serine and threonine residues (16%); their electropositive hydrogen
atoms may facilitate conductance. Most strikingly, the electrostatic potential within the
AtSLAC1 pore is electropositive throughout (Fig. 3a). This polarization, promoted by
charges on extra-membranous loops, no doubt contributes significantly in discrimination
against cations.
The relative anion permeability sequence of SLAC1 determined by us and others, I− > NO3−
> Br− > Cl− 6,7, corresponds to selectivity sequence 1 compiled by Wright and Diamond29
for a range of anion-selective proteins. This sequence correlates inversely with the hydration
energies of monovalent anions – anions with a lower hydration energy have a greater
channel permeability. It is thought to be generated in proteins with weak, low field-strength,
anion binding sites, where selectivity is largely determined by the energetic cost of anion
dehydration. These selectivity results are thus consistent with the SLAC1 structure, where
the pore lacks any obvious anion binding site.
Distinctiveness of the SLAC1 channel
SLAC1 anion channels are entirely novel in structure and, apparently, in the mechanism for
ion conductance. The best characterized of anion channels belong to the CLC family of Cl−
channels and transporters30-32. CLC channels have an altogether different architecture from
the SLAC1 channel, and the mechanism for selectivity is also very different. Bacterial CLC
transporters bind halide ions at three sites in a highly constricted pore30. By contrast, the
SLAC1 pore has a relatively uniform diameter across the membrane, except where closed by
the gating phenyl group, and we do not find discrete ion binding sites. CLC selectivity is
governed by specific residues surrounding these binding sites30,32. The anion selectivity
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sequence for the CLC channels of Cl−>Br−>NO −3>I−, opposite of that in SLAC1, is
consistent with the high field-strength anion binding sites in CLC channels29. Interestingly,
as for AtSLAC1, an Arabidopsis CLCa channel also preferentially transports nitrate ions33,
and an E. coli CLC channel is converted to preference of nitrate when a generally conserved
serine at the central site is substituted with proline as in AtCLCa32.
SLAC1 also differs radically from other structurally characterized anion channels and
transporters. These include the VDAC1 voltage-gated anion channel from mitochondrial
outer membranes, which has a porin-like beta-barrel structure34-36, and a light-driven
halorhodopsin chloride pump, which has a transmembrane conductance pathway similar to
that of the proton-pumping pore of bacteriorhodopsin37. Although its channel structure is
still only known by homology to other ABC transporters, CFTR is another obviously
distinct chloride channel38. Cys-loop receptors also include anion channels39, and these are
similar to SLAC1 in having five-helix pores25, but here selectivity is governed by charged
groups at the entrance to the pore, which distinguish the anion-selective GABAA and
glycine receptors from the cation-selective acetylcholine and serotonin 5HT3 receptors39.
Finally, the recently identified TMEM16A gene for calcium-activated chloride channels40-42
appears to encode an 8-TM protein that is again distinct from SLAC1.
Stomatal guard cells show both rapidly activated (R-type) and slow (S-type) anion channel
activity43. Although slac1 guard cells have very defective S-type activity, their R-type
currents are normal4. Guard cell protoplasts from the slac1-2 mutant abnormally accumulate
Cl−, K+, malate and fumarate3, whereas SLAC1 shows negligible malate conductance7. As
for SLAC1-associated K+ movements, other channels or transporters must be responsible for
SLAC1-associated malate movements. Recent studies indicate that AtALMT12, an
aluminium-activated malate transporter (ALMT) family member, is a malate-dependent R-
type anion channel44 needed for stomatal closure45.
Conclusions
We find that many functional properties of the plant SLAC1 anion channel are explained
well by the structure of an uncharacterized bacterial TehA protein that has been associated
with tellurite resistance. SLAC1 and TehA belong to distinct subfamilies within one branch
of a larger SLAC1 superfamily, but AtSLAC1 and HiTehA are sufficiently similar (19%
sequence identity) that the SLAC1 homology model is predictive for function, including a
verified placement of the identifying slac1-2 mutation G194D and a phenylalanine gate. One
remaining puzzle concerns the structural change that activating phosphorylation elicits in
SLAC1, and another puzzle is the biochemical role of the TehA homologs in bacteria. In a
companion paper26, we examine functional and structural properties of TehA in bacteria,
showing that it is anion channel, although actually not conferring tellurite resistance, and
identifying a mutant variant with properties suggestive of an activated state. Thus, SLAC1
and TehA likely represent a large family of selective anion channels controlled by
environmental stimuli.
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METHODS
Selection of target sequences
TehA from E. coli (EcTehA) was a centrally selected sequence for expansion as a
NYCOMPS targeted family. Target selection criteria utilized at NYCOMPS are described in
details elsewhere46. Briefly, the EcTehA sequence was run against a dataset of about 40,000
predicted alpha helical integral membrane protein sequences from prokaryotic genomes
(NYCOMPS98 dataset46) using PSI-BLAST50. Sequences that matched EcTehA with an E-
value lower than 10−3 in an alignment extending over at least 50% of both predicted TM
regions and passing our post-seed-expansion filtering criteria46 were passed to the protein
production pipeline.
Protein expression screening
Full-length homologs from the following 38 species, including 2 sequences each from 5 of
these, were amplified from genomic DNA by PCR: Thermoplasma acidophilum,
Lactococcus lactis subsp. lactis Il1403, Streptococcus pyogenes M1 GAS, Streptococcus
pneumoniae TIGR4, Haemophilus influenzae Rd KW20, Methanocaldococcus jannaschii
DSM 2661 (2), Escherichia coli K12, Salmonella typhimurium LT2, Clostridium
perfringens ATCC 13124, Xanthomonas campestris pv. campestris str. ATCC 33913,
Streptococcus agalactiae 2603V/R, Shewanella oneidensis MR-1, Streptococcus mutans
UA159, Archaeoglobus fulgidus DSM 4304, Vibrio parahaemolyticus RIMD 2210633 (2),
Pseudomonas syringae pv. tomato str. DC3000 (2), Enterococcus faecalis V583,
Pyrococcus horikoshii OT3, Bordetella bronchiseptica RB50, Streptomyces coelicolor A3,
Corynebacterium glutamicum ATCC 13032, Picrophilus torridus DSM 9790, Acinetobacter
sp. ADP1, Vibrio fischeri ES114, Pseudomonas fluorescens Pf-5, Sulfolobus acidocaldarius
DSM 639, Colwellia psychrerythraea 34H, Neisseria meningitidis MC58, Rhodobacter
sphaeroides 2.4.1, Vibrio cholerae O1 biovar eltor str. N16961 (2), Marinobacter aquaeolei
VT8, Acinetobacter baumannii ATCC 17978, Klebsiella pneumoniae subsp. pneumoniae
MGH 78578, Streptomyces avermitili MA-4680, Bordetella parapertussis 12822 (2),
Streptococcus thermophilus LMG 18311, Salmonella enterica subsp. enterica serovar
Paratyphi A str. ATCC 9150, and Anaeromyxobacter dehalogenans 2CP-C.
Selected cDNAs were cloned into a modified pET vector (Novagen, Inc) that fuses a FLAG
and deca-histidine tag at the C-terminus, which are cleavable by TEV protease. Proteins
were expressed in E. coli BL21(DE3) plysS by a high throughput format (0.6ml in a deep
well block) and purified after lysis by sonication using metal affinity purification in a buffer
containing N-Dodecyl-β-D-Maltopyranoside. Samples were passed over an analytical size
exclusion column in 12 different detergent-containing mobile phases, which included N-
dodecyl-β-D-maltopyranoside (DDM), N-decyl-β-D-altopyranoside (DM), N-nonyl-β-D-
altopyranoside (NM), N-octyl-β-D-altopyranoside (OM), N-octyl-β-D-Glucopyranoside
(OG), N-nonyl-β-D-Glucopyranoside (NG) and lauryl dimethyl amine oxide (LDAO).
Multi-angle light scattering with refractive index detection was used to analyze the
oligomeric state51. The E. coli and H. influenzae proteins were judged to be monodisperse
and stable and were passed to scale up.
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Scaled-up production and purification
For scale up, in brief, transformed BL21 plysS cells were grown at 37°C in 2× YT media to
OD 0.6-0.8 after being inoculated with 1% of the overnight culture. The culture was induced
with 0.4mM IPTG and continued to grow at 37 °C for another 4 hours. The cells were
harvested by centrifugation and stored at −80°C before use. Selenomethionyl (SeMet) TehA
was expressed in a similar way, but using containing SeMet in place of methionine in
defined minimal media. Cells were resuspended in a buffer containing 50 mM Tris-HCl (pH
8.0) and 200 mM NaCl and lysed using a French Press with two passes at 15-20,000 psi.
Cell debris were removed by centrifugation at 10,000g for 20min, and the membrane
fraction was isolated from that supernatant by ultra-centrifugation at 150,000g for 1 hr.
The membrane fraction was homogenized in a solubilization buffer containing 50 mM Tris
(pH 8.0) and 200 mM NaCl, and incubated with final concentration of 1% (w/v) dodecyl-β-
D-maltopyranoside (DDM, Anatrace, Inc) for 1 hr at 4°C. The non-dissolved matter was
remove by ultracentrifugation at 150,000g for 30 min, and the supernatant was loaded to a
5ml Hitrap Ni2+-NTA affinity column (GE Healthcare), pre-equilibrated with the same
solubilization buffer supplemented with 0.05% DDM. After 20 column volume buffer wash,
the protein was eluted with 250mM imidazole in the solublization buffer. The Flag and 10-
His tags were removed by adding super TEV at 1:100 mass ratio and incubating at 4°C
overnight. Tag removal was confirmed by SDS-PAGE, and the resulting sample was
concentrated to around 10mg/ml. Preparative size exclusion chromatography was carried out
on a Superdex-200 column for further purification, removal of TEV protease and the
cleaved tag, and for buffer and detergent exchange. The gel-filtration buffer contained 10
mM Tris (pH 8.0), 200 mM NaCl, 1mM EDTA, 0.5 mM Tris [2-carboxyethyl] phosphine
(TCEP), and 2×CMC of detergent. In the case of HiTehA, the protein was well behaved and
stable in nearly all tested detergents, and we have purified it from DDM, DM, NM, OM, OG
and LDAO.
Protein characterization
We performed N-terminal amino-acid sequencing of purified HiTehA and EcTehA prior to
TEV protease treatment. Results from these analyses proved that true initiating methionine
residue is located 14 residues after one annotated as N-terminal. The intervening nucleotide
sequence contains a Shine-Delgarno sequence.
For cross-linking experiments, purified HiTehA was incubated with 10 mM disuccinimidyl
glutarate at room temperate for 30 min, and 100mM Tris-HCl pH 7.5 was added to stop the
reaction. The incubated sample when run on a 8-25% gradient SDS-PAGE gel showed a
ladder consistent with a trimeric structure.
Crystallization and data collection
Purified protein was concentrated to ~10mg/ml for initial crystal trials in a Mosquito robot
with commercial screens from Hampton research, Emerald Biosystems and Molecular
Dimension. We obtained crystals of HiTehA from protein in detergent DDM, DM, NM,
OM, OG and LDAO, but only those from LDAO and OG gave diffraction to beyond 4Å
spacings. Crystals that proved useful were all grown at 4°C using the sitting-drop vapor
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diffusion method. After extensive optimization we reached conditions supporting very high
resolution. Wild-type HiTehA and most of its variants crystallized from 1mM ZnSO4,
50mM Hepes-Na pH 7.8 and 28% PEG600. HiTehA F262A crystallized from 200mM
Li2SO4, 100mM glycine pH9.3 and 33% PEG400). Addition of 10mM spermidine as an
additive helped to obtain slightly better diffracting crystals. Cryoprotection was achieved by
adding 5% ethylene glycol or PEG400 to the crystallization solution.
Structure determination and refinement
Native and SeMet SAD data set were collected at NSLS beamline X4A and processed using
the software HKL200052. Crystals of HiTehA grew in space group R3 with a=b=c = 85.0Å
and α=β=γ = 93.5°. All subsequent manipulations were done in the hexagonal setting of this
space group with a=b= 96.0Å and c = 136.7Å. The asymmetric unit contains one TehA
protomer and 65% solvent volume. The structure was determined at 2.0Å resolution from
single-wavelength anomalous diffraction (SAD) using selenomethionine-substituted protein
crystals. Assessment of data quality for phasing, location of heavy atom sites and initial
phases were calculated using the HKL2MAP interface to SHELX programs53.
All the secondary structure elements were clearly visible in the experimental electron
density map. Automatic model building was done in Arp/wArp54 and completed manually
in the program COOT55. The model was refined against native data at 1.20Å resolution
using the program Refmac5.5 in CCP456, with anisotropic B factor restrained refinement
applied. Subsequent structural analyses of mutant variants were refined as isomorphous
structures.
Site-directed mutagenesis
Site-directed mutants were constructed using the QuikChange Site-Directed Mutagenesis Kit
(Stratagene, La Jolla, CA 92037) and expressed from pET vectors in E. coli BL21(DE3)
plysS cells as for the wild-type protein.
Electrophysiology
All constructs were cloned into plasmid pGHME2, linearized, and transcribed into cRNA
using T7 polymerase (mMessage mMachine, Ambion). Oocytes were injected with 50 nl of
cRNA solution each, at a constant concentration of 0.5 mg/ml for HiTehA, or 0.5 mg/ml for
AtSLAC1 constructs, with or without 0.5 mg/ml of AtOST1 constructs and recorded in
voltage-clamp experiments 2 days after injection. For mixed expression experiments, 25nl of
cRNA solution was injected for each AtSLAC1 component. Two-electrode voltage clamp
recordings were performed to measure HiTehA or AtSLAC1 currents as described6,7. The
pipette solutions contained 3 M KCl. For voltage-clamp current recordings, the bath solution
contained 1 mM MgCl2, 1 mM CaCl2, 10 mM Mes/Tris (pH 5.6) and 30 mM CsCl; For
anion selectivity measurements, the bath solutions contained 50 mM Cl−, NO −3, malate, or
30 mM SO 2-3 sodium salts, 45 mM Na-gluconate, 1 mM Ca-gluconate2, 1 mM Mg-
gluconate2, 1 mM K-gluconate, as well as 10 mM Tris/Mes (pH 5.6). Osmolality was
adjusted with D-mannitol to 220 mOsmol/kg. The bath electrode was a 3M KCl agar bridge.
The testing potentials were +50 to −110 or −130 mV in 20 mV decrements with 7.5 s
duration. The holding and return potentials were 0 mV with a 1.45 s or 2 s, respectively. I-V
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relations of HiTehA or AtSLAC1 were generated from the current reached after 0.5 s in
testing potentials. The Goldman-Hodgkin-Katz equation was applied to estimate
permeability ratios for monovalent ions as described6. For divalent anions, the permeability
ratios were derived according to Fatt and Ginsborg57.
Bioinformatic analysis of SLAC-related proteins
Sequences related to SLAC1 were analyzed comprehensively by PSI-BLAST50. Searches at
E<10−3 starting from five disparate homologs each identified a common pool of over 900
proteins, which when pooled were used for sub-classification into families and subfamilies.
Details of these analyses are reported in footnotes to Table S1.
Molecular figures were produced in PyMOL58.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We thank Filippo Mancia, Larry Shapiro and Ming Zhou for helpful discussions; Patricia Rodriguez and Arianne
Morrison for help with cloning and expression; Jonah Cheung, Mark Collins, Chris Min, Sheng Ye and Zhen Zhang
for advice in protein chemistry and crystallography; John Schwanof and Randy Abramowitz for help with
synchrotron experiments; John Riley for help with Xenopus oocyte injections; and Min Su for advice on the PSI-
BLAST analysis of SLAC relatives. This project was supported in part by an award to the New York Consortium
on Membrane Protein Structure (NYCOMPS) from the NIGMS Protein Structure Initiative. Beamline X4A at the
National Synchrotron Light Source (NSLS), a DOE facility at Brookhaven National Laboratory, is supported by the
New York Structural Biology Center.
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Figure 1. Sequence analysis for the SLAC1 superfamily
a, Family tree. The presentation was computed by the program COBALT47 from
representative subfamily sequences (Table S1), including Arabidopsis thaliana SLAC1 for
SF1Ai, Haemophilus influenzae TehA for SF1Ci, Escherichia coli TehA for SF1Cii, Vibrio
parahaemolyticus for SF2A, Staphylococcus aureus for SF2B, Aspergillus fumigatus Ssu1
for SF3A, and Schizosaccharomyces pombe Mae1 for SF3B. b, Structure-based sequence
alignment of TehA from Haemophilus influenzae (HiTehA) and SLAC1 from Arabidopsis
thaliana (AtSLAC1). The TehA structure has been used to restrict sequence gaps to inter-
helical segments. Superior coils define extents of the HiTehA helical segments; red letters
mark residue identities; red boxes are drawn for residues that are >95% identical within the
plant subfamily SF1A for AtSLAC1 or within the TehA subfamily for HiTehA; red
diamonds mark HiTehA residues that line the central pore; and the colored inferior bar
encodes ConSurf sequence variability48 for the SF1 family of 204 non-redundant proteins.
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Figure 2. Crystal structure of HiTehA and homology model of AtSLAC1
a, Electron density distribution from the HiTehA crystal structure at 1.2Å resolution. The
map has (2Fo-Fc) coefficients based on the superimposed model. Contours are at 2.5σ. b,
Ribbon diagram of the HiTehA trimer. Each protomer is colored spectrally from blue at its
N-terminus to red at its C-terminus. c, DelPhi49 electrostatic potential at the extracellular
surface. Electronegative and electropositive potential are colored in degrees of red and blue
saturation, respectively. d, Electrostatic potential49 at the intracellular surface. e, Ribbon
diagram of an HiTehA protomer viewed from outside the membrane. The ribbon is colored
spectrally as in 2b. f, Ribbon diagram of an HiTehA protomer viewed from within the
membrane from below the view of e. g, Surface of a homology model of AtSLAC1, viewed
as in f, and colored by sequence variability48. h, Surface of AtSLAC1 as in g, but colored by
electrostatic potential49.
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Figure 3. Putative structure of the SLAC1 conductance pore
a, Cross-section through the homology model of AtSLAC1. The model is viewed as in 2i,
with the electrostatic potential49 shown on the external surface of the molecular envelope.
The side chain of Phe450 is shown as a stick model (red) on the backbone ribbon colored
yellow. b, Cross-section as in a, but colored by surface conservation48 as in 2h. c, Pore-
lining residues in the SLAC1 homology model. A cylinder model (left), spectrally colored as
in 2b, provides a key for viewing the rolled-open structure (left) with pore-lining residues of
AtSLAC1 shown on the TModd helices. d, Ribbon diagrams of HiTehA TM9 (left) and TM7
(right) viewed from within the conductance pore. The side chains of Pro207 and Phe262 are
shown as well as a kink-stabilizing water HOH25 that is coordinated by the NH of Gly263
and by C=O groups of Gly202 and Ala259. Density contours are shown for the water
molecule.
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Figure 4. Ionic conductance measurements
a, Typical microelectrode voltage-clamp current traces from oocytes injected with various
channel cRNAs. Left column, oocytes injected with cRNAs encoding wild-type HiTehA
channels (WT), or F262A, G15D/F262A or G15D mutants. Right two columns, oocytes
injected with cRNAs for WT AtSLAC1, or F450A, G194D/F450A or G194D mutants, with
or without co-injection of AtOST1. Dotted lines represent zero current levels. Extracellular
solution contained 30mM CsCl. Schematic keys at the left place phenyl and/or aspartyl
gates. b, Effects of gating residue mutations. Mean chloride currents, measured at −90 mV,
are shown comparing WT HiTehA and its mutant series F262A,G,T,V,L with WT AtSLAC1
and its corresponding series F450A,G,T,V,L, both alone and co-expressed with AtOST1.
Full I-V relations are shown in Fig. S5. c, Effect of gating residues on relative AtSLAC1
anion permeabilities. Relative permeabilities (P[X]/P[Cl]) for chloride, nitrate, sulfite and
malate of WT, F450A and F450T SLAC1 channels were measured from the change in
current reversal potential with Cl− or anion X− as the sole permeant anion in the bath
solution (Methods, Table S6).
Chen et al.
Page 19
Nature. Author manuscript; available in PMC 2013 January 18.
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Figure 5. Structural features at the SLAC1 homolog gate
a,b, Cross-sections through the conductance pores of HiTehA F262A and HiTehA G15D.
The view and presentations are as in 3a, except that helices are colored purple. c, Molecular
basis for conformational strain in gating residue Phe262 of HiTehA. Helices TM1 (left),
TM9 (center) and TM7 (right) are viewed from within the pore and presented as ribbon
diagrams with selected side chains drawn in stick representation. The local low-energy
conformation for the phenyl ring (χ2 = 90°) is shown in thin lines with short contacts
indicated in dashed lines: Leu18 Cδ2 – Phe262 Cδ1, d = 2.4 Å; Val210 Cγ2 – Phe262 Cε2, d
= 2.8 Å. d, Conformational shifts consequent to release of strain in gating residue Phe262.
Cα backbone structures of F262A, F262V and F262L HiTehA were first superimposed onto
WT HiTehA, and residues 258-266 were drawn in stereo as superimposed. All backbone
atoms are shown for peptides 262±1, but only Cα atoms are shown otherwise. The WT
backbone and phenyl group are green; other backbone are all magenta; side chains of
Ala262, Val262 and Leu262 are cyan, blue and red, respectively; oxygen-directed bonds are
red and nitrogen-directed bonds are blue. e, Typical microelectrode voltage-clamp current
traces from oocytes injected with wild-type (WT) or F450L AtSLAC1 cRNA. Experimental
conditions and displays are as in 4a.
Chen et al.
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|
3M7N
|
archaeoglobus fulgidus exosome with RNA bound to the active site
|
Quantitative analysis of processive RNA degradation
by the archaeal RNA exosome
Sophia Hartung1, Theresa Niederberger1, Marianne Hartung2, Achim Tresch1 and
Karl-Peter Hopfner1,*
1Center for Integrated Protein Sciences and Munich Center for Advanced Photonics at the Gene Center,
Department of Biochemistry, Ludwig-Maximilians-University Munich, Feodor-Lynen-Strasse 25, 81377 Munich
and 2General Electric - Global Research, Freisinger Landstrasse 50, 85748 Munich, Germany
Received February 9, 2010; Revised March 18, 2010; Accepted March 21, 2010
ABSTRACT
RNA exosomes are large multisubunit assemblies
involved
in
controlled
RNA
processing.
The
archaeal exosome possesses a heterohexameric
processing
chamber
with
three
RNase-PH-like
active sites, capped by Rrp4- or Csl4-type subunits
containing RNA-binding domains. RNA degradation
by RNA exosomes has not been studied in a quan-
titative manner because of the complex kinetics
involved, and exosome features contributing to effi-
cient RNA degradation remain unclear. Here we
derive a quantitative kinetic model for degradation
of a model substrate by the archaeal exosome.
Markov Chain Monte Carlo methods for parameter
estimation allow for the comparison of reaction
kinetics between different exosome variants and
substrates.
We
show
that
long substrates
are
degraded in a processive and short RNA in a more
distributive manner and that the cap proteins influ-
ence degradation speed. Our results, supported by
small angle X-ray scattering, suggest that the
Rrp4-type cap efficiently recruits RNA but prevents
fast RNA degradation of longer RNAs by molecular
friction,
likely
by
RNA
contacts
to
its
unique
KH-domain. We also show that formation of the
RNase-PH like ring with entrapped RNA is not
required for high catalytic efficiency, suggesting
that the exosome chamber evolved for controlled
processivity, rather than for catalytic chemistry in
RNA decay.
INTRODUCTION
The eukaryotic and archaeal RNA exosomes and their
distant relative, the bacterial degradosome, are large
multiprotein assemblies that function as central cellular
RNA
processing
and
degradation
machineries.
The
RNA exosome was originally found in yeast as an essen-
tial protein complex with 30 ! 50 exonuclease activity.
First, identified for the 30-processing of the yeast 5.8S ribo-
somal RNA (1), the yeast RNA exosome subsequently
turned out to be important for the trimming and degrad-
ation of the 30-end of several nuclear RNA precursors (2).
In addition, the exosome was shown to be also active in
the cytoplasm by controlling mRNA turnover (3), and by
its implication in various mRNA surveillance pathways
like the non-sense-mediated and the non-stop decay
pathways (4–7). Due to its involvement in all the different
RNA processing and surveillance pathways the exosome is
apparently one of the central exonucleases of a yeast cell
[for reviews see for instance (8,9)].
Structural homologues of the yeast exosome were sub-
sequently identified in humans, previously known as the
PM-Scl
(polymyositis–scleroderma
overlap
syndrome)
complex, and in archaea (10–12). A variety of structural
studies revealed a conserved architecture of exosome like
complexes (13–18): exosomes consist of nine conserved
core subunits, six RNase PH type subunits and three
subunits with S1 and KH or zinc-ribbon domains. The
six RNase-PH like domains form a ring, arranged as
trimers of pseudo-dimers. In archaea, the ring is formed
by three (archaeal)aRrp41:aRrp42 dimers, while human
and yeast exosomes contain six different RNase PH type
subunits.
The archaeal exosome possesses a central chamber
within the RNase PH ring which contains three phos-
phorolytic active sites. The actual active site is located in
the aRrp41 subunits, but the whole aRrp41:aRrp42 dimer
is involved in positioning the RNA. These sites degrade
single-stranded RNA (ssRNA) in a phosphate dependent
manner in 30 ! 50 direction. They also catalyse the reverse
reaction of adding nucleoside diphosphates to the 30-end of
RNA (13), liberating inorganic phosphate. In archaea, this
*To whom correspondence should be addressed. Tel: +49 89 2180 76953; Fax: +49 89 2180 76999; Email: hopfner@lmb.uni-muenchen.de
5166–5176
Nucleic Acids Research, 2010, Vol. 38, No. 15
Published online 14 April 2010
doi:10.1093/nar/gkq238
The Author(s) 2010. Published by Oxford University Press.
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/
by-nc/2.5), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.
activity has been attributed to formation of poly-A-rich
tails on RNA (19). A proposed RNA entry pore at one
side of the chamber restricts entry to mostly unstructured
ssRNA, providing an explanation for controlled RNA deg-
radation. Furthermore, three Csl4 or Rrp4 type putative
RNA recognition subunits are located on top of the
(Rrp41:Rrp42)3 ring and frame the proposed RNA entry
pore. Current models suggest that these domains recognize
RNA substrates and help to funnel them into the process-
ing chamber.
Although the human exosome is structurally related to
the archaeal complex, including S1 and KH domain con-
taining subunits (Csl4, Rrp4 and Rrp40), it has lost
phosphorolytic activity (14). Instead, it gained additional
ectopic subunits: the hydrolytic RNase Rrp44 (20,21)
has both exonucleolytic and endonucleolytic activities,
located in RNB and PIN domain subunits, respectively
(14,20,22–24). A second hydrolytic RNase (Rrp6) was
identified as transient part of the nuclear complex (10).
Recent results indicate that despite the ectoptic placement
of the nuclease active sites, RNA is still threaded through
the nuclease deficient RNase PH type ring (25).
A variety of groups have biochemically observed
processive
RNA
degradation,
in
particular
for
the
archaeal
exosome.
From
structural
studies,
it
was
proposed that RNA is channelled through an entry pore
between the S1 domains of aCsl4 or aRrp4 trimers into the
processing chamber, where the 30-end of the RNA is pos-
itioned in one of the three phosphorolytic active sites,
subsequently degrading RNA base-per-base (16,26,27).
The presence of the cap proteins Csl4 and Rrp4 in
general
increases
the
degradation
efficiency
of
the
exosome, but it is unclear how they do so. For instance,
if the cap proteins recruit RNA, one would expect an
increase in the general binding affinity. However, once
RNA has entered the processing chamber, high affinity
binding to the ectopic domains should slow down
processive degradation. Another mechanism that is not
yet understood is why processivity depends on the length
of the RNA molecules (28). To address these questions
and to develop means to quantitatively analyse processive
degradation, we performed quantitative high-resolution
RNase degradation activity assays with different variants
of the Archaeoglobus fulgidus exosome. We evaluated dif-
ferent kinetic models and developed a Markov Chain
Monte Carlo (MCMC) analysis to fit the model to the
data and derive appropriate rate constants of individual
RNA degradation steps. Our data identify different struc-
tural contributors to processivity, suggesting that ectopic
RNA-binding domains, the entry pore and the active site
are different contributors to processive degradation. The
methods
should
be
easily
applicable
also
to
other
processive enzymes, including the hydrolytic nucleases of
the eukaryotic exosomes.
MATERIALS AND METHODS
Protein expression and purification
The Archaeaoglobus fulgidus RNA exosome with different
Rrp4 and Csl4 caps, derivatives and mutants were
expressed and purified as described (29). Site directed mu-
tations
were
introduced
using
the
QuickChange
Site-directed Mutagenesis Kit (Stratagene) and verified
by sequencing. Oligonucleotide sequences are provided
in the Supplementary Table S2.
Crystallization and structure determination
An amount of 120 mM Csl4-exosome (Csl4:Rrp41:Rrp42)3
or its Y70ARrp42 mutant (= 27 g/l) were incubated with
400 mM RNA (3.3-fold excess, 6-mer CCCCUC) for
10 min on ice. Protein:RNA complexes were crystallized
by sitting drop vapour diffusion technique by mixing 1 ml
protein and 1 ml of reservoir solution (0.1 M NaAcetate,
pH 4.6, 30% 3-Methyl-1,5-pentadiol (MPD), 100 mM
NaCl) at 20C. Datasets were recorded at the ID-14-2
beamline (ESRF, Grenoble, France) to 2.4 A˚
(wild-type
exosome) and at the PX I beamline (SLS, Villigen,
Switzerland) to 3.0 A˚ (Y70ARrp42 mutant) and processed
with X-ray Detector Software (XDS) (30). A model of the
apo-Csl4-exosome complex (29) was used as a search
model for molecular replacement using PHASER (31).
Refinement to 2.4 A˚ and 3.0 A˚ , respectively was performed
with CNS (32) and PHENIX (33). In the additional
electron density RNA nucleotides were positioned using
COOT (34). Refinement of the complete complexes was
followed by iterative cycles of manual model completion
with COOT and positional and B-factor refinement with
CNS (Supplementary Table S1).
Small angle X-ray scattering
For small angle X-ray scattering (SAXS) studies, the
(Rrp41:Rrp42:Rrp4)3 complex was purified as described
above. To purify the exosome with endogenously bound
Escherichia coli RNA the protocol was modified as
follows: RNA was not washed offwith high salt, and in
all buffers the salt concentration was 250 mM or lower.
After the Ni-NTA affinity chromatography, the complex
was loaded on an anion exchange column to remove
unbound nucleic acids and the procedure was repeated
to assure the total removal of free RNA. Not until only
one distinct peak was eluted, the fractions were pooled,
concentrated and flash frozen. The apo-complex was
measured at 5, 10 and 15 mg/ml and the RNA complex
was
concentrated
to
an
absorption
at
280 nm
of
A280 = 55 and measured in a 1: 0, 1: 1 and 1: 2 dilution
to evaluate the concentration dependency of scattering.
Both complexes did not show concentration dependent
aggregation and were not affected by long exposure to
high-energy X-rays. SAXS data collection was performed
in 20 mM Tris pH 7.4 and 200 mM NaCl buffer at the
SIBYLS beamline (Advanced Light Source, Berkeley,
CA, USA) (35). The radius of gyration was calculated
using the Guinier plot in the linear region (constraint: s
Rg <1.3) and the calculation of the pair distribution
function was done with GNOM within PRIMUS (36).
Ab initio modelling of the solution structures was done
with GASBORp (37) and more than 10 identically
calculated models were aligned and averaged using
DAMAVER and SUPCOMB (38). For analysis of the
Nucleic Acids Research, 2010, Vol. 38, No. 15
5167
bound RNA, the protein was separated from the RNA by
running the complex on a denaturing 6 M urea and 20%
polyacrylamide gel and elution of the RNA from the gel.
The pelleted RNA was sent to Vertis Biotechnologie AG,
where the sample was poly(A)-tailed using poly(A) poly-
merase followed by ligation of an RNA adapter to the
50-phosphate of the small RNAs. First-strand cDNA syn-
thesis was then performed using an oligo(dT)-adapter
primer and M-MLV H- reverse transcriptase. The result-
ing cDNAs were PCR-amplified to 20–30 ng/ml in 19
cycles using standard Taq DNA polymerase. We cloned
the cDNA products with EcoRV into pET21 vectors,
transformed and amplified the plasmids and isolated and
sequenced single clones.
Cross-linking
Site-specific crosslinking of the K37CRrp41:D143CRrp42
mutant was performed with a HBVS (1,6-Hexane-bis-
vinylsulfon) crosslinker. The crosslinking reaction was
performed with a 100-fold excess of crosslinker under
oxygen-free conditions in a glove-box. We removed
crosslinked
protein
from
non-crosslinked
protein
complexes using a Superose 6 size-exclusion column,
equilibrated
with
a
running
buffer
containing
4 M
guanidinium chloride. Protein from the peak correspond-
ing to a crosslinked Rrp41/Rrp42 dimer was refolded in
50 mM Tris (pH 7.4), 200 mM NaCl, 500 mM arginine and
5% glycerol by dilution. The refolded protein was again
applied onto a Superose 6 column in 50 mM Tris (pH 7.4)
and 200 mM NaCl. Only the correctly refolded protein,
verified by the formation of a hexamer in size exclusion
chromatography, was used for further experiments. As a
control sample, the same complex without crosslinker was
partly denatured, purified and refolded in the same way as
the crosslinked protein.
RNase activity assays
We carried out RNase activity assays using 32P-labelled
poly(rA)-oligoribonucleotides with different lengths as
substrate (26). RNA was incubated with [g-32P] ATP
(Hartmann Analytics) and T4 polynucleotide kinase
(NEB)
for
45 min
at
37C
and
purified
by
using
MicroSpin G-25 columns (GE Healthcare). For each
reaction, the protein (30 nM for the Csl4- and the
Rrp4-capped
wild-type
exosome
and
the
interface
mutant; 60 nM for the cap-less exosome and the single
site mutants R65ERrp41 and Y70ARrp42; 120 nM for the
crosslinked cap-less exosome) was incubated with RNA
in buffer containing 20 mM Tris (pH 7.8), 60 mM KCl,
10 mM MgCl2, 10% glycerol, 2 mM DTT, 0.1% PEG
8000, 10 mM NaH2PO4 (pH 7.8) and 0.8 U/ml RNasin
(Promega) at 50C. Different time points were taken and
the reaction was stopped by adding one volume of loading
dye [0.75 g/l bromphenol blue, 0.75 g/l xylene cyanol, 25%
(v/v) glycerol, 50% formamide]. The reaction products
were
resolved
on
a
20%
polyacrylamide/6 M
urea
sequencing gel running at 50C and were analysed
by phosphorimaging (GE Healthcare). The gel bands
were
quantified
using
the
ImageQuant
Software
(GE Healthcare) and data analysis, simulation and
fitting was done with MatLab (Mathworks).
Models and kinetic data analysis
Kinetic models are shown in Figure 3A. They are
described by four parameters: association rate ka,i, dissoci-
ation rate kd,i, cleavage rate kc,i and polymerization rate
kp,i, one for each RNA of length i = 4, 5, . . ., 30. The cor-
responding set of differential equations that quantitatively
describe RNA degradation is shown in the supplement
Data (Chapter 1). Since the reaction takes place in an
excess
of
inorganic
phosphate
(10 mM
phosphate
compared to only 3.6 mM ADP at the time all RNA mol-
ecules are totally degraded), we may assume no polymer-
ization takes place, i.e. kp,i = 0 for all i. Consistently, we
saw no synthesis of longer RNAs in our reactions. To
obtain empirical estimates of the posterior parameter dis-
tribution, we implemented a MCMC approach based on
the Metropolis–Hastings algorithm. The key ingredients
are the likelihood function, the prior, and the proposal
distribution. The likelihood function penalizes the estima-
tion error produced by a given model. More precisely, it
penalizes the residuals, i.e. the deviation of the measured
RNA amounts at each time point from the amounts that
have been predicted from the current parameter set. We
assume that the residuals are independent realizations of
Gaussian distributions with zero mean. Since the vari-
ances of these Gaussians are not known a priori, we
assume a two-parameter error model with an additive
and a multiplicative error component which has been
proposed (39) in the context of spot quantification on
arrays. We initialize the error model very conservatively
(presuming
large
measurement
errors).
During
the
MCMC run, the error model is updated continuously by
replacing it with an empirical estimate derived from the
residuals that occurred in the Markov chain so far.
The prior encodes prior knowledge/assumptions on
the distribution of the parameters. It is sensible to
require the kinetic parameters to vary smoothly with the
RNA length i. This is made explicit by penalizing the
difference of two successive kinetic parameters kx,i+1 and
kx,i using a Gaussian prior on these differences. We em-
phasize that this does not impose any restrictions on the
absolute level of the parameter values. The comparison of
the parameter levels obtained by different experiments is
virtually unaffected by our prior choice and therefore
practically unbiased. The proposal distribution generates
a new parameter set as a candidate for the next MCMC
step that is based on the current parameter set. We simply
use a multivariate log-normal distribution with fixed
diagonal covariance matrix, which is centred at the
current parameter set.
It turns out that the parameters of the model as stated
above are not identifiable. We therefore fixed kd,i to
one global constant kd, whereas the association param-
eters ka,i are sampled individually. The parameters kc,i
are set equal to one length-independent parameter kc.
The details of this approach and its justification through
extensive simulations are given in the supplementary Data
(Chapters 2–4).
5168
Nucleic Acids Research, 2010, Vol. 38, No. 15
RESULTS
RNA is not degraded with constant velocity
Despite intense structural and biochemical research on
RNA exosomes, a kinetic model, quantitative analysis of
processive RNA degradation and a biochemical identifi-
cation of elements that contribute to processive degrad-
ation have not been studied, due to the complex kinetics
involved. To address these issues, we performed RNase
assays with 50-radioactively labelled 30-mer oligo(A)
RNAs and the A. fulgidus Csl4- (Csl4:Rr41:Rrp42)3 and
Rrp4-exosomes
(Rrp4:Rr41:Rrp42)3.
The
reaction
products and their time evolution were resolved on
a
denaturing
sequencing
gel
and
quantified
by
phosphorimaging (Figure 1), controls are shown in the
supplemental material (Supplementary Figures S4 and
S5). Several characteristic features of substrate degrad-
ation by exosomes are revealed:
First, RNA is not degraded at a constant speed, but
the degradation of substrate has several phases and is
distinct in different isoforms. In the Csl4 exosome
(Figure 1A), after a slower first processing step, longer
RNAs (>12–13 nt) are degraded very fast, seen by
the low amount of intermediates in this range; shorter
RNAs (<12–13 nt) are degraded slower and accumulate
first before they are further degraded. On the contrary,
the first processing step is faster in the Rrp4- than in the
Csl4-exosome (Figure 1B). However, oligo-rA substrates
>24 nt are degraded slower, intermediate substrates
(24–13 nts) faster, and RNAs <13 nt slower again.
This result is astonishing, considering homooligomeric se-
quences are used and the effect is consequently not
sequence
dependent.
In
addition,
the
unexpected
slow-fast-slow kinetics of the Rrp4 isoform reveals a
quite complex length dependency of RNA degradation
speeds.
Second, the final degradation product is a 3-mer.
Further
degradation
is
extremely
slow,
comparable
to spontaneous background hydrolytic cleavage under
the present conditions. We hypothesized that features
of the active site might interact specifically with the
fourth base at the 30-end. Previous structural analysis
with the Sulfolobus solfataricus exosome has shown that
at least 4 nt are stably bound in the phosphoropytic active
sites (26), but in the case of the Pyrococcus furiosus
exosome some nucleotides were recognized (16). To get
direct
structural
information
for
the
A.
fulgidus
exosome:RNA
interaction,
used
in
this
study,
we
crystallized
our
Csl4-exosome
with
a
6-mer
RNA
molecule (Figure 2; Supplementary Table S1). Four nu-
cleotides from the 30-end are clearly visible in the unbiased
Fo–Fc electron density, with weaker density for the two
additional nucleotides. Interestingly, the side chain of
Y70Rrp42
shows
p-stacking
with
the
fourth
base
(counting from the active site) and this seems to be a
conserved feature among archaeal exosomes (16,26).
This interaction specifically stabilizes the first 4 nt, while
RNA positions+5 and+6 behind Y70Rrp42 appear not to
be specifically recognized. To test the role of Y70Rrp42, we
determined
the
co-crystal
structure
of
the
Csl4-
exosome-Y70A mutant with a CCCCUC oligonucleotide.
In fact, we only see clear electron density for 4 nt in the
active site and the electron density at position+4 is weaker
and less defined compared to the wild-type. Thus,
the 3-mer as degradation end-product is likely the
cause of inefficient recognition of RNA’s with <4 nt at
the active site.
Figure 1. Visualization of RNase activity of the archaeal exosome on denaturing polyacrylamide gels: the input (I) is a 30-mer polyA RNA
radioactively labelled at the 50-end that is degraded from the 30-end to a final product (FP) of a 3-mer. Time points were taken in increasing
intervals [in minutes: 0:10; 0:20; 0:30; 0:40; 0:50; 1:00; 1:10; 1:20; 1:40; 2:00; 2:20; 2:40; 3:00; 3:30; 4:00; 4:30; 5:00; 5:50; 6:00; 6:30; 7:00; 7:30; 8:00;
9:00; 10:00; 12:00; 14:00; 16:00; 18:00; 20:00; 25:00; 30:00; 35:00; 40:00; (B) ends at 8:00 min]. RNA degradation does clearly not occur with constant
speed and the (Csl4:Rrp41:Rrp42)3 exosome (A) degrades RNA with a different time dependency than the (Rrp4:Rrp41:Rrp42)3 exosome (B).
Nucleic Acids Research, 2010, Vol. 38, No. 15
5169
A kinetic model for processive RNA degradation
by exosomes
To obtain a comprehensive picture of the exosomal RNA
decay, we need to analyse the reaction speeds in a quan-
titative manner. The amount of RNA as function of time
of intermediate i of an rA n-mer may be described by
several rate constants (Figure 3A): an association rate
constant ka,i of the 30-end of RNA to the active site; a
corresponding dissociation rate constant kd,i; a rate of for-
mation of intermediate i by cleavage of intermediate i+1,
Figure 3. Three different models to describe the kinetics of RNA degradation by the exosome were tested: (A) scheme for the general kinetic
model, which includes cleavage and polymerization rates kc and kp as well as association and dissociation rates ka and kd for all RNAs from 30–4 nt.
(B–D) Quantified concentrations of RNA intermediates from Figure 1A, along with least square fits to different kinetic models. (B) Strict processivity
considers only 27 different cleavage rates kc,30 –kc,4. (C) cleavage-and-polymerization considers 27 different cleavage rates kc,30 –kc,4, 27 different
polymerization rates kp,30–kp,4 and one initial association rate ka,30 (=55 rates). With models (C) and (B), no reasonable fit could be obtained. (D) By
including association, dissociation and cleavage and making rational simplifications (see text) we can convincingly fit the data with a model con-
taining 28 different rate constants.
Figure 2. Crystal structure of 6-mer RNA bound to the active site of the archaeal exosome. Rrp41 is shown in light and Rrp42 in dark green. The
2Fo–Fc electron density is contoured at 1.0s and only shown for the RNA and the side chain of Y70Rrp41. (A) In the wild-type exosome Y70 is
stacking with the fourth base of the bound RNA, and only weak density can be seen for the fifth and sixth base. (B) Electron density for the fourth
base of the RNA is much weaker in the Y70ARrp41 mutant compared to the wild-type and no density can be detected at this contour level for
additional nucleotides.
5170
Nucleic Acids Research, 2010, Vol. 38, No. 15
kc,i+1; and by adenylation (polymerization) of intermedi-
ate i1, kp,i-1; a rate of disappearance of intermediate i by
cleavage of i, kc,i and by adenylation of i, kp,i. The system
kinetics is then given by a set of differential equations
(Supplementary Data).
However, it is possible that this more general model can
be further simplified. For instance, we likely can neglect
adenylation (kp,i = 0) because our reaction conditions
contain 10 mM phosphate compared to only 3.6 mM
ADP
at
the
time
all
RNA
molecules
are
totally
degraded,
strongly
shifting
the
reversible
reaction
towards degradation. In addition, the exosome might
be strictly processive, i.e. association and dissociation
rates of RNA intermediates are negligible compared to
the
cleavage
rates
(ka,i = kd,i = 0).
Furthermore,
all
cleavage rates may be independent of the length of
RNA, because they could be a local active site property
(kc,i = kc,j). Hence we analysed three simplified models
(Figure 3B–D). Once initial values for the rate constants,
enzyme concentration and RNA substrate concentration
(rA 30-mer) are provided, this corresponding set of differ-
ential equations can be used to calculate the concentra-
tions of all RNA intermediates over time. We then
minimized the resulting least square deviations between
the
calculated
and
experimental
concentrations
of
reaction intermediates by optimizing the rate constants
using the ‘fminsearch’ parameter optimization procedure
as implemented in Matlab.
Using the ‘strict processivity’ model with 27 independ-
ent
variables
(ka,i = kd,i = kp,i = 0)
(Figure
3B),
we
obtained no reasonable fit of the experimental data. A
second model including the adenylation reaction (55 inde-
pendent variables, ka,i = kd,i = 0) could also not properly
interpret the data (Figure 3C). Thus, simply adding more
parameter does not automatically lead to reasonable fits
and the RNA degradation activity cannot be convincingly
explained by strict processivity. Consequently, we added
association and dissociation of RNA intermediates to the
equations and used the following alternative simplifica-
tions: (i) adenylation is omitted (kp,i = 0; see above); (ii)
the same cleavage rate is used for all RNA molecules
(kc,i = kc,j),
i.e.
cleavage
rate
is
a
local
active
site
property
and
not
dependent
on
RNA
length.
We
estimated starting values for kc and validated this simpli-
fication from analysis of the initial exponential decay of
RNAs substrates with different initial lengths (data not
shown). (iii) Due to our experimental approach, we
cannot experimentally distinguish between bound and
free RNA since the gel bands represent the sum of free
and exosome-bound RNA intermediates of length i. For
that reason, we cannot reconstruct dissociation-, associ-
ation- and cleavage rate constants independently of each
other. Consequently, we do not treat the association and
the dissociation rate constants independently, but analyse
the ratio of ka,i/kd,i by setting kd,i’s to a constant low value,
leaving ka,i free to vary. Variation of the value for kd did
not
result
in
significant
changes
in
the
analysis
(Supplementary Data). These three reasonable simplifica-
tions leave essentially one free parameter per intermediate
plus one overall cleavage rate constant. Although this
model has less degrees of freedom than the second
model (55 versus 28), it can convincingly interpret the ex-
perimental data for the both Csl4 and Rrp4 exosome
variants and most mutants (Figure 3D).
MCMC analysis of degradation
To address the problem of multidimensional parameter
fitting and to assess the variance in parameter esti-
mation, we established MCMC simulations. Because of
the difficulty in determination of separate values for
the single rate constants, we defined an RNA length-
dependent quantity vi
vi ¼ kc,i
Km,i
ð1Þ
with Km,i the Michaelis–Menten constant
Km,i ¼ kc,i+kd,i
ka,i
ð2Þ
vi is called ‘catalytic efficiency’ or ‘specificity constant’, as
it is a measure of the velocity of RNA intermediate i deg-
radation by the exosome. We are now in a position to test
exosome features important for vi. We observe that for the
Csl4 exosome, vi is highly dependent on the RNA length:
the initial RNA processing step, likely determined by the
initial association of RNA with the exosome, is generally
slow. Once RNA is bound, vi is large and relatively
constant for RNA lengths >13 nt. vi then progressively
decreases for RNA lengths <13 nt until the final 3-mer
appears (Figures 4B and 5A). This length dependency may
be explained by the exosome structure: RNA molecules
longer than 13 nt might still reach through the ‘neck’,
and this topological interaction will induce a higher
‘local concentration’ of RNA at the active site with
increased vi. Short RNAs, on the contrary, will lose this
contact and due to their smaller size more easily diffuse
out of the processing chamber, therefore decreasing vi.
To test this idea, we analysed the Y70A mutant of the
Csl4-exosome. The length profile of the catalytic efficiency
has a similar shape than for the wild-type, although the
catalytic efficiency is lower for all RNA intermediates
(Figure 5B). For RNAs >13 nt, the difference in vi is
2- to 3-fold (about one log unit). However, the drop in
vi for RNAs <13 nt is progressively more pronounced
compared to the wild-type and towards short RNAs
(<8 nt), the mutant is 20- to 150-fold (three to five log
units) slower than the wild-type. This is consistent with the
idea that for long RNAs the neck provides additional
interaction and thus overcomes in part the destabilizing
effect of Y70A. For shorter RNAs, the active site becomes
the sole attachment, leading to a rapid drop of catalytic
efficiency in the Y70A mutant.
We also analysed the ‘neck’ mutant R65E, which has
been shown to severely reduce exosome activity (16,26,27).
This mutant exhibited a substantially delayed onset of
degradation, presumably because RNA is unable to effi-
ciently enter the active site (data not shown). A likely
reason is the formation of non-productive RNA:protein
complexes with RNA trapped on the outside of the
exosome (29). At present, our model cannot deal with
Nucleic Acids Research, 2010, Vol. 38, No. 15
5171
this scenario and we could not convincingly include—as
only variant—the R65E mutant in the analysis. However,
the data of the analysis of R65E are provided in the sup-
plementary Figure S15 and following.
Role of exosome ring formation and ring dynamics for
RNA binding
Although the initial binding of RNA appears slow, it
seems unlikely that the 30-end directly finds its way
through the small hole in the neck. It is perhaps more
likely that the ring structure ‘breathes’—as observed e.g.
in hexameric helicases—and allows some lateral entry at
the neck. To explore this idea we analysed a crosslinked
exosome, where the ring is rigidified by three site specific
crosslinks, and a mutant that disrupts the ring structure
into Rrp41:Rrp42 pairs. We compared these isoforms with
the
corresponding
wild-type,
the
cap-less
hexameric
(Rrp41:Rrp42)3 ring (Figure 5C). From the structural
analyses, it was observed that the Rrp41 and Rrp42
subunits possess two interfaces. One interface is larger,
and characterized by contiguous b-sheets between Rrp41
and Rrp42 (40). The other interface is smaller, presumably
more dynamic, and was chosen for the crosslinking and
mutagenesis
analysis
(Supplementary
Figure
S2).
(Rrp41:Rrp42)3 exhibit a biphasic length dependence of
vi, similar to the Csl4-exosome. However, the catalytic
efficiency of (Rrp41:Rrp42)3 is 5- to 10-fold higher
Figure 5. Comparisons
of
the
catalytic
efficiency
vi
of
different
exosome variants versus RNA lengths: (A) differences in the cap
proteins influence catalytic activity. This is shown by comparison of
vi from the cap-less exosome (Rrp41:Rrp42)3 in magenta, the Csl4
capped exosome (Csl4:Rrp41:Rrp42)3 in red and the Rrp4 capped
exosome (Rrp4:Rrp41:Rrp42)3 in blue. (B) Tyr70Rrp42 close to the
active site is especially important to efficiently degrade small RNAs.
The wild-typ Csl4 exosome is shown in red and the Y70ARrp42 mutant
in green. (C) The role of the ring architecture and dynamics for cata-
lytic activity is shown by comparing wild-type cap-less exosome
(Rrp41:Rrp42)3 in magenta with the dimeric and open interface
mutant (Rrp41:Rrp42)1 and a rigidified crosslinked variant that is less
dynamic in yellow. A total of 1000 parameter sets have been randomly
drawn from the stationary phase of the Markov chain. Thus for each
RNA length and each timepoint, we obtained 1000 estimates whose
distribution is displayed by boxplots.
Figure 4. Catalytic efficiency vi for all RNA intermediates present
during the degradation of a 30-mer RNA by the Csl4-Rrp41-Rrp42
exosome was determined with MCMC simulations. (A) shows the
traceplot and (B) the final parameter set (burnin = 150 000). It can be
seen that the MCMC chains vary in convergence speed as well as in
variability. The boxplots in (B) illustrate the main advantage of the
MCMC approach: it not only offers a set of parameters that best
describe the measured data, but it also yields a posterior distribution
for each catalytic efficiency parameter and thus provides a more com-
prehensive summary of the data.
5172
Nucleic Acids Research, 2010, Vol. 38, No. 15
across the RNA spectrum than that of the Csl4-exosome,
indicating that the Csl4 cap subunits do not substantially
promote
degradation
of
this
model
substrate.
In
addition,
vi
drops
for
RNAs
<13 nt
even
for
the
(Rrp41:Rrp42)3 particle, indicating that the neck not
e.g. the S1 domains of caps are responsible for the
higher catalytic efficiency on longer RNAs.
To explore the effect of the hexameric ring formation,
we mutated Lys51 to Glu, located in the ‘smaller’ interface
between alternating Rrp41 and Rrp42 pairs. This resulted
in stable Rrp41:Rrp42 dimers that do not assemble into
hexamers
anymore
(Supplementary
Figure
S3).
The
Rrp41:Rrp42 dimers exhibit catalytic efficiencies that are
only slightly lower than the (Rrp41:Rrp42)3 particle for
RNAs >13 nt, and almost identical to the corresponding
hexamers for RNAs <13 nt. As a result, the drop around
13 nt from a faster to a slower degradation is much less
pronounced in the Rrp41:Rrp42 dimer, further supporting
the idea that encapsulation in the neck is responsible for
higher degradation speeds. The relatively high activity of
the dimers is possibly also a result of the effective ‘tripli-
cation’ of active sites, i.e. only one RNA molecule can be
degraded by a (Rrp41:Rrp42)3, while three RNA mol-
ecules can be degraded by three Rrp41:Rrp42 dimers. In
addition, while RNA probably dissociates faster from the
dimers, this effect could be compensated by a faster asso-
ciation of RNA to the readily accessible active sites in the
open dimers.
The opposite is observed, when the ring structure is
crosslinked. We introduced cysteines on the outside of
the
RNase-PH
ring
and
crosslinked
the
three
Rrp41:Rrp42 dimers via a thiol specific bifunctional
crosslinker. This procedure resulted in a hexameric
RNase PH ring with wild-type-like size and shape accord-
ing
to
gel
filtration
(Supplementary
Figure
S3).
Comparison of ni between the crosslinked isoform with
the (Rrp41:Rrp42)3 hexamer, revealed a dramatically
reduced ni (500- to 2000-fold) indicating that rigidifying
the exosome by the crosslink severely affects catalytic ef-
ficiency. We cannot formally rule out that the crosslinking
affects activity by other means, but considering that the
hexamer disrupting mutation at the same interface does
not severely reduce activity, a plausible scenario is that the
rigidified exosome does not allow efficient association with
RNA anymore.
Thus, taken together, the self-compartmentalization of
exosomes is probably not an evolution for high activity,
but rather for controlled RNA degradation.
Effect of the cap structures
To learn about the role of the cap proteins in exosome
activity, we compared the rate constants of the cap-less
exosome with the Csl4 and the Rrp4 exosome. The
initial rates for degradation of the 30-mer rA are similar
for the Csl4-exosome and the cap-less version, but consid-
erably faster for the Rrp4-exosome (Figure 5A). This in-
dicates that cap proteins can influence recognition and
recruitment of RNA substrates and that this step is
more efficient for the Rrp4 exosome. However, while
RNA degradation for medium and short RNAs is quite
comparable between the Csl4 and Rrp4 exosomes, there is
an interesting difference for long RNA molecules (>24 nt).
The Rrp4 exosome is quite slower for RNAs >24 nt, faster
for RNA between 24 and 13 nt, and then progressively
slower for RNAs <13 nt. This remarkable length depen-
dency is clearly evident in degradation profiles (Figure 1).
The most likely explanation is that long RNAs might still
have contacts with Rrp4, where a more specific binding
site holds them partially back from rapid degradation. In
principle, this could be viewed as molecular friction. When
RNAs are shorter, they loose contact to Rrp4 and degrad-
ation speed is increased. The Csl4 protein and the Rrp4
protein differ in their domain structures. While Csl4
contains
a
Zn-ribbon
domain,
Rrp4
possesses
a
KH-domain, which is a typical RNA-binding domain
and could recognize the oligo-rA. Such a binding could
be responsible for the faster first degradation step, because
it more efficiently sequesters RNAs on the exosome
surface, but may subsequently slow down degradation
until RNAs are too short to maintain simultaneous
contacts at the KH domain and active site.
However, the Rrp4 isoform is more efficient for smaller
RNA species than the Csl4 and capless isoforms. Since
these shorter RNAs cannot form dual contacts with the
active site and outside the caps, the Rrp4 could also influ-
ence the dynamics or other properties of the RNase-PH
ring, for instance to help in loading of RNA into the ring
structure.
SAXS structure of the Rrp4 exosome with endogenously
purified bacterial RNA
To explore the role of the Rrp4 cap in efficiently recruiting
RNAs further, we performed SAXS studies with a
nuclease deficient nine-subunit Rrp4 exosome bound to
RNA:
we
had
noticed
that
this
nuclease
deficient
Rrp4-exosome
(D180A
in
Rrp41)
very
efficiently
co-purifies with E. coli RNA. To determine the kind of
RNA that binds to the exosome we run it on a denaturing
gel together with RNAs with known sizes and could
estimate the size of the RNA to be between 55 and 65 nt
(Supplementary Figure S1). Cloning and sequencing of
bound RNA molecules revealed a set of much shorter in-
homogeneous mixed sequences (Supplementary Table S3).
It is possible that the bound RNAs are a mixture of
various mRNAs from E. coli, although the isolated
RNA is larger than the identified sequences and it is
possible that highly structured RNAs such as tRNAs are
underrepresented due to inefficient amplification and
cloning.
Comparison
of
the
SAXS
structure
of
apo-Rrp4-exosome with the RNA bound complex shows
an increase in the radius of gyration from 39.6 A˚ to 46.8 A˚
when RNA is bound and the corresponding pair distribu-
tion functions contains longer vectors (Figure 6A), likely
because
additional
scattering
elements
from
RNA
protrude from the compact protein core. The resulting
ab initio model of the complex overlaid with the crystal
structure of the apo-complex clearly indicates additional
mass from the bound RNA (Figure 6B and C). This clear
additional mass is distributed in the centre of the cap
structure on top of the neck region but also protrudes
Nucleic Acids Research, 2010, Vol. 38, No. 15
5173
away from the complex. When looking at the overlay with
the crystal structure it appears that the RNA is bound at
the KH and the S1-domains. The SAXS analysis supports
the model that RNA binds near the KH-domain on the
outside of the caps and reveals a low-resolution image of
trapped exosome–RNA complexes.
DISCUSSION
RNA exosomes are large, self-compartmentalized nucle-
ases, implicated in processive, controlled degradation of a
large variety of RNAs. While the archaeal exosome
possesses three phosphorolytic active sites within the com-
partment, the eukaryotic exosomes apparently have lost
this activity but adopted hydrolytic RNase subunits that
are bound at the outside of the evolutionary conserved
core. Nevertheless, recent data suggest that RNA is still
threaded through the eukaryotic core exosome before it is
degraded in ectopic hydrolytic active sites, suggesting that
the core particle retained critical ‘structural’ functions re-
garding RNA degradation such as increased processivity
or controlled RNA degradation (25).
To be able to quantitatively address RNA exosome
activities, we derived a kinetic model for the complex
RNA degradation of the archaeal RNA exosome using
Markov Chain Monte Carlo analysis. The kinetic model
gives a realistic assessment of the velocity of the exosome
and mutant variants during processive degradation of a
rA 30-mer oligonucleotide. The considerable effort we had
to put into the MCMC simulation pays offeventually. We
are now able to derive a realistic joint posterior distribu-
tion of kinetic parameters, enabling us to quantify the
relation of different parameters in either the same or in
distinct exosome mutants. This would have been impos-
sible with a conventional least squares fit of the data,
which produces very unstable parameter estimates (see
Supplementary Data for a comparison), although the
obtained fits are very good (Figure 3D).
With this in hand, we find several interesting and unex-
pected features of RNA degradation activities. First,
kinetic evaluation of RNA degradation of exosomes
needs to include association and dissociation rate con-
stants. Thus the kinetics cannot be treated as strictly
processive, at least for RNA species in the assessed
Figure 6. SAXS structure of the Rrp4 exosome with endogenously purified bacterial RNA. (A) SAXS data of the Rrp4 exosome (green) and the
Rrp4 exosome with RNA (orange) (curves show the scattering intensity I(q) as a function of the scattering angle 2y and X-ray wavelength , where
q = (4sin/y)) and the pair-distribution function describing intramolecular distances; in the presence of RNA longer distances occur and the radius of
gyration increases. (B) Average of 10 independent ab initio models for the apo exosome and the RNA-bound complex superimposed with the crystal
structure. The additional density for the RNA is clearly visible.
5174
Nucleic Acids Research, 2010, Vol. 38, No. 15
length range. This does not necessarily imply that RNA
dissociates and rebinds completely from the exosome.
Longer RNAs may be retained within the neck as well
as cap domains, while binding and dissociating from the
active site in the processing chamber. The association and
dissociation
constants
can
thus
be
understood
as
‘ratcheting’ constants that influence the rate of translation
along the RNA to and from the active site. For short
substrates that are unable to simultaneously bind neck
and active site this connection is lost, the dissociation in-
creases, degradation speeds drop and the exosome changes
from being fast and processive to a slower distributive
enzyme.
Our results also show how neck region and active site
features contribute to exosome activity. Although we
could
not
quantitatively
address
the
importance
of
Arg65 in the neck with the simplified model in hand,
this residue appears to be important for loading RNA
into the processing chamber, but not for efficient degrad-
ation once RNA is bound. This conclusion is derived from
the observation that while the initial degradation is sub-
stantially delayed, the appearance of smaller RNA species
is qualitatively similar to the wild-type Csl4 exosome.
Taken together with the observation that crosslinking
severely reduces processing and the RNase PH ring
needs
to
breath
or
display
some
conformational
dynamics, it is unlikely that RNA is simply threaded
into the processing chamber like a yarn through the eye
of a needle. Rather, we propose that initial RNA binding
includes some lateral entry near the neck.
We are also in the position now to address the influence
of the cap proteins Rrp4 and Csl4. These proteins possess
a variety of domains with unclear function in exosome
activity.
While
eukaryotic
exosomes
have
defined
heterotrimeric caps, the stoichiometry of cap proteins in
archaeal exosomes is not defined in vitro and perhaps
variable in vivo. For the archaeal exosome, the Csl4
capped isoform displays similar degradation kinetics to
that of the capless variant, and the function of this type
of cap remains to be shown. However, the Rrp4 isoform
substantially differs from the other two variants and our
analysis suggests that Rrp4 more efficiently recruits RNA
to the exosome. In fact, RNA from the heterologous
expression in E. coli is very tightly bound to the Rrp4
exosome. It must be noted that the gene coding for
Rrp4 is in the same operon as genes for Rrp41 and
Rrp42, indicating that this cap is perhaps a ‘default’
isoform of the exosome, while the Csl4 cap, located else-
where in the genome, might be differentially regulated.
The cap structures, however, also influence the degrad-
ation of short RNAs. This is to some extent surprising,
since short RNAs (<13 nt) cannot bind to the caps and the
active site at the same time. However, the Rrp4 subunit
more intimately interacts with the RNase PH ring than the
Csl4 protein and might influence also the dynamics of
the RNase PH type ring. Likewise, binding of RNA to
the KH domains, consistent with the lateral density of
RNA in the SAXS models, may position it better for
loading into the processing chamber.
In sum, we present here a robust method to analyse the
complex degradation kinetics of a partially processive
degradation enzyme in a quantitative manner, with esti-
mates of the posterior distribution of the model param-
eters. We applied this analysis to degradation of RNA by
several isoforms and variants of the archaeal exosome and
could reveal a variety of features that are important for
catalytic efficiency. The objective of this manuscript is to
derive a general method that can now be used to unravel
the biochemistry of exosomes in a more quantitative
manner. The method can now form a basis for compre-
hensive analysis of different substrates, other RNA se-
quences, as well as mutants of this system or related
systems such as the eukaryotic exosome.
ACCESSION NUMBERS
3M7N, 3M85.
SUPPLEMENTARY DATA
Supplementary Data are available at NAR Online.
ACKNOWLEDGEMENTS
The authors thank Christian Luginsland for help in
protein purification, Katharina Bu¨ ttner for exosome con-
structs and Katja Lammens and Gregor Witte for helpful
discussions. The authors thank the staffof the European
Synchrotron Radiation Facility (beamline 14–2) and the
Swiss Light Source (beamline PX I) for help with diffrac-
tion data collection and Michal Hammel from the
Advanced Light Source (SIBYLS beamline) for help
with scattering data collection.
FUNDING
Deutsche
Forschungsgemeinschaft
(HO2489/3
and
SFB646);
Center
for
Integrated
Protein
Science
Munich. Funding for open access charge: Deutsche
Forschungsgemeinschaft.
Conflict of interest statement. None declared.
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Nucleic Acids Research, 2010, Vol. 38, No. 15
|
3M7W
|
Crystal Structure of Type I 3-Dehydroquinate Dehydratase (aroD) from Salmonella typhimurium LT2 in Covalent Complex with Dehydroquinate
|
Insights into the Mechanism of Type I Dehydroquinate
Dehydratases from Structures of Reaction Intermediates*
Received for publication,October 8, 2010, and in revised form, October 28, 2010 Published, JBC Papers in Press,November 18, 2010, DOI 10.1074/jbc.M110.192831
Samuel H. Light‡§, George Minasov‡§, Ludmilla Shuvalova‡§, Mark-Eugene Duban‡, Michael Caffrey¶,
Wayne F. Anderson‡§, and Arnon Lavie¶1
From the ‡Center for Structural Genomics of Infectious Diseases and §Department of Molecular Pharmacology and Biological
Chemistry, Feinberg School of Medicine, Northwestern University, Chicago, Illinois 60611 and the ¶Department of Biochemistry
and Molecular Genetics, University of Illinois, Chicago, Illinois 60607
The biosynthetic shikimate pathway consists of seven en-
zymes that catalyze sequential reactions to generate choris-
mate, a critical branch point in the synthesis of the aromatic
amino acids. The third enzyme in the pathway, dehydro-
quinate dehydratase (DHQD), catalyzes the dehydration of
3-dehydroquinate to 3-dehydroshikimate. We present three
crystal structures of the type I DHQD from the intestinal
pathogens Clostridium difficile and Salmonella enterica.
Structures of the enzyme with substrate and covalent pre- and
post-dehydration reaction intermediates provide snapshots of
successive steps along the type I DHQD-catalyzed reaction
coordinate. These structures reveal that the position of the
substrate within the active site does not appreciably change
upon Schiff base formation. The intermediate state structures
reveal a reaction state-dependent behavior of His-143 in which
the residue adopts a conformation proximal to the site of cata-
lytic dehydration only when the leaving group is present. We
speculate that His-143 is likely to assume differing catalytic
roles in each of its observed conformations. One conformation
of His-143 positions the residue for the formation/hydrolysis
of the covalent Schiff base intermediates, whereas the other
conformation positions the residue for a role in the catalytic
dehydration event. The fact that the shikimate pathway is ab-
sent from humans makes the enzymes of the pathway potential
targets for the development of non-toxic antimicrobials. The
structures and mechanistic insight presented here may inform
the design of type I DHQD enzyme inhibitors.
Present in bacteria, fungi, and plants but absent in higher
eukaryotes, the seven enzymes of the shikimate pathway cata-
lyze sequential reactions to generate chorismate. Chorismate
serves as a precursor of many biologically important aromatic
compounds including the ubiquinones, folates, and aromatic
amino acids (1). The essential nature of these proteins in a
number of species, in combination with an absence of human
homologs, makes the shikimate pathway an attractive target
for the development of non-toxic antimicrobials, anti-fungals,
and herbicides (2–4). Given the complexities inherent to in-
hibitor design, a comprehensive understanding of the struc-
tural and mechanistic framework underlying the function of
the shikimate pathway enzymes should aid in the develop-
ment of novel shikimate-targeting inhibitors.
The third step in the shikimate pathway, consisting of the
dehydration of dehydroquinate to dehydroshikimate (Fig. 1),
can be catalyzed by two unrelated enzymes, termed type I and
type II dehydroquinate dehydratases (DHQDs).2 These two
enzyme families lack sequence or structural homology and
employ distinct reaction mechanisms (5–10). The type I
DHQDs utilize a covalent Schiff base (imine) intermediate
that results in a cis-elimination, whereas the type II reaction
lacks a covalent intermediate and undergoes a trans-elimina-
tion (5–10). The phylogenetic distribution of the two enzyme
types is somewhat disorderly, with closely related species of-
ten possessing different types. In general, the type I enzyme is
found in plants and fungi as a domain within a multifunc-
tional protein and in some bacteria as an 29-kDa mono-
functional protein that assembles into a homo-dimer. In con-
trast to the type I DHQD, the type II enzyme is found within a
mostly non-overlapping subset of bacteria and exists as an
17-kDa protein that assembles into a homo-dodecamer (8,
10–12).
Previously reported structures of the Salmonella typhi,
Staphylococcus aureus, and Archaeoglobus fulgidus type I
DHQD have characterized the enzyme in an apo and co-
valently bound post-dehydration intermediate state (8, 13–
15). Here we present crystal structures of the type I DHQD
from the two intestinal pathogens, the Gram-positive Clos-
tridium difficile (cdDHQD) and Gram-negative Salmonella
enterica (seDHQD), and characterize previously unobserved
substrate and pre-dehydration reaction intermediate bound
states of the enzyme. The similar mode of substrate and reac-
tion intermediate binding and the reaction state-dependent
behavior of a conserved active site histidine are identified, and
their functional implications are discussed.
EXPERIMENTAL PROCEDURES
Cloning, Protein Overexpression, and Purification—Follow-
ing published protocols (16), seDHQD and cdDHQD genes
* This work was supported, in whole or in part, by National Institutes of
Health Contract HHSN272200700058C (to W. F. A.) through the NIAID.
The atomic coordinates and structure factors (codes 3M7W, 3NNT, and 3JS3)
have been deposited in the Protein Data Bank, Research Collaboratory for
Structural Bioinformatics, Rutgers University, New Brunswick, NJ
(http://www.rcsb.org/).
1 To whom correspondence should be addressed: 900 South Ashland Ave.,
MBRB Room 1108, Chicago, IL 60607. Fax: 312-355-4535; E-mail
Lavie@uic.edu.
2 The abbreviations used are: DHQD, dehydroquinate dehydratase; cd-
DHQD, C. difficile DHQD; seDHQD, S. enterica DHQD.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 286, NO. 5, pp. 3531–3539, February 4, 2011
© 2011 by The American Society for Biochemistry and Molecular Biology, Inc.
Printed in the U.S.A.
FEBRUARY 4, 2011•VOLUME 286•NUMBER 5
JOURNAL OF BIOLOGICAL CHEMISTRY 3531
from C. difficile strain 630 and S. enterica subspecies enterica
serovar typhimurium strain LT2 genomic DNA were PCR-
amplified and cloned into pMCSG7 and pMCSG19 vectors,
respectively, each of which contains N-terminal hexahistidine
tag followed by a tobacco etch virus protease cleavage se-
quence. Insert-containing plasmids were transformed into
Escherichia coli BL21 (DE3) strain from Agilent (Santa Clara,
CA). Protein expression and purification were performed us-
ing standard Center for Structural Genomics of Infectious
Diseases protocols (17, 18). Crystallization screens were set
up immediately following purification, and the remaining en-
zyme was aliquoted, frozen in liquid nitrogen, and stored at
80 °C for future use.
DHQD Assays—Immediately before performing assays,
protein was thawed and diluted to the appropriate concentra-
tion. Assays were performed at 37 °C in potassium phosphate
buffer (100 mM, pH 7.5). Reaction was initiated by the addi-
tion of protein to a mixture of buffer and 3-dehydroquinic
acid (Sigma-Aldrich). Formation of the conjugated enone
carboxylate in dehydroshikimate was followed by mea-
suring the increase in absorbance at 234 nm relative to
substrate ( 12 mM1cm1) (4, 19). To determine the
kinetic parameters, triplicate measurements of the reaction
rate were determined at varying concentrations of 3-dehy-
droquinic acid. Data were fitted to the Michaelis-Menten
equation using the enzyme kinetics module in SigmaPlot
version 8.02.
Protein Crystallization and Data Collection—Protein con-
centrated to 7.5 mg/ml in a buffer containing 0.5 M NaCl and
10 mM Tris-HCl (pH 8.3) was used to set up sitting drops at a
ratio of 1:1 protein to reservoir. Further detail regarding crys-
tallization conditions is presented in Table 1. The substrate-
bound structure was obtained by co-crystallization of the Lys-
170 3 Met (K170M) mutant protein with 2 mM
dehydroquinic acid. The pre-dehydration reaction intermedi-
ate bound structure was obtained by soaking a crystal in
mother liquor containing 5 mM dehydroquinic acid for 15 min
prior to freezing in liquid nitrogen. The post-dehydration re-
action intermediate bound structure was obtained by co-crys-
tallization with 1 mM dehydroshikimic acid. All crystals where
immersed in mother liquor before being frozen in liquid ni-
trogen. Diffraction data were collected at 100° K at the Life
Sciences Collaborative Access Team at the Advanced Photon
Source, Argonne, IL.
Structure Determination and Refinement—Data were pro-
cessed using HKL-3000 for indexing, integration, and scaling
(20). Structures were solved with Phaser (21) using the
S. typhi apo DHQD structure (Protein Data Bank (PDB) code
1GQN) as a starting model for the cdDHQD structure and the
apo seDHQD structure (PDB code 3L2I) for determination of
seDHQD structures. Structures were refined with Refmac
(22) and manually corrected based on electron density maps
displayed in Coot (23). All figures were prepared in the
PyMOL Molecular Graphics System, Version 1.3 (Schro¨-
dinger, LLC). Atomic coordinates and structure factors were
deposited in the PDB under codes 3M7W (pre-dehydration
complex), 3NNT (K170M substrate complex), and 3JS3 (post-
dehydration complex).
RESULTS AND DISCUSSION
C. difficile and S. enterica DHQDs Display a Similar Overall
Structure—The cdDHQD and seDHQD share 56% sequence
identity (Fig. 2A). Structures of the reaction intermediate
bound seDHQD and cdDHQD protein were solved by molec-
ular replacement and refined to a resolution of 1.95 and 2.20
Å, respectively. The DHQD monomer from each bacterium is
characterized by an eight-stranded -barrel motif and is ob-
served to dimerize with helices 6, 7, 8, and 9, making anti-
parallel intermolecular contacts, in a manner similar to previ-
ously reported DHQDs (8, 13–15). The overlay of cdDHQD
and seDHQD shows a high degree of structural conservation
(Fig. 2B).
Differing Side Chain Conformation of His-143 in Pre- and
Post-dehydration States—To gain insight into the reaction
mechanism of DHQD, we obtained structures of DHQD com-
plexed with ligand. A soak of seDHQD crystals with the sub-
strate, 3-dehydroquinic acid, produced a structure in which
the substrate covalently linked at its 3-position to the Schiff
base-forming Lys-170 was observed (Fig. 3, A and B). The
bound ligand displays clear electron density corresponding to
the 1-hydroxyl leaving group (Fig. 3A, arrow), and therefore
this structure represents a pre-dehydration intermediate state
of the reaction. Co-crystallization of the cdDHQD with the
product, 3-dehydroshikimic acid, also produced a crystal
structure in which a Lys-1703-bound covalent species is ob-
served (Fig. 3, C and D). In this case, the bound ligand lacks
electron density corresponding to the leaving group. Interest-
ingly, an ordered water molecule is located directly above
where the leaving group was observed in the pre-dehydration
ligand (Fig. 3C, arrow) and is thus positioned to be the mole-
cule dehydrated from the substrate. This structure, which is
similar to previously reported type I DHQD intermediate
complexes (8, 14), represents a post-dehydration intermediate
state of the reaction. Together, these structures provide snap-
shots of successive steps in the reaction and thus allow for the
identification of structural changes that occur over the course
of the reaction.
A particularly striking reaction state-dependent structural
behavior is revealed by an overlay of the pre- and post-dehy-
dration reaction intermediate bound active sites. In the pre-
3 cdDHQD and seDHQD have different chain lengths, but for the sake of
consistency, all residue numbering within the text refers to the position
of the residue in the seDHQD protein.
FIGURE 1. The substrate and product of the dehydration reaction cata-
lyzed by DHQDs. Indicated is the numbering convention used throughout
the text for the substrate, intermediate, and product and the pro-R and
pro-S hydrogen relevant to the mechanism of elimination of the 1-hydroxyl
group.
Multiple Roles for Active Site Histidine in Type I DHQDs
3532
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 286•NUMBER 5•FEBRUARY 4, 2011
dehydration complex, His-143 is proximal to both elements of
the catalytic dehydration. The His-143 N2 atom is 2.7 Å from
the modeled pro-R C-2 hydrogen of the substrate and forms a
2.8 Å hydrogen bond with the 1-hydroxyl leaving group (Fig.
4A). In the post-dehydration covalent complex, the histidine
side chain is displaced 1.5 Å away from the site of catalytic
dehydration to the position it adopts in all previously reported
apo and post-dehydration reaction intermediate DHQD
structures (8, 13–15). In this position, the His-143 N1 atom
forms a 2.9 Å hydrogen bond with the conserved residue
Glu-86 (Fig. 4B).
To address the possibility that the displacement of His-143
observed in the pre-dehydration reaction intermediate bound
structure is a non-biologically relevant artifact of the low pH
of 4.6 of the crystallization condition, we used a similar crystal
soaking protocol to test a crystal grown at neutral pH. A com-
parable behavior of His-143 was observed in this neutral pH
crystal structure (data not shown), suggesting that the confor-
mation of His-143 seen in the pre-dehydration structure is
representative of the behavior of the residue over a wide range
of physiological pH values.
Having ruled out pH in explaining the displacement of
His-143, the only difference between the pre- and post-
dehydration reaction intermediate bound structures is the
presence of the leaving group in the pre-dehydration li-
gand. As such, the differential behavior of His-143 between
the two structures can be attributed to the presence of the
leaving group in the pre-dehydration reaction state. In par-
ticular, induction of the pre-dehydration conformation of
His-143 is likely due to the potential for formation of a fa-
vorable hydrogen-bonding interaction between the N2
atom of the histidine and the leaving hydroxyl group (Fig.
4). In the apo and post-dehydration reaction states, with-
out the potential for His-143 to hydrogen-bond with the
leaving group, adoption of the His-143 post-dehydration
conformation is likely due to the potential for the N1 atom
of the histidine to hydrogen-bond with Glu-86 in this posi-
tion. These structures define a never before characterized
leaving group-dependent conformation of His-143 in
which the residue is proximal to the site of dehydration
only when the leaving group is present.
Intermediate State Crystal Structures Suggest Role for His-
143 in Catalysis—Several lines of experimental evidence
suggest that this conserved active site histidine may act to
shuttle the proton from the C-2 position in the ring to the
1-hydroxyl leaving group. Hanson and Rose (9) used ste-
reo-specific labeling experiments to demonstrate that type
I dehydroquinate dehydration results in cis-elimination.
That is, the abstracted proton comes from the same side of
the ring as the leaving group. This finding is consistent
with a single residue both abstracting the proton from the
ring and protonating the leaving group (9). Based on di-
ethyl polycarbonate treatment resulting in an inactivation
of the protein, Deka et al. (24) hypothesized that the con-
served histidine might be the proton-transferring residue.
Mutagenesis studies might be expected to conclusively ad-
dress the role of this residue in proton shuttling. Although
Leech et al. (25) showed that the E. coli His-143 3 Ala
(H143A) mutant had a profound loss of activity, potentially
supportive of this histidine acting as the proton-shuttling
entity, it was unclear whether or not the effect of the muta-
tion was due solely to the role of this residue in Schiff base
formation and hydrolysis. Based on the body of kinetic evi-
dence, the proximity of the pre-dehydration conformation
TABLE 1
Crystallization conditions, data collection, and refinement statistics
Values for highest resolution shell are in parentheses.
Species
S. enterica
S. enterica
C. difficile
Variant
Wild type
K170M
Wild type
PDB code
3M7W
3NNT
3JS3
Color in figures
Cyan
Gray
Pink
Active site ligand
Pre-dehydration covalent intermediate
3-Dehydroquinate
Post-dehydration covalent intermediate
Crystallization conditions
170 mM NH4OAc, pH 4.6, 25.5% (w/v)
PEG 4000, 15% (v/v) glycerol
50 mM K2PO4, pH 6, 20% (w/v) PEG 8000
100 mM Tris, pH 8.5, 30% (w/v) PEG 500
Space group
C2
P1
P21
Unit cell dimensions
a, b, c (Å)
a 184.33, b 66.58, c 128.23
a 36.73, b 43.55, c 79.94
a 60.47, b 139.62, c 66.77
, , (°)
a 90.00, 119.08, 90.00
91.18, 101.27, 109.05
90.00, 90.63, 90.00
Resolution (Å)
29.69–1.95 (2.00–1.95)
29.65–1.60 (1.64–1.60)
29.55–2.20 (2.26–2.20)
Number of reflections
99109 (7189)
55405 (3978)
55907 (4028)
Completeness (%)
99.9% (99.1)
96.8 (95.2)
99.8% (98.0)
Redundancy
3.7 (3.6)
2.0 (2.3)
3.8 (3.8)
Rsym I
0.086 (0.466)
0.030 (0.357)
0.066 (0.614)
I/(I)
14.9 (2.8)
23.2 (2.3)
17.3 (2.2)
Molecules per aua
6
2
4
No. of atoms
Protein
11646
3992
8116
Water
1128
525
322
Ligand
156
26
44
Rwork/Rfree
0.154/0.206
0.158/0.187
0.189/0.241
r.m.s.b deviations
Bond lengths (Å)
0.012
0.011
0.012
Bond angles (°)
1.45
1.41
1.54
Average B-factors
Protein
25.4
25.5
26.9
Waters
35.9
37.3
28.2
Ligand
21.3
25.6
37.4
a asu, asymmetric unit.
b r.m.s., root mean square.
Multiple Roles for Active Site Histidine in Type I DHQDs
FEBRUARY 4, 2011•VOLUME 286•NUMBER 5
JOURNAL OF BIOLOGICAL CHEMISTRY 3533
of His-143 to both the pro-R C-2 proton and the 1-hy-
droxyl leaving group, and the absence of another feasible
proton-transferring residue, we propose a reaction mecha-
nism consistent with previously hypothesized models (25–
28) but in which His-143 catalyzes the dehydration event
by its N2 atom abstracting the pro-R C-2 proton and do-
nating it to the 1-hydroxyl leaving group (Fig. 5A).
Reaction State-dependent His-143 Behavior Suggests Con-
formation-dependent Catalytic Roles for the Residue—In addi-
tion to its role in the catalytic dehydration event, His-143 has
a well established role in the formation and hydrolysis of the
Schiff base intermediates. Leech et al. (25) found that the
H143A mutant had a profoundly deficient reaction rate. They
observed that the mutant enzyme slowly accumulated co-
FIGURE 2. Sequence and structure similarity between the cdDHQD and seDHQD type I DHQDs. A, sequence alignment of cdDHQD and seDHQD. Align-
ment was done in ClustalW2 version 2 using the default settings. Secondary features were defined using the cdDHQD structure in ESPript version 2.2. B,
superposition of the biological dimer of the seDHQD pre-dehydration complex (cyan) and the cdDHQD post-dehydration complex (pink) structures (back-
bone root mean square deviation 1.16 Å). Lys-170 and the reaction intermediates to which it is covalently bound are shown as sticks. Helices of the dimer
interface are labeled.
Multiple Roles for Active Site Histidine in Type I DHQDs
3534
JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 286•NUMBER 5•FEBRUARY 4, 2011
valently bound reaction intermediate at the active site and
interpreted this result to mean that the mutant enzyme has a
diminished capacity both to form and later to hydrolyze the
covalent Schiff base. Based on these findings, Leech et al. (25)
proposed a model in which His-143 acts to facilitate the pro-
tonation or deprotonation of the carbinolamine reaction in-
termediate depending upon whether the Schiff base is being
formed or hydrolyzed (Fig. 5B).
Considering that adoption of the His-143 pre-dehydration
conformation appears to be leaving group-dependent, the
hydrolysis of the Schiff base, which occurs post-dehydration,
must initiate with His-143, adopting its post-dehydration
conformation. As such, His-143 would appear to assume dif-
fering functionalities in each of its conformations. In its pre-
dehydration conformation, the residue catalyzes the catalytic
dehydration event (Fig. 5A), whereas in its post-dehydration
conformation, the residue catalyzes Schiff base hydrolysis
(Fig. 5B). Although the conformation His-143 adopts when it
catalyzes Schiff base formation cannot be inferred from these
structures, clearly, His-143 plays a complex and highly tuned
role in which it shifts between two conformations to catalyze
multiple aspects of the type I DHQD reaction coordinate.
Similar Positioning of Substrate and Reaction Intermediates
within the Active Site Supports a Catalytic Role for the Schiff
Base—Stereoelectronic principles dictate that the Lys-170
amine nucleophile will form the covalent Schiff base after ap-
proaching the carbonyl carbon of the substrate at the Bu¨rgi-
Dunitz (N–C–O) angle of 107° (29, 30). According to this
model of bond formation, the position of the Lys-170 amine
relative to the 3-carbon of the substrate should differ mark-
edly during its approach to the carbonyl when compared with
after the Schiff base has formed. To meet these positional re-
quirements, either Lys-170 or the substrate must undergo
substantial movement as the substrate transitions from a non-
covalent to Schiff base bound state. The simplest mechanism
by which the enzyme might accommodate both the Bu¨rgi-
Dunitz approach and the corresponding active site rearrange-
ments would be for the substrate to initially dock in an orien-
tation that positioned it for the nucleophilic approach.
Subsequently, in a process correlated with bond formation,
the substrate could move to its covalent intermediate bound
position.
In fact, this expectation is supported by studies of several
enzymes that generate a similar Schiff base intermediate by
the nucleophilic attack of a lysine on a substrate ketone. Crys-
tallographic studies of dihydrodipicolinate synthase (31), fruc-
tose 1,6-bisphosphate (32), and 2-keto-3-deoxy-6-phospho-
gluconate aldolase (PDB ID 3LAB) all show that the
orientation of the covalent Schiff base intermediate is rotated
180° within the active site when compared with the non-
covalently bound substrate. These observations are consistent
with the substrate initially docking in a conformation that
FIGURE 3. Crystal structures of DHQD in pre- and post-dehydration covalent intermediate states. A, structure of seDHQD active site with covalent pre-
dehydration reaction intermediate. Carbon atoms are depicted in cyan, oxygens are in red, nitrogens are in blue, and sulfurs are in yellow. Difference maps
were calculated with the reaction intermediate omitted from the model. The Fo Fc map is contoured at the 3.0 level (red), and the 2Fo Fc map is con-
toured at the 1 level (blue). An arrow indicates the 1-hydroxyl leaving group. B, schematic rendering of the pre-dehydration reaction intermediate shown
in A. Distances between atoms are shown in angstroms. C, structure of the cdDHQD active site with covalent post-dehydration reaction intermediate.
Model and maps are the same as A, except that carbons are shown in pink. The arrow points toward an ordered water molecule at a position consistent with
that dehydrated from the substrate. D, schematic rendering of the post-dehydration reaction intermediate shown in C. Distances between atoms are shown
in angstroms.
Multiple Roles for Active Site Histidine in Type I DHQDs
FEBRUARY 4, 2011•VOLUME 286•NUMBER 5
JOURNAL OF BIOLOGICAL CHEMISTRY 3535
allows for a Bu¨rgi-Dunitz approach before moving to the in-
termediate bound position in a process concurrent with bond
formation. Whether this mode of substrate nucleophile ap-
proach is specific to these enzymes or is general to the Schiff
base-forming enzymes is presently unknown.
Based on the severe effect of the Lys-170 3 Ala mutation
on kcat but minimal effect on Km, it has been argued that the
Schiff base is likely to play a direct catalytic role in the reac-
tion mechanism of DHQD (25). Suggestions on the mecha-
nism by which the Schiff base might promote catalysis have
focused on how it could distort the ring of the substrate in a
manner that would stereoelectronically promote elimination
or function to stabilize the carbanion reaction intermediate
(25, 33, 34). However, in light of the stereoelectronic argu-
mentation and experimental data describing substrate bind-
ing in other Schiff base-forming enzymes, in DHQD, it is also
likely that prior to Schiff base formation, the substrate adopts
an orientation that is different from the covalent reaction in-
termediate within the active site. In that case, it is conceivable
that Schiff base formation is required to position the substrate
in the proper orientation in which for catalysis to occur, pre-
senting a scenario wherein the Schiff base plays only an indi-
rect role in catalysis.
To gain insight into the initial substrate binding event
within Schiff base-forming enzymes generally and to deter-
mine whether the Schiff base is directly involved in DHQD
catalysis, we set out to capture a complex of DHQD with sub-
strate. We reasoned that if the non-covalently bound sub-
strate was similarly positioned relative to the covalently
bound reaction intermediate, then formation of the Schiff
base must not result in significant change in the position of
the substrate within the active site and therefore cannot be
promoting catalysis by positioning the substrate into the cata-
lytically required orientation. In that case, having ruled out
the proposed non-catalytic (orientation-determining) role of
the Schiff base, it could be concluded that the Schiff base is
likely to have only a direct involvement in catalysis.
To obtain a crystal structure of seDHQD in a non-covalent
complex with substrate, site-directed mutagenesis was uti-
lized to convert the reactive Schiff base-forming Lys-170 3
Met (K170M). Methionine was chosen because it is the resi-
due closest in shape to a lysine but cannot form a Schiff base.
As previously reported (25), mutation of Lys-170 resulted in a
dramatic loss in reactivity (Table 2). Co-crystallization of the
K170M mutant enzyme with dehydroquinic acid produced a
structure in which the substrate was observed non-covalently
bound at the active site (Fig. 6A). A comparison of the pre-
dehydration intermediate bound structure and K170M sub-
strate-bound structure shows the ring core of the substrate to
be similarly positioned but slightly shifted (0.3 Å) within the
active site, likely to avoid a clash with the Met-170 terminal
methyl group (Fig. 6B). The similar orientation of substrate
and reaction intermediate demonstrates a differential mode of
substrate binding when compared with other Schiff base-
forming enzymes. The dispensability of Lys-170 in substrate
binding, as reasoned above, provides support for the Schiff
base playing a direct catalytic role in the reaction mechanism
of the protein.
K170M Substrate-bound Structure Provides Insight but Also
Generates New Questions Regarding Schiff Base Formation—
Although a previous study has shown a role for His-143 in
Schiff base formation (25), the mechanism of His-143 involve-
ment remains ill-defined. Based on mutagenesis studies,
Leech et al. (25) proposed that His-143 catalyzes the conver-
sion of the carbinolamine intermediate to the Schiff base in-
termediate (Fig. 5B). However, it is possible that His-143 af-
fects other steps in Schiff base formation. The K170M
substrate-bound structure reveals that His-143 adopts partial
occupancies of both of the conformations observed in the
wild type structures (Fig. 6A). Both positions of His-143 are
proximal to the 3-carbonyl oxygen of the substrate (Fig. 6C).
This structural observation positions the residue to protonate
the carbonyl oxygen and/or to accentuate the dipole of the
carbonyl by drawing charge away from the carbonyl carbon,
which might promote the Lys-170 nucleophilic attack. The
proximity of His-143 to the carbonyl oxygen prior to the for-
mation of the covalent intermediate suggests that the residue
may play a functional role in an earlier step of Schiff base for-
mation than has previously been recognized.
FIGURE 4. The conformation of His-143 is dependent on the presence of
the 1-hydroxyl group of the reaction intermediate. A, in the pre-dehy-
dration reaction intermediate structure (cyan), the N2 atom of His-143 is
within interaction distance to the 1-hydroxyl (2.8 Å) and the pro-R C-2 hy-
drogen (2.7 Å). Thus, it can facilitate dehydration by transferring the hydro-
gen atom to the leaving hydroxyl group. B, superposition of the pre- (cyan)
and post- (pink) dehydration states of DHQD. In the pre-dehydration state,
His-143 interacts with the 1-hydroxyl group. Upon leaving of this group, as
revealed by the post-dehydration intermediate structure, His-143 rotates to
a position where it interacts with Glu-86 (2.9 Å).
Multiple Roles for Active Site Histidine in Type I DHQDs
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JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 286•NUMBER 5•FEBRUARY 4, 2011
Although providing new insight into the significance of the
Schiff base in catalysis, the K170M substrate-bound structure
presents new questions regarding how formation of the Schiff
base occurs. The observed position of the substrate is incon-
sistent with the Lys-170 N atom approaching the carbonyl
carbon of the substrate at an ideal stereoelectronic angle.
Modeling Lys-170 into the K170M substrate-bound structure
reveals that, without significant perturbations to the main
chain, the maximal approach angle that the Lys-170 amine
can achieve falls well short of the 107° Bu¨rgi-Dunitz angle
(Fig. 6D). As such, the bond must form either by a non-Bu¨rgi-
Dunitz approach of the substrate in its observed position or
by a Bu¨rgi-Dunitz approach of the substrate in an unobserved
position within the active site.
Possible exceptions to the Bu¨rgi-Dunitz nucleophilic ap-
proach in the context of enzyme catalysis are not without
precedent (35). In conjunction with previous observations,
the mode of substrate binding observed here potentially sup-
ports a more widespread exception to the established mode of
nucleophilic approach in enzyme catalysis. Alternatively, the
approach of Lys-170 and formation of the covalent bond may
occur when the substrate transiently adopts a conformation
that places its carbonyl carbon at the Bu¨rgi-Dunitz angle; this
requirement may be met during the process of substrate
docking into the active site. In that case, in the wild type en-
zyme, the covalent adduct may form before the substrate
reaches its observed position in the K170M substrate-bound
structure. The mechanism of Schiff base formation is the
topic of ongoing study.
Conclusions—We report three crystal structures that ad-
dress several issues about how type I DHQDs function. The
substrate and pre-dehydration covalent intermediate bound
structures provide the first view of these reaction states. The
different conformation of His-143 in pre- and post-dehydra-
tion intermediate states defines a leaving group-dependent
behavior of the residue. The proximity of His-143 to the C-2
proton and leaving group of the pre-dehydration reaction in-
termediate supports a role for this residue in the transport of
the proton to the leaving group in the catalytic elimination
and provides a rare example of the requirement of the leaving
group for a residue to adopt the conformation consistent with
its presumed catalytic role. Previous kinetic studies and the
structural data presented here suggest a reaction mechanism
in which His-143 moves between two conformations while
undergoing a series of protonation/deprotonation events to
FIGURE 5. Proposed role of His-143 in type I DHQD-catalyzed reaction. A, putative role of His-143 in the catalytic dehydration. Following forma-
tion of the covalent Schiff base linker between Lys-170 and the substrate 3-dehydroquinate 1, His-143 assumes its pre-dehydration position, where
its N2 atom forms a key hydrogen-bonding interaction with the 1-hydroxyl group of the reaction intermediate. In this position, the His-143 N2 atom
abstracts the C-2 pro-R proton of the substrate to generate the carbanion intermediate 2, which gives rise to the enamine intermediate 3. The proto-
nated His-143 then delivers its N2 proton to facilitate departure of the 1-hydroxyl leaving group, which generates the ene-iminium intermediate 4.
Because the H143 N2 atom can no longer form the critical interaction with the 1-hydroxyl leaving group, a shift to the post-dehydration position of
the residue ensues. Finally, following Schiff base hydrolysis, the formally dehydrated product 3-dehydroshikimate is released. Boxed intermediates 1
and 4 represent likely states of the reaction captured by pre- and post-dehydration crystal structures. B, role of His-143 in Schiff base formation and
hydrolysis based on Leech et al. (25). In the formation of the Schiff base, attack by the Lys-170 N atom on the 3-carbonyl carbon leads to formation
of the carbinolamine intermediate. His-143 then delivers a proton to facilitate departure of the hydroxyl leaving group to generate the Schiff base
intermediate. Following the catalytic hydrolysis described in A, His-143 adopts its post-dehydration conformation and catalyzes the reverse reaction
to hydrolyze the Schiff base and regenerate the active site.
TABLE 2
Kinetic characterization of seDHQD and cdDHQD
Errors were calculated by fitting of the kinetic data to the Michaelis-Menten
equation and expressed as S.D.
kcat
Km
kcat/Km
s1
M
s1M1
seDHQD wild type
210 5
21 3
10 1
seDHQD K170M
0.015 0.007
33 4
4.5 104 2.3 104
cdDHQD wild type
125 4
36 9
3.5 0.9
Multiple Roles for Active Site Histidine in Type I DHQDs
FEBRUARY 4, 2011•VOLUME 286•NUMBER 5
JOURNAL OF BIOLOGICAL CHEMISTRY 3537
catalyze multiple steps in the formation and hydrolysis of the
Schiff base as well as the catalytic dehydration.
The K170M substrate-bound structure provides insight
into the role of the Schiff base in catalysis. The similar mode
of binding of substrate in the K170M variant and reaction
intermediate in the wild type enzyme eliminates the possibil-
ity that the functional role of the Schiff base is to orient the
substrate within the active site. Based on these results, we
conclude that the Schiff base must have a direct role in ca-
talysis. It is anticipated that the more detailed knowledge
of the DHQD kinetic pathway provided by this work will
aid in the design of novel anti-bacterials, which are partic-
ularly relevant for the emerging pathogens C. difficile and
S. enterica.
Acknowledgments—We thank Dr. Elisabetta Sabini for facilitating
the communications with the groups involved in this work and Dr.
Scott Peterson and Dr. Keehwan Kwon for providing DHQD expres-
sion clones. Use of the Advanced Photon Source at Argonne Na-
tional Laboratory was supported by the U. S. Department of Energy,
Office of Science, Office of Basic Energy Sciences, under Contract
DE-AC02-06CH11357. Use of the Life Sciences Collaborative Access
Team (LS-CAT) Sector 21 was supported in part by the Michigan
Economic Development Corporation and the Michigan Technology
Tri-Corridor for the support of this research program (Grant
085P1000817).
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FEBRUARY 4, 2011•VOLUME 286•NUMBER 5
JOURNAL OF BIOLOGICAL CHEMISTRY 3539
|
3M81
|
Crystal structure of Acetyl xylan esterase (TM0077) from THERMOTOGA MARITIMA at 2.50 A resolution (native apo structure)
|
Functional and structural characterization of a thermostable
acetyl esterase from Thermotoga maritima
Mark Levisson1,*, Gye Won Han2,3,*, Marc C. Deller2,3, Qingping Xu2,4, Peter Biely5, Sjon
Hendriks1, Lynn F. Ten Eyck6,7, Claus Flensburg8, Pietro Roversi8, Mitchell D. Miller2,4,
Daniel McMullan9, Frank von Delft2,3,‡, Andreas Kreusch10, Ashley M. Deacon2,4, John van
der Oost1, Scott A. Lesley2,3,10, Marc-André Elsliger2,3, Servé W. M. Kengen1,†, and Ian A.
Wilson2,3,†
1Laboratory of Microbiology, Wageningen University, 6703 HB, Wageningen, The Netherlands
2Joint Center for Structural Genomics, http://www.jcsg.org 3Department of Molecular Biology, The
Scripps Research Institute, La Jolla, California 92037 4Stanford Synchrotron Radiation
Lightsource, SLAC National Accelerator Laboratory, Stanford University, Menlo Park, California
92045 5Institute of Chemistry, Slovak Academy of Sciences, 845 38 Bratislava, Slovakia
6Department of Chemistry and Biochemistry, University of California at San Diego, La Jolla,
California 92093-0505 7San Diego Supercomputer Center, University of California at San Diego,
La Jolla, California 92093-0505 8Global Phasing Ltd. Sheraton House, Castle Park, Cambridge
CB3 0AX, United Kingdom 9Protein Therapeutics Department, Genomics Institute of the Novartis
Research Foundation, San Diego, California 92121 10Protein Sciences Department, Genomics
Institute of the Novartis Research Foundation, San Diego, California 92121
Abstract
TM0077 from Thermotoga maritima is a member of the carbohydrate esterase family 7 and is
active on a variety of acetylated compounds, including cephalosporin C. TM0077 esterase activity
is confined to short-chain acyl esters (C2-C3), and is optimal around 100°C and pH 7.5. The
positional specificity of TM0077 was investigated using 4-nitrophenyl-β-D-xylopyranoside
monoacetates as substrates in a β-xylosidase-coupled assay. TM0077 hydrolyzes acetate at
positions 2, 3 and 4 with equal efficiency. No activity was detected on xylan or acetylated xylan,
which implies that TM0077 is an acetyl esterase and not an acetyl xylan esterase as currently
annotated. Selenomethionine-substituted and native structures of TM0077 were determined at 2.1
Å and 2.5 Å resolution, respectively, revealing a classic α/β-hydrolase fold. TM0077 assembles
into a doughnut-shaped hexamer with small tunnels on either side leading to an inner cavity,
which contains the six catalytic centers. Structures of TM0077 with covalently bound
phenylmethylsulfonyl fluoride (PMSF) and paraoxon were determined to 2.4 Å and 2.1 Å,
respectively, and confirmed that both inhibitors bind covalently to the catalytic serine (Ser188).
Upon binding of inhibitor, the catalytic serine adopts an altered conformation, as observed in other
esterase and lipases, and supports a previously proposed catalytic mechanism in which this Ser
hydroxyl rotation prevents reversal of the reaction and allows access of a water molecule for
completion of the reaction.
†Correspondence to: Ian A. Wilson, Ph.D., Department of Molecular Biology, The Scripps Research Institute, La Jolla, California
92037; (858) 784-2939 Fax: (858) 784-2980; wilson@scripps.edu or Servé W. M. Kengen, Ph.D., Laboratory of Microbiology,
Wageningen University, 6703 HB, Wageningen, The Netherlands; 31 317 483737, Fax: 31 317-483829; serve.kengen@wur.nl.
*ML and GWH contributed equally to this work.
‡Current address: The Structural Genomics Consortium, Roosevelt Drive, Headington, Oxford OX3 7DQ, UK
NIH Public Access
Author Manuscript
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Published in final edited form as:
Proteins. 2012 June ; 80(6): 1545–1559. doi:10.1002/prot.24041.
NIH-PA Author Manuscript
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Keywords
Acetyl esterase; Thermotoga maritima; crystal structure; α/β hydrolase; inhibitor; serine rotation
INTRODUCTION
Thermotoga maritima is a hyperthermophilic bacterium that grows optimally at 80°C and is
able to metabolize a variety of simple and complex carbohydrates, including glucose,
sucrose, starch, cellulose, and xylan 1. Its carbohydrate utilization potential was confirmed
by analysis of its sequenced genome 2. The xylan degrading pathway of T. maritima has
been studied using microarrays 2–4, and several genes encoding transporters, xylanases, and
a β-xylosidase have been identified. Among the enzymes with a differential expression
pattern in the microarray was a predicted acetyl xylan esterase (locus tag TM0077,
axeA) 3,5. Depending on the source, the xylan backbone may contain a varying degree of
acetylated xylose residues. Therefore, in addition to xylanases and xylosidases, the complete
degradation of xylan requires esterases/deacetylases 6.
Presently, esterases and deacetylases that are active on carbohydrate substrates have been
classified into 16 families by Henrissat and coworkers (Carbohydrate-Active enZymes
Server (CAZy)) 7. According to this classification, the predicted acetyl xylan esterase from
T. maritima would be a member of family 7 of the carbohydrate esterases (CE7). In addition
to the acetyl xylan esterase activity, enzymes in the CE7 family are rather unusual in that
they display a high specific activity towards the antibiotic cephalosporin C [(Fig. 1(a-b)] 8.
Cephalosporins belong to the β-lactam class of antibiotics, which also includes penicillin,
and affect bacterial cell growth by inhibiting the penicillin-binding-protein that cross-links
peptide glycans required for cell wall formation 9. The production of deacetylated
cephalosporins is of great interest because these compounds are valuable building blocks for
the production of semi-synthetic β-lactam antibiotics10,11.
To explore the catalytic capacity of the predicted acetyl xylan esterase from T. maritima and
gain a better insight into the structure and function of the family 7 carbohydrate esterases,
TM0077 was expressed and purified, and three-dimensional structures of the native enzyme
and its complexes with phenylmethylsulfonyl fluoride (PMSF) and paraoxon inhibitors,
were determined by x-ray crystallography. Furthermore, the enzyme was functionally
characterized, and various biochemical properties including the positional specificity of the
esterase were investigated.
MATERIALS AND METHODS
Gene cloning
TM0077 was selected as part of the Joint Center for Structural Genomics (JCSG) effort on
complete structural coverage of the T. maritima soluble proteome as a large-scale center for
high-throughput structure determination funded under the NIHGMS Protein Structure
Initiative (PSI) 12. The gene encoding TM0077 (GenBank: AAD35171.1, GI:4980565;
SwissProt: Q9WXT2) was amplified by polymerase chain reaction (PCR) from genomic
DNA using PfuTurbo DNA polymerase (Stratagene) and primers corresponding to the
predicted 5′ and 3′ ends. The PCR product was cloned into plasmid pMH1, which encodes
an expression and purification tag (MGSDKIHHHHHH) at the amino terminus of the
protein. The cloning junctions were confirmed by DNA sequencing.
Levisson et al.
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TM0077-SeMet protein production and purification
Protein production was performed in a selenomethionine-containing medium using the
Escherichia coli methionine auxotrophic strain DL41. Expression was induced by the
addition of 0.15% L-arabinose. At the end of fermentation, cells were harvested and
subjected to one freeze/thaw cycle, and subsequently sonicated in Lysis Buffer [50 mM Tris
pH 7.9, 50 mM NaCl, 1 mM MgCl2, 0.25 mM Tris(2-carboxyethyl)phosphine hydrochloride
(TCEP), 1 mg/ml lysozyme] and the lysate was centrifuged at 3,400 × g for one hour. The
soluble fraction was applied to nickel-chelating resin (GE Healthcare) pre-equilibrated with
Equilibration Buffer [50 mM potassium phosphate pH 7.8, 300 mM NaCl, 10% (v/v)
glycerol, 0.25 mM TCEP] containing 20 mM imidazole. The resin was washed with
Equilibration Buffer containing 40 mM imidazole, and the protein was eluted with Elution
Buffer [20 mM Tris pH 7.9, 300 mM imidazole, 10% (v/v) glycerol, 0.25 mM TCEP]. The
eluate was buffer exchanged with Buffer Q [20 mM Tris pH 7.9, 5% (v/v) glycerol, 0.25
mM TCEP] containing 50 mM NaCl and applied to a RESOURCE Q column (GE
Healthcare) pre-equilibrated with the same buffer. The protein was eluted using a linear
gradient of 50 to 500 mM NaCl in Buffer Q and purified further with a HiLoad 16/60
Superdex 200 column (GE Healthcare), using Crystallization Buffer [20 mM Tris pH 7.9,
150 mM NaCl, 0.25 mM TCEP] as the mobile phase. For crystallization trials, the peak
Superdex 200 fractions were concentrated to ~15 mg/mL by centrifugal ultrafiltration
(Millipore). Molecular weight and oligomeric state of TM0077 were determined using a 1
cm × 30 cm Superdex 200 column (GE Healthcare) coupled with miniDAWN static light
scattering (SEC/SLS) and Optilab differential refractive index detectors (Wyatt
Technology). The mobile phase consisted of 20 mM Tris pH 8.0, 150 mM NaCl, and 0.02%
(w/v) sodium azide.
Native TM0077 production and purification
For protein production, E. coli DL41 cells were grown in LB medium for 8 hours (an
OD600 well above 2.0 was reached). Subsequently, the culture was induced by adding
0.15% L-arabinose and incubated another 16 hours at 37°C. Cells were harvested by
centrifugation at 10,000 × g for 20 min. The cell pellet was resuspended in 30 ml of Lysis
Buffer 2 [50 mM Tris-HCl pH 8.0, 50 mM NaCl, 10 mM imidazole, 0.25 mM TCEP]. The
cells were disrupted by two passages through a French press at 110 MPa. The crude cell
extract was treated with DNAse I at room temperature for 30 min and subsequently
centrifuged at 43,000 × g for 30 min in order to remove cell debris. The supernatant was
heated at 70°C for 25 min and then centrifuged to remove the precipitated proteins. The
supernatant was filtered and loaded onto a nickel-chelating column packed with 20 ml of Ni-
NTA His-Bind Resin (Novagen) and equilibrated in 50 mM Tris-HCl pH 8.0, 300 mM
NaCl, 2% (v/v) glycerol, and 0.25 mM TCEP. The column was washed with 20 mM
imidazole in the same buffer, and proteins were subsequently eluted with a linear gradient of
20–500 mM imidazole in the same buffer. Fractions containing esterase activity were pooled
and loaded onto a HiPrep Desalting column (GE Healthcare) equilibrated with 20 mM Tris-
HCl pH 8.0, 150 mM NaCl, and 0.25 mM TCEP. The homogeneity of the protein was
checked by SDS-PAGE, and activity staining of the SDS-PAGE gel was performed using α-
napthyl acetate, as described previously 13. The protein concentration was determined at 280
nm using a NanoDrop ND-1000 Spectrophotometer.
Crystallization
Crystals of selenomethionine-substituted TM0077 were obtained by hanging drop vapor
diffusion against a 250 μl crystallization solution consisting of 20% (w/v) PEG-3000, 0.1 M
HEPES pH 7.5, 0.2 M NaCl. Drops consisted of 0.5 μl protein and 0.5 μl crystallization
solution. Native TM0077 was crystallized using nanodrop vapor diffusion techniques
against a crystallization solution consisting of 0.2 M calcium acetate hydrate, 20% (w/v)
Levisson et al.
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PEG 3350, pH 7.3 at 20°C. Protein was concentrated to 22.8 mg/ml. Drops consisted of 100
nl protein and 100 nl of crystallization solution and a 60 μl reservoir of crystallization
solution. Crystals of TM0077 in complex with inhibitors PMSF or paraoxon were obtained
at 4°C in the same conditions with the same reagents as the native crystals. PMSF or
paraoxon were added in a molar ratio of 1:3 (protein:inhibitor).
Data collection
For cryoprotection, the TM0077-SeMet crystal was transferred to crystallization solution
supplemented with 15% (v/v) glycerol. The crystal was mounted in a cryoloop and
subsequently flash-cooled in liquid nitrogen. X-ray data were collected at 100 K on
beamline BL9-2 at the Stanford Synchrotron Radiation Lightsource (SSRL, Menlo Park,
CA) using a Quantum 4 CCD detector (ADSC). A TM0077-SeMet MAD data set was
collected to 2.1 Å resolution and the data were indexed in monoclinic space group P21, with
unit cell parameters a = 152.6 Å, b = 131.0 Å, and c = 157.8 Å, and β=118.9°, and 12
molecules in the asymmetric unit. Data were indexed and integrated with DENZO and then
scaled with SCALEPACK 14.
Native TM0077, TM0077-PMSF complex (TM0077-PMS) and TM0077-paraoxon complex
(TM0077-DEP) crystals were transferred to crystallization solution supplemented with 10%
(v/v) ethylene glycol and flash-cooled to 100K. Data were collected at beamline 5.0.3 of the
Advanced Light Source (ALS, Berkeley, CA) and processed with the HKL2000 package 14.
The native data set was collected to 2.5 Å resolution, and TM0077-PMS and TM0077-DEP
data sets were collected to 2.4 and 2.1 Å, respectively. All data were indexed in
orthorhombic space group P212121, with unit cell parameters approximately a=103Å
b=104Å c=221Å (See Table 1), and six molecules in the asymmetric unit. Data reduction
and refinement statistics for TM0077-SeMet, TM0077-Native, TM0077-PMS and TM0077-
DEP are summarized in Table I.
Structure solution and refinement
The TM0077-SeMet structure was solved by MAD phasing method using a two-wavelength
MAD dataset. At the time of the initial data collection (2001), the structure determination of
Se-MAD TM0077 posed a significant challenge to crystallographic programs, which were
still under active development. As a result, modifications were made in various structure
determination and refinement programs to achieve success. For initial phasing, SHELXD 15
was used to find candidate SeMet substructure sites. Attempts to complete phasing were
unsuccessful due to the translational non-crystallographic symmetry (NCS) (not recognized
initially). Self-consistent sets (partial sets) were found using the CCP4 program
PROFESSS 16 and additional SeMet sites were found by SHELXD, and added to these
partial sets. The SHARP 17 run did not complete initially; however, updates of SHARP and
ARP/wARP eventually helped to resolve issues and an initial trace was obtained by ARP/
wARP. The structure was then refined with BUSTER 18 using tight NCS restraints to an
Rcryst and Rfree of 18.6% and 22.3%, respectively. Model building was performed using
O 19 and the structure was refined using Refmac5 20. Refinement statistics are summarized
in Table I. The final model contains 12 protein molecules (chains A-L) in the asymmetric
unit each consisting of residues 2-323. The MolProbity 21 Ramachandran plot analysis
showed that 97.4% of all residues are in favored regions with a single outlier, Gln120 of
chain B, which is supported by unambiguous electron density. Ramachandran outlier
Gln120 of chain B of TM0077-SeMet is due to crystal packing with chain C. The backbone
carbonyl oxygens of Gln120 and Gly119 of chain B makes hydrogen bonds with the
backbone nitrogen of Gln140 of chain C (3.19 and 3.11 Å, respectively).
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The native TM0077 structure, TM0077-PMS and TM0077-DEP structures were solved by
molecular replacement using PHASER 22,23 with the TM0077-SeMet hexamer coordinates
(pdb: 1vlq; A-F chains) as a search model. One hexamer was successfully located and the
structure was further refined with Refmac5 20 using tight NCS restraints to an Rcryst and
Rfree of 16.7% and 21.2% (native TM0077), 16.0% and 20.8% (TM0077-PMS) and 16.7%
and 20.5% (TM0077-DEP), respectively. Iterative cycles of refinement and building were
performed with Refmac5, Phenix 24,25 and Coot 26. All other crystallographic manipulations
were carried out with the CCP4 package 16. Refinement statistics are summarized in Table I.
The final model of native TM0077 contains residues 3-324 (chains A, B, C, D and F) and
3-323 (chain E) in the asymmetric unit. Analysis of main-chain torsion angles using
MolProbity 21 showed that 97.8% of the residues are in favored regions of the
Ramachandran plot with 0.2% outliers (Asn302 of chains B, C and D), which are supported
by unambiguous electron density. The final model of TM0077-PMS contains residues 3-323
for all chains in the asymmetric unit with 97.5% of the residues in favored regions with
0.2% outliers (Asn302 of chains A, B, D and F). The final model of TM0077-DEP contains
residues 0-324 for (chains A, B, C and F) and 0-323 (chains D and E) in the asymmetric
unit, respectively, with 97.6% of the residues in favored region of the Ramachandran plot
with 0.2% outliers (Asn302 of B, C, D and F chains). Ramachandran outlier Asn302 in the
TM0077-Native, TM0077-PMS and TM0077-DEP structures is a neighbor to the catalytic
triad residue His303 and may reflect a slightly different state for these structures compared
to the Se-Met structure.
Structure validation and deposition
The quality of the crystal structure was analyzed using the JCSG Quality Control server
(http://smb.slac.stanford.edu/jcsg/QC). This server processes the coordinates and data
through a variety of validation tools including AutoDepInputTool 27 MolProbity 21,
WHATIF 5.0 28, RESOLVE 29, MOLEMAN2 30 as well as several in-house scripts, and
summarizes the results. Protein quaternary structure analysis were performed using the PISA
server 30. Figures were prepared with PyMOL (DeLano Scientific) 31. RMSD values were
calculated using the ProCKSI-Server 32. The structural data have been deposited in the
RCSB Protein Data Bank (PDB) with accession codes 1vlq for TM0077-SeMet, 3m81 for
TM0077-native, 3m83 for TM0077-DEP and 3m82 for TM0077-PMS.
Enzyme assays
Esterase activity was measured using p-nitrophenyl esters as described previously 13.
Briefly, the standard assay consisted of activity measurements with 0.2 mM p-nitrophenyl
acetate as substrate in 50 mM citrate-phosphate (pH 6) at 70°C. The p-nitrophenol liberated
was measured continuously at 405 nm on a Hitachi U-2001 spectrophotometer with a
temperature-controlled cuvette holder. Extinction coefficients of p-nitrophenol were
determined prior to each measurement. Kinetic parameters were determined by direct fitting
the data, obtained from multiple measurements, to the Michaelis–Menten curve (Tablecurve
2d, version 5.0).
The effect of pH on esterase activity was studied in the pH range from 5 to 10. The buffers
used were 50 mM citrate-phosphate (pH 5–8) and 50 mM CAPS (3-(cyclohexylamino) 1-
propanesulphonic acid) (pH 9.5–10). The pH of the buffers was set at room temperature, and
temperature corrections were made using their temperature coefficients: −0.0028 pH/°C for
citrate-phosphate buffer and −0.018 pH/°C for CAPS buffer. The effect of temperature on
esterase activity was studied in the range of 40–100°C using 0.2 mM p-nitrophenyl acetate
as substrate. Enzyme thermostability was determined by incubating the enzyme in a 50 mM
Tris-HCl, 150 mM NaCl (pH 7.8) buffer at 90°C and 100°C for various time intervals.
Residual activity was determined in the standard assay.
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Inhibition kinetics of PMSF and paraoxon were determined as described for the
acetylcholinesterase from electric eel 33. All experiments were performed at 70°C in 50 mM
citrate-phosphate (pH 6) buffer and 0.2 mM p-nitrophenyl acetate as substrate. The kinetic
constants for the inhibition of TM0077 with PMSF and paraoxon were measured in the
concentration range of 1.0–10.0 mM and 0.2–1.0 mM, respectively.
Deacetylase activity was determined using high-performance liquid chromatography
(HPLC) by measuring the amount of acetic acid released from the substrates cephalosporin
C, 7-aminocephalosporanic acid, glucose-pentaacetate and acetylated xylan. Xylan was
acetylated by the method described by Johnson 34. The reaction mixture contained 0.9 ml of
substrate solution (dissolved in 50 mM Tris-HCl, pH 7.5) and 0.1 ml of enzyme solution,
and was incubated at 37°C for various time intervals (0–10 min). The reaction was stopped
by adding 0.2 ml of stop solution (100 mN H2SO4 and 30 mM crotonate) and placing the
sample on ice. The conditions for HPLC were as follows: column, KC811 Shodex;
detection, RI and UV detectors; solvent, 3 mN H2SO4; flow rate, 1.5 ml/min; temperature,
30°C; internal standard, crotonate. One unit of enzyme activity was defined as the amount of
enzyme that releases one μmol of acetic acid per minute.
Activity on xylan was measured quantitatively using DMSO-extracted xylan (1%
polysaccharide solution in 0.1 M sodium phosphate buffer pH 6) at 60°C 35. Xylan will
precipitate as a consequence of deacetylation, resulting in a rapid turbidity of the solution.
Positional specificity assay
The positional specificity of TM0077 was investigated using an enzyme-coupled assay on
monoacetylated 4-nitrophenyl β-D-xylopyranosides (pNP-Xyl) as described 36. The β-
xylosidase XloA (locus tag: TM0076) from T. maritima was cloned into the vector pET24d
in frame with a C-terminal His6-tag. The enzyme was expressed and purified as described
above for native TM0077. Activity of XloA was confirmed by measuring the release of p-
nitrophenol at 405 nm from the substrate 4-nitrophenyl β-D-xylopyranoside.
The enzyme-coupled assay was performed at 60°C in a total volume of 125 μl, which
contained 0.1 M sodium phosphate (pH 6 or 7), 2-O-, 3-O-, or 4-O-acetyl pNP-Xyl, the β-
xylosidase XloA, and TM0077. Stable 50x-concentrated stock solutions of the substrates
were prepared in DMSO. The reaction was started by addition of 2.5 μl of a stock solution
to a preheated reaction mixture consisting of phosphate buffer, auxiliary β-xylosidase XloA
in excess, and TM0077. The reaction was terminated by addition of 800 μl of a 2% solution
of Na2CO3. Liberated p-nitrophenol was determined at 405 nm against substrate and
enzyme blanks. A short incubation time for activity determination was used to suppress
acetyl migration on the xylopyranosyl-ring, which is significant at pH 6 or 7 37. The kinetic
constants were determined at pH 7 and 60°C with reaction times of 2 or 5 minutes.
RESULTS and DISCUSSION
In silico analysis
TM0077 consists of 325 amino acids with a calculated molecular mass of 37 kDa. Sequence
analysis, using the SignalP 3.0 server, revealed that TM0077 has no predicted signal
sequence and is, therefore, believed to be an intracellular enzyme. Analysis of the gene
organization indicates that the TM0077 gene co-localizes with genes encoding a xylanase
(TM0070) 38, ABC transporter components (TM0071-TM0075), and a β-xylosidase
(TM0076) 39. BLAST-P analysis showed that TM0077 has highest similarity to putative
acetyl esterases, acetyl xylan esterases and cephalosporin C deacetylases. Among the
BLAST results, a predicted acetyl xylan esterase-related protein from T. maritima (locus
tag: TM0435) was also identified. TM0077 was compared with other members of the CE7
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family using structure-based, multi-sequence alignment and the putative catalytic triad,
Ser188, Asp274, and His303, was identified from conservation throughout the analyzed
sequences. The putative nucleophilic serine (Ser188) is located within a conserved
pentapeptide consensus sequence, Gly-Xaa-Ser-Gln-Gly, typical of this family. Previously, a
signature sequence motif, [RGQ]-(x:~70)-[GxSQG]-(x:~115)-[HE] (where x indicates any
amino acid), had been suggested for the CE7 family based on an aminoacid alignment of 12
sequences 40. In an updated alignment consisting now of 50 sequences, we observed many
sequences that have this signature motif, but it is not conserved throughout the entire family
(See Supporting Information and Fig. S1 for the multi-sequence alignment).
Overall structure
The crystal structure of seleno-methionine incorporated TM0077 (TM0077-SeMet) was
determined to 2.1 Å resolution by multi-wavelength anomalous dispersion (MAD) (Table I)
with twelve molecules per asymmetric unit. A native apo structure (TM0077-Native) was
determined in a different space group (see Methods) to 2.5 Å by molecular replacement,
using TM0077-SeMet as a search model, with six molecules in the asymmetric unit (Table
I). Each monomer of the native hexamer contained a calcium ion (see below) bound by
Lys22, Glu26, and Asp25 via a bridging water molecule. Superposition of the TM0077-
SeMet and the TM0077-Native structures gave a root-mean-square difference (RMSD) of
0.12 Å over 321 Cα atoms, which indicates that these structures are nearly identical as
expected.
In general, the TM0077 structure resembles the canonical α/β-hydrolase fold, which
consists of a central, twisted, eight-stranded β-sheet surrounded by α-helices on both sides,
with β2 antiparallel to the other strands. TM0077 deviates slightly from the canonical α/β-
hydrolase fold at two locations: a three-helix insertion after strand β6 and an extension of
the N-terminus (Fig. 2). Insertions after β6 or β7 are common for α/β-hydrolases and are
proposed to help shape the substrate-binding site 41. The N-terminus is extended by two
helices (αA-1 and αA-2) and an antiparallel β-strand (β-1) that aligns with the other eight β-
strands (β1-β8) and extends the central β-sheet. This nine-stranded β-sheet is highly twisted,
and β-1 and β8 at the extreme edges are rotated approximately 130° relative to each other.
Helices αA and αB both contain a short 310-helix segment at their N-terminus. Helices
αA-1, αA-2, αB, αC, αD, αD1, αD2, αD3, αE, and the 310-helix η2 are located on one side
of the central β-sheet, and helices αA, αF and the 310-helix η1 are on the other side.
A structural similarity search was performed using the program DALI 42. Monomer A of the
TM0077-SeMet structure was used as a search model and similarity was found with
cephalosporin C deacetylases, acetyl xylan esterases, acylamino-releasing enzymes,
dipeptidyl peptidases and some esterases and lipases. TM0077 is structurally most similar to
cephalosporin C deacetylase (CAH) from B. subtilis (PDB: 1ods) 40, acetyl xylan esterase
(AXE) from B. pumilus (PDB: 3fvr and 2xlb), acetyl xylan esterase (AXE1) from
Thermoanaerobacterium sp. JW/SL YS485 (PDB: 3fcy), and acylpeptide hydrolase/esterase
apAPH from Aeropyrum pernix K1 (PDB: 1ve6) 43. The sequence identity between
TM0077 and CAH is 41% and the two structures align with a Z-score of 46 and an RMSD
of 1.5 Å over 312 Cα atoms. The sequence identity with apAPH is 17% with a Z-score of
23.3 and an RMSD of 2.3 Å over 230 Cα atoms. Superpositions of TM0077 with CAH and
with apAPH are shown in Fig. 3.
Quaternary structure
The crystal structure of TM0077-SeMet contains two hexamers in the asymmetric unit that
are related by a non-crystallographic two-fold axis. Each hexamer contains a dimer of
trimers with a back-to-back arrangement (Fig. 4). The apo and the complex crystals
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contained one hexamer in the asymmetric unit. Crystallographic packing analysis using
PISA (EBI) 44 indicated that the relevant physiological oligomeric state of TM0077 is a
hexamer, which was confirmed by size exclusion chromatography coupled with static light
scattering. Further analyses of the hexameric assembly indicated that two main interfaces
play an essential role in complex formation. The first interface between subunit A and B
(green and cyan in Fig. 4) (identical to C/D and E/F) is stabilized by seven hydrogen bonds
on average and has a buried surface area of 1024 Å2 contributed by each chain. The second
interface between A and F (green and purple in Fig. 4) (B/C and D/E) is stabilized by 17
hydrogen bonds on average with a buried surface area of 1079 Å2 contributed by each chain
However, a multiple sequence alignment of TM0077 with other CE7 esterases showed that
the residues involved in these two main interfaces are not conserved. Other secondary
interfaces bury around 514 Å2 contributed by each chain. The hexamer has a total buried
surface area of 18,860 Å2, which is approximately 30% of the total surface area.
Approximately 3,143 Å2 per monomer is, therefore, buried upon complex formation.
The TM0077 hexamer has a doughnut-shape when viewed from the side, with the six active
sites located in the interior of the complex, where they line an oval-shaped cavity [Fig. 4(a)].
This cavity is accessible via two entrances, one on each side of the flat hexamer. Each of
these entrances is approximately 13 Å wide and connects to a short tunnel or pore spanning
approximately 10 Å to reach the inner cavity. Interestingly, in the TM0077-SeMet hexamer,
the entrance to the internal cavity is blocked by three phenylalanine residues (Phe4), one for
each of three monomers that compose half of the hexamer [Fig. 4(b)]. Residue Phe4 is
located in the mobile N-terminus (high B-values), which may indicate some flexibility or
multiple conformations.
Calcium ions were identified, by the electron density and coordination geometry, supported
by their presence in the crystallization reagents, in the native TM0077, TM0077-PMS and
TM0077-DEP structures, but not in the TM0077-SeMet structure. The SeMet protein was
crystallized without any calcium in the crystallization reagents. In each subunit of the
hexamer, one calcium ion is located at the N-terminal region of helix αA-1, and is
coordinated by the backbone carbonyl of Lys22 and the Glu26 carboxylate. The remainder
of the calcium coordination sphere is filled with waters from a neighboring solvent channel
present in all molecules in the asymmetric unit. The Asp25 carboxylate contributes to the
calcium binding via one of the coordinating water molecules. Another calcium ion is bound
in a crystal packing interface between chain A and chain C′ of a crystallographic symmetry-
related hexamer. This calcium is coordinated by the carboxylates of GluA45 and AspA58
from one chain and the carboxylate from Glu C’45 (bidentate coordination) of the
symmetry-related chain with three water molecules completing a capped-octahedral
coordination sphere. An equivalent calcium binding site is also observed in the crystal
packing interface between chains D and B′. No significant increase or reduction of activity
of TM0077 was observed in the presence of calcium ions or EDTA. Therefore, it seems that
these calcium ions are not important for activity. On the other hand, calcium may help
stabilize the structure. No calcium was present in the B. subtilis CAH structure 40; however,
Lys22, Glu26 and Ser25 are conserved and may also act as a calcium binding site.
Enzyme activity
The activity of TM0077 was investigated using p-nitrophenol esters with varying acyl-chain
length, ranging from C2 to C18. TM0077 is only active on the short-chain p-nitrophenol
esters of acetate and propionate and does not hydrolyze esters with acyl chains longer than
four carbons. No significant difference was found in the catalytic efficiency (kcat/Km) for the
hydrolysis of p-nitrophenyl with acyl chains containing 2 to 3 carbons (Table II) [Fig. 1(c)].
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The effect of temperature on activity was studied using p-nitrophenyl acetate as substrate.
The esterase activity increased from 40°C upwards until 100°C [Fig. 5(a)]. An Arrhenius
analysis resulted in linear plots in the temperature ranges of 40–60°C and 60–100°C [Fig.
5(a); inset], with calculated activation energies for the formation of the enzyme substrate-
enzyme complex of 33.7 and 21.9 kJ/mol, respectively. The transition or break in linearity
of the Arrhenius plot at 60°C (1000/T (K) = 3.0) could indicate some conformational change
of the enzyme. TM0077 is fairly resistant to thermal inactivation. An approximate 50%
transient increase in activity is seen during the first 10 to 20 minutes when the enzyme is
incubated at 90°C. After 30 minutes, inactivation of function occurs by first order kinetics
with a half-life of approximately 120 minutes [Fig. 5(b)]. A transient activation has also
been observed for other thermophilic esterases, such from Sulfolobus shibatae 45, and it is
believed that a high temperature is needed in order to obtain the optimal conformation for
catalysis. TM0077 was not stable at 100°C, resulting in a half-life of less than 5 minutes.
However, the optimum temperature and thermal stability of TM0077 are still considerably
higher than those reported for other characterized CE7 esterases, including the
Thermoanaerobacterium enzyme that has a temperature optimum of 80°C and a half-life of
1h at 75°C 46.
The effect of pH on activity was measured in the pH range of 4.8 to 9.2 using the substrate
p-nitrophenyl acetate. TM0077 displayed maximum activity at approximately pH 7.5 [Fig.
5(c)], which is comparable to other CE7 esterases, such as the acetyl xylan esterases from
Thermoanaerobacterium sp. strain JW/SL-YS485 46.
Positional specificity
The positional specificity of TM0077 was tested on three monoacetates of 4-nitrophenyl β-
D-xylopyranoside (pNP-Xyl). To determine the enzyme activity, the β-xylosidase XloA 39
(TM0076) from T. maritima is required as an auxiliary enzyme. This thermostable XloA
enzyme was, therefore, cloned, heterologously expressed, purified to homogeneity, and its
activity was confirmed by measuring release of p-nitrophenol from the substrate pNP-Xyl
(data not shown). The β-xylosidase was not active on the three monoacetates of pNP-Xyl. In
the XloA-coupled assay, TM0077 hydrolyzed acetate from positions 2, 3 and 4 of pNP-Xyl
with similar catalytic efficiency. The results are summarized in Table II.
In addition, TM0077 was investigated for its ability to remove acetyl groups from 7-
aminocephalosporanic acid (7-ACA), cephalosporin C, glucose penta-acetate, N-acetyl-D-
glucosamine, xylan and acetylated xylan. TM0077 has no activity for acetylated and non-
acetylated xylan polymers, indicating that it is, indeed, an acetyl esterase and not an acetyl
xylan esterase. As expected for an acetyl esterase, TM0077 displayed high activity on
glucose penta-acetate with a turnover number of 2680 s−1. Like other members of CE7,
TM0077 was also able to hydrolyze the acetyl groups from both cephalosporin C and 7-
ACA with a turnover number of 376 s−1 and 1140 s−1, respectively. TM0077 was not able to
hydrolyze the acetyl group from N-acetyl-D-glucosamine, indicating that it is specific for
ester bonds and unable to hydrolyze amide bonds.
Inhibitor assays and TM0077 structures complexed with PMSF and paraoxon
PMSF and paraoxon [Fig. 1(d,e)] are competitive irreversible inhibitors of esterases.
Inhibition proceeds by the formation of a reversible Michaelis complex, followed by an
irreversible step and inhibition can, therefore, be characterized by two parameters: a
dissociation constant and a binding rate constant. The inhibition kinetics for paraoxon and
PMSF were investigated in the presence of p-nitrophenyl acetate, as described previously 47,
and the dissociation and rate constants were 0.5 ± 0.1 mM and 0.13 ± 0.02 s−1 for paraoxon,
and 1.1 ± 0.2 mM and 0.020 ± 0.001 s−1 for PMSF, respectively. The acetyl xylan esterase
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from Bacillus pumilus (BpAxe) has slightly reduced sensitivity to paraoxon (dissociation
and rate constants of respectively 5.4 mM and 0.012 s−1), likely due to steric hindrance of
two tyrosine residues (Tyr91 and Tyr206) that hamper the binding of paraoxon. Although
these residues are essentially conserved in TM0077 (Tyr92 and Phe213), TM0077 is more
sensitive to paraoxon than BpAxe48. In comparison to EST2 of Alicyclobacillus
acidocaldarius 49 and EstA of T. maritima 47, the TM0077 dissociation constant is slightly
higher, but the rate constant is comparable. No significant stimulation or reduction of
activity of TM0077 was observed in the presence of divalent metal ions or
ethylenediaminetetraacetic acid (EDTA).
To obtain more information about inhibitor binding and any possible conformational
changes during catalysis, TM0077 was co-crystallized with the inhibitors PMSF and
paraoxon and the PMSF (TM0077-PMS) and paraoxon (TM0077-DEP) structures were
determined to 2.4 Å and 2.1 Å, respectively (Table I). The electron density map of TM0077
with PMSF showed clear density for the PMSF covalent modification. The fluorine was
cleaved from the PMSF molecule during the binding reaction and the phenylmethyl sulfonyl
(PMS) moiety is covalently bound to the Oγ atom of Ser188. The native apo and PMS-
bound structures superimpose well with RMSD’s of 0.09–0.11 Å over 320–321 Cα atoms.
Electron density maps of the paraoxon-bound structure displayed clear density for a diethyl-
phosphate moiety covalently bound to the Oγ atom of Ser188. This covalent modification
indicates that the p-nitrophenol group of paraoxon was cleaved off during co-crystallization,
and a tetrahedral product reminiscent of the first transition state was formed during carboxyl
ester hydrolysis. The native apo and paraoxon-bound structures superimpose with RMSD’s
of 0.12–0.32 Å over 320–322 Cα atoms. Attempts to obtain co-crystals of TM0077 with
cephalosporin C, even at a low temperature of 4°C, were unsuccessful.
Analysis of the active site
TM0077 has a classic catalytic triad, consisting of Ser188 as the nucleophile, His303 as the
proton acceptor/donor, and Asp274 as the acidic residue stabilizing the histidine (Fig. 6).
The catalytic serine Ser188 is located within a conserved pentapeptide sequence, Gly-X-Ser-
X-Gly (GGSQG), characteristic of esterases and lipases. The positions of Ser188, Asp274,
and His303 are consistent with their expected locations in the canonical fold of the α/β-
hydrolase family. Ser188 is located at the nucleophile elbow in a sharp turn between β5 and
helix αC. The presence of three glycine residues (Gly186, Gly187, and Gly190) in close
proximity to Ser188 prevents steric hindrance and facilitates access to the nucleophile
elbow. Asp274 and His303 are located in loops between β7 and helix αE, and between β8
and helix αF, respectively. The oxyanion hole is formed by the backbone amide groups of
Tyr92 and Gln189. The catalytic triad and oxyanion hole are located in a depression on the
surface of TM0077. This ellipsoid pocket (S1), which is approximately 12 Å wide, extends
15 Å from the catalytic serine. A smaller pocket (S2), approximately 5 Å long, extends to
the other side of the catalytic serine [Fig. 6(a)]. The volume of both pockets combined (S1 +
S2) is 1082 Å3 (CASTp analysis; 50). The substrate-binding pocket is bordered by residues
from helices αA and αF, and its base is formed by residues from β-strands 4, 5, 6, and their
adjacent C-terminal loops. The overall pocket is hydrophobic, although it does have some
polar residues (Gln88, Asp210, and Gln314), which may interact with the substrate.
In the native apo structure, the Ser188 hydroxyl makes a hydrogen bond with the imidazole
of His303 [Fig. 6(b)]. Extra density was observed near the side chain of Ser188 and was
interpreted as a chloride ion based on electron density size and shape as well as the
geometry of the interactions with surrounding residues. This chloride ion is bound at the
entrance of the oxyanion hole, forming hydrogen bonds with the backbone amides of Tyr92
and Gln189. In the PMSF-bound structure, the phenyl ring of the inhibitor is located in the
small active site groove surrounded by hydrophobic residues Tyr92, Trp124, Pro228, Ile276,
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and His303 [Fig. 6(c)]. The sulfonyl group of PMSF makes hydrogen bonds with the
backbone amides of Tyr92 and Gln189. In the paraoxon-bound structure, the diethyl-
phosphate (DEP) moiety is stabilized by hydrogen-bonding interactions with the oxyanion
hole. One of the two ethyl arms of bound paraoxon points toward the larger pocket in the
protein, while the other follows the groove of the small pocket. The two ethyl arms are
stabilized by packing against Tyr92, Trp124, Pro228, Ile276, and His303 [Fig. 6(d)].
Two rotamers of the catalytic serine
Although no large conformational changes were observed upon binding of PMSF or
paraoxon, a different rotamer of the catalytic serine side chain was observed compared to
native TM0077 [Fig. 7(a,b)]. Similar changes have been observed in several other esterases
and have been shown to play a key role in the catalytic mechanism (see CONCLUSION for
more details). In the native structure, the catalytic Ser188 Oγ is in the plane of the imidazole
ring of His303, as most commonly observed in the resting state of esterases and lipases 51.
The Ser188 Oγ forms a hydrogen bond (2.6 Å) with His303 Nε2. In the PMSF- and
paraoxon-bound structures, the conformation of the catalytic serine changes; the Ser188 Oγ
rotates about 110°, increasing the distance (3.1 Å and 2.8 Å for PMSF and paraoxon bound
structures, respectively) to the His303 imidazole ring. In the TM0077-SeMet structure, the
catalytic serine is also rotated over ~110°, with a distance to the imidazole ring of 3.0 Å
[Fig. 7(c.)]. A probable explanation for this observation could be the protonation of His303,
since TM0077-SeMet was crystallized at a lower pH (pH 4.2) compared to the native
TM0077 (pH 7.3). Furthermore, extra electron density was identified in the TM0077-SeMet
structure, suggesting a partially occupied acyl intermediate on Ser188. However, as this
density is not sufficiently clear and interpretable to fit an acyl intermediate, water molecules
were modeled instead. No rearrangements of any other residues in the active site were
observed.
CONCLUSION
TM0077 from the hyperthermophilic bacterium T. maritima was predicted from its gene
sequence to be an acetyl xylan esterase. We have expressed and purified TM0077 and
experimentally demonstrated that it has ester-hydrolyzing activity. The TM0077 activity was
restricted to short acyl chain esters (C2 and C3) when artificial p-nitrophenyl-esters were
used as substrates. In addition, the enzyme has high specific activity on glucose penta-
acetate. However, no activity was detected on xylan or acetylated xylan. Thus, TM0077
should be reclassified as an acetyl esterase, and not as an acetyl xylan esterase as currently
annotated 52. Furthermore, the lack of any apparent signal sequence suggests that the protein
is not secreted. Thus, the predicted intracellular location of TM0077 is compatible with a
role other than the deacetylation of extracellular xylan. Based on these results, we conclude
that the likely biological function of TM0077 is removal of the remaining acetyl groups
from the short, end products of xylan degradation that are imported into the cytoplasm. The
resulting deacetylated xylose oligomers are the substrates for a β-xylosidase. This role for
TM0077 is in good agreement with the clustering of the TM0077 gene with other genes
involved in xylose metabolism. However, it cannot be ruled out that TM0077 may also act
on other small, acetylated compounds.
TM0077 is the first esterase from the CE7 family to be tested for its positional specificity for
the deacetylation of 4-nitrophenyl-β-D-xylopyranoside. TM0077 hydrolyzes acetate at the 2,
3 and 4 positions of 4-nitrophenyl-β-D-xylopyranoside with similar efficiency. Conversely,
the CtAxe esterase from Clostridium thermocellum in the CE4 family shows a clear
preference for hydrolyzing acetate at the 2 position 53, and Penicillium purpurogenum AXE
II esterase, a member of the CE5 family, also has a preference for acetate at position 2 54.
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This lack of preference for a specific position of the acetate group correlates with the
relative broad substrate specificity of the CE7 esterases.
Esterases and deacetylases in the CE7 family are unusual in that they are active towards both
acetylated xylo-oligosaccharides and the antibiotic cephalosporin C [Fig. 1(a,b)]. Therefore,
TM0077 was investigated for activity towards the substrates 7-ACA and cephalosporin C.
The activity of TM0077 on these substrates is approximately ten-fold higher than that of the
acetyl xylan esterase from B. pumilus 55 or the acetyl esterase from Thermoanaerobacterium
sp. strain JW/SL YS485 56. TM0077 has a higher hydrolytic activity on 7-ACA compared to
cephalosporin C, as described for other CE7 esterases 40,55,56. Nonetheless, it is unlikely that
both compounds are natural substrates, because the stability of these compounds at the
optimal growth temperature (80°C) of T. maritima is very low.
Crystal structures of TM0077 in complex with inhibitors PMSF and paraoxon revealed that,
upon binding of PMSF or paraoxon, the reaction is trapped at the acylation step via the
formation of a covalent tetrahedral reaction product. In the complexed structures, the
negatively charged oxygen of the tetrahedral intermediate, derived from the substrate
oxyanion, is stabilized by hydrogen bonds with the backbone amide groups of Tyr92 and
Gln189. Comparison of the TM0077 complexed structures with the native structure shows
that the catalytic serine (Ser188) Oγ rotates about 110°, thereby increasing the distance
between Ser188 Oγ and His303 Nε2. Such a conformational change of the catalytic serine
has been observed in several other esterases, including Fusarium solani cutinase 57,
Penicillium purpurogenum acetyl xylan esterase 51, Rhodococcus sp. strain MB1 cocaine
esterase 58, Bacillus subtilis lipase 59, Rhodococcus sp. strain H1 heroine esterase 60, and
Aspergillus niger feruloyl esterase 61. The classical model for the catalytic mechanism of
esterases consists of a sequential two-step hydrolysis. The first reaction involves
nucleophilic attack by the catalytic serine on the substrate carbonyl carbon, resulting in an
acyl-enzyme and the liberation of an alcohol. In the second reaction, a water molecule
performs a nucleophilic attack on the acyl-enzyme, the acyl-enzyme bond breaks and the
carboxylate is released 62. Although the catalytic mechanism of esterases is well established,
it is unclear why the initially generated tetrahedral intermediate does not collapse back to the
reactant complex during the nucleophilic attack of the substrate. A previously proposed
mechanism that would prevent this collapse is the spatial reorganization of the catalytic
residues during the initial catalytic step, causing the residues to separate and thereby drive
the reaction forward 62–64. The apo and inhibitor bound structures of TM0077, presented
herein, support this proposed mechanism. Moreover, in a recent study of the serine protease
mechanism, it was suggested that subtle atomic motions of the catalytic serine and histidine
residues during the catalytic cycle favor the forward reaction 65. Thus, rotation of Ser188 Oγ
of TM0077 may be required to inhibit reversal of the reaction. In addition, such changes
may facilitate the access of water to the catalytic histidine so that the second step of the
reaction can go to completion.
Deacetyl cephalosporins are valuable building blocks for the production of semisynthetic β-
lactam antibiotics. These compounds are derived from cephalosporin C or 7-
aminocephalosporanic acid via enzymatic or chemical processes 10. The thermostable
TM0077 esterase may be valuable in the preparation of derivatives of β-lactam antibiotics.
Recently, the substrate specificity of the acetyl xylan esterase from P. purpurogenum was
engineered to accept a range of fatty acid esters of up to 14 carbons compared to its wild-
type preference for acetate54. It might also be possible to engineer TM0077 and enable the
(de)acetylation of cephalosporins at the C10 position with various acyl chains. Because of
its high stability and activity on 7-ACA and cephalosporin C, TM0077 presents an attractive
candidate for the production of new semi-synthetic antibiotics.
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Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We gratefully acknowledge contributions from George Sheldrick for modifications of the SHELXD program, and
for Global Phasing Ltd. that made significant improvements in the automation of autoSHARP. We also thank
Victor Lamzin for updates of chain docking of the ARP/wARP program, and Gerard Bricogne and Eleanor Dodson
for helpful discussion on phasing for the large TM0077-SeMet structure, and Willem J. van Berkel for valuable
discussion on the catalytic mechanism of TM0077. Portions of this research were carried out at the Stanford
Synchrotron Radiation Lightsource (SSRL) and the Advanced Light Source (ALS). The SSRL is a Directorate of
SLAC National Accelerator Laboratory and an Office of Science User Facility operated for the U.S. Department of
Energy Office of Science by Stanford University. The SSRL Structural Molecular Biology Program is supported by
the DOE Office of Biological and Environmental Research, and by the National Institutes of Health, National
Center for Research Resources, Biomedical Technology Program (P41RR001209), and the National Institute of
General Medical Sciences. The ALS is supported by the Director, Office of Science, Office of Basic Energy
Sciences, Materials Sciences Division, of the U.S. Department of Energy under Contract No. DE-
AC02-05CH11231 at Lawrence Berkeley National Laboratory. Genomic DNA from Thermotoga maritima MSB8
(DSM3109) (ATCC #43589D-5) was obtained from the American Type Culture Collection (ATCC). The content is
solely the responsibility of the authors and does not necessarily represent the official views of the National Institute
of General Medical Sciences or the National Institutes of Health.
Grant sponsor: NIH Grant numbers U54 GM094586 and U54 GM074898 (Protein Structure Initiative); Grant
sponsor: The Graduate School VLAG Wageningen, the Netherlands (ML).
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Figure 1.
Substrates and inhibitors of the CE7 family of enzymes. Structures of (A) acetylated
xylooligosaccharide, (B) cephalosporin C, (C) p-nitrophenyl-acetate, (D)
phenylmethylsulfonyl fluoride (PMSF), and (E) paraoxon.
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Figure 2.
Overall fold and topology of TM0077. (A) Stereo view of a TM0077 protomer. The β-
strands are labeled numerically (-1 to 8) with the core strands in red, α-helices are labeled
alphabetically (A-2 to F) and 310-helices are labeled with an Eta (η1 and η2) with the core
helices in cyan. The three-helix insertion after β6 is colored green and the N-terminal
extension is colored sky blue. The figure was generated using Pymol 31. (B) Topology
diagram of TM0077, with the helices displayed as cylinders and the strands displayed as
arrows following the color and label scheme of (A). The location of residues forming the
catalytic triad is also indicated.
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Figure 3.
Structural superposition of TM0077 with structurally related esterases. Superposition of
TM0077 (yellow) with (A) the cephalosporin C deacetylase (CAH) from B. subtilis (PDB:
1ods; blue) 40 and (B) the α/β-hydrolase domain of the acylpeptide hydrolase/esterase
apAPH from A. pernix K1 (PDB: 1ve6; grey) 43.
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Figure 4.
TM0077 oligomeric assembly. (A) Surface representation of the biological unit of the
TM0077-Native hexamer with each monomer in a different color (left). The “cross section”
shows the entrances on either side of the assembly and the internal cavity (center), and a 90°
rotated view of the TM0077-Native hexamer, with a close-up view of the open central hole
(right). (B) Surface representation of the biological hexamer unit of CAH from B. subtilis 40
(left) and the TM0077-SeMet hexamer with a close-up view of the blocked central hole
(right).
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Figure 5.
Effect of temperature and pH on esterase activity. (A) The esterase activity was studied
using pNP-C2 as a substrate at temperatures ranging from 40–100°C. The inset shows the
temperature dependence as an Arrhenius plot. (B) Thermal stability of TM0077 at 90°C. (C)
The effect of pH on esterase activity studied using pNP-C2 as a substrate at pH values in the
range of 4.8–9.2.
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Figure 6.
TM0077 catalytic site. (A) Surface representation of the TM0077 catalytic site, with His303,
Asp274 and the intermediate DEP-modified Ser188 shown as sticks. The two binding
pockets are indicated with S1 and S2. (B) Apo TM0077 with a bound chloride ion (green
sphere), (C) TM0077 with PMS-modified Ser188 and (D) TM0077 with DEP-modified
Ser188. The catalytic residues are shown as sticks, with the hydrogen bonds shown as
dashed lines. Carbon atoms are in green (apo), cyan (PMS) or blue (DEP), oxygen atoms in
red, sulfur atoms in yellow and phosphate in orange. Electron density omit maps shown for
inhibitor modified Ser188 contoured at 1σ show that the PMS and DEP are covalently
bonded to Ser188 in (C) and (D), respectively. Distances are shown in Ångströms.
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Figure 7.
Conformational change of Ser188 Oγ. The Oγ atom of the Ser188 is rotated ~110° between
the native apo structure (cyan) and (A) the complexed PMS-modified Ser188 structure
(pink), (B) the DEP-modified Ser188 structure (light blue) and (C) the SeMet structure
(purple). The different hydrogen bonds made for the Ser Oγ in the native versus complexed
structures are shown as dashed black lines with distances in Ångströms.
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Table I
Summary of crystal parameters, data collection, and refinement statistics
TM0077-SeMet
TM0077-Native
TM0077-PMS
TM0077-DEP
Space group
P 21
P 21 21 21
P 21 21 21
P 21 21 21
Unit cell parameters
a=152.64Å b=130.95Å
c=157.82Å β=118.90°
a=103.46Å
b=103.79Å
c=221.02Å
a=103.57Å
b=104.50Å
c=221.61Å
a=103.80Å
b=104.43Å
c=221.64Å
Data collection
λ1 MAD-Se
λ2 MAD-Se
Wavelength (Å)
0.9791
0.9183
0.9765
0.9765
0.9765
Resolution range (Å)
29.6 – 2.10
29.6 – 2.10
48.8 – 2.50
49.0 – 2.40
49.0 – 2.12
No. observations
1,119,236
1,100,249
1,222,016
765,546
989,949
No. unique reflections
293,140
291,757
83,045
94,681
123,070
Completeness (%)
93.0 (61.8)a
92.6 (60.8)
100 (100)
100 (100)
89.8 (53.5)
Mean I/σ(I)
9.1 (2.4)a
9.6 (2.2)
14.4 (2.9)
11.5 (3.4)
15.3 (2.2)
Rmerge on I (%)
12.3 (52.5) a
11.9 (57.9)
20.7 (109.7) c
18.0 (67.4)
9.5 (51.9)
Rmeas on I (%)
14.3 (62.2) a
13.9 (68.7)
21.4 (113.6)
19.2 (71.9)
10.2 (60.2)
Rpim on I (%)
7.2 (32.7) a
7.1 (36.2)
5.5 (29.2)
6.7 (24.9)
3.5 (29.2)
Highest resolution shell (Å)
2.15 – 2.10
2.15 – 2.10
2.56 – 2.50
2.46 – 2.40
2.18 – 2.12
Model and refinement statistics
Resolution range (Å)
29.6 – 2.10
48.8 – 2.50
49.0–2.40
49.0 – 2.12
No. reflections (total)
293,097 b
83,045
94,680
122,994
No. reflections (test)
14,726
4,200
4,742
6,188
Completeness (% total)
92.8
100.0
100.0
89.8
Data set used in refinement
λ1 MAD-Se
Cutoff criteria
|F| > 0
|F| > 0
|F| > 0
|F| > 0
Rcryst
0.186
0.167
0.160
0.167
Rfree
0.223
0.212
0.208
0.205
Stereochemical parameters
Restraints (RMSD observed)
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TM0077-SeMet
TM0077-Native
TM0077-PMS
TM0077-DEP
Bond angle (°)
1.48
1.47
1.53
1.44
Bond length (Å)
0.018
0.017
0.017
0.015
Av. isotropic B-value (Å2)
27.9
24.7
19.4
19.6
ESU based on Rfree
0.17
0.25
0.22
0.18
Water molecules/other solvent molecules
2,464/1
507/24
946/17
987/23
PDB ID
1vlq
3m81
3m82
3m83
aHighest resolution shell
ESU = Estimated overall coordinate error 16,66.
Rmerge=ΣhklΣi|Ii(hkl)-<I(hkl)>|/ΣhklΣiIi(hkl), Rmeas(redundancy-independent Rmerge)=Σhkl[Nhkl/(Nhkl-1)]1/2Σi|Ii(hkl)-<I(hkl)>|/ΣhklΣiIi(hkl), and Rpim(precision-indicating Rmerge)=Σhkl[1/
(Nhkl-1)]1/2Σi|Ii(hkl)-<I(hkl)>|/ΣhklΣiIi(hkl) 67–69.
Rcryst = Σ| |Fobs|-|Fcalc| |/Σ|Fobs| where Fcalc and Fobs are the calculated and observed structure factor amplitudes, respectively.
Rfree = as for Rcryst, but for 5.0 % of the total reflections chosen at random and omitted from refinement.
bTypically, the number of unique reflections used in refinement is slightly less than the total number that were integrated and scaled. Reflections are excluded due to systematic absences, negative
intensities, and rounding errors in the resolution limits and cell parameters.
cRmerge of the highest resolution shell is high due to high redundancy (14.7). However, the completeness and mean I/σ of the highest resolution shell are reasonable, and these data were included in the
refinement.
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Table II
Kinetic parameters for hydrolysis of various esters
Ester
Km (mM)
kcat (s−1)
kcat/Km (s−1 mM−1)
pNP-Acetate
0.185 ± 0.026
57.5 ± 2.2
310.8 ± 45.3
pNP-Propionate
0.137 ± 0.013
41.3 ± 1.1
301.5 ± 29.7
2-O-acetyl pNP-Xyl
3.6 ± 0.5
76.1 ± 19.2
21.1 ± 6.1
3-O-acetyl pNP-Xyl
4.2 ± 0.4
70.1 ± 7.7
16.7 ± 2.4
4-O-acetyl pNP-Xyl
4.0 ± 0.1
78.6 ± 12.9
19.7 ± 3.3
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|
3M82
|
Crystal structure of Acetyl xylan esterase (TM0077) from THERMOTOGA MARITIMA at 2.40 A resolution (PMSF inhibitor complex structure)
|
Functional and structural characterization of a thermostable
acetyl esterase from Thermotoga maritima
Mark Levisson1,*, Gye Won Han2,3,*, Marc C. Deller2,3, Qingping Xu2,4, Peter Biely5, Sjon
Hendriks1, Lynn F. Ten Eyck6,7, Claus Flensburg8, Pietro Roversi8, Mitchell D. Miller2,4,
Daniel McMullan9, Frank von Delft2,3,‡, Andreas Kreusch10, Ashley M. Deacon2,4, John van
der Oost1, Scott A. Lesley2,3,10, Marc-André Elsliger2,3, Servé W. M. Kengen1,†, and Ian A.
Wilson2,3,†
1Laboratory of Microbiology, Wageningen University, 6703 HB, Wageningen, The Netherlands
2Joint Center for Structural Genomics, http://www.jcsg.org 3Department of Molecular Biology, The
Scripps Research Institute, La Jolla, California 92037 4Stanford Synchrotron Radiation
Lightsource, SLAC National Accelerator Laboratory, Stanford University, Menlo Park, California
92045 5Institute of Chemistry, Slovak Academy of Sciences, 845 38 Bratislava, Slovakia
6Department of Chemistry and Biochemistry, University of California at San Diego, La Jolla,
California 92093-0505 7San Diego Supercomputer Center, University of California at San Diego,
La Jolla, California 92093-0505 8Global Phasing Ltd. Sheraton House, Castle Park, Cambridge
CB3 0AX, United Kingdom 9Protein Therapeutics Department, Genomics Institute of the Novartis
Research Foundation, San Diego, California 92121 10Protein Sciences Department, Genomics
Institute of the Novartis Research Foundation, San Diego, California 92121
Abstract
TM0077 from Thermotoga maritima is a member of the carbohydrate esterase family 7 and is
active on a variety of acetylated compounds, including cephalosporin C. TM0077 esterase activity
is confined to short-chain acyl esters (C2-C3), and is optimal around 100°C and pH 7.5. The
positional specificity of TM0077 was investigated using 4-nitrophenyl-β-D-xylopyranoside
monoacetates as substrates in a β-xylosidase-coupled assay. TM0077 hydrolyzes acetate at
positions 2, 3 and 4 with equal efficiency. No activity was detected on xylan or acetylated xylan,
which implies that TM0077 is an acetyl esterase and not an acetyl xylan esterase as currently
annotated. Selenomethionine-substituted and native structures of TM0077 were determined at 2.1
Å and 2.5 Å resolution, respectively, revealing a classic α/β-hydrolase fold. TM0077 assembles
into a doughnut-shaped hexamer with small tunnels on either side leading to an inner cavity,
which contains the six catalytic centers. Structures of TM0077 with covalently bound
phenylmethylsulfonyl fluoride (PMSF) and paraoxon were determined to 2.4 Å and 2.1 Å,
respectively, and confirmed that both inhibitors bind covalently to the catalytic serine (Ser188).
Upon binding of inhibitor, the catalytic serine adopts an altered conformation, as observed in other
esterase and lipases, and supports a previously proposed catalytic mechanism in which this Ser
hydroxyl rotation prevents reversal of the reaction and allows access of a water molecule for
completion of the reaction.
†Correspondence to: Ian A. Wilson, Ph.D., Department of Molecular Biology, The Scripps Research Institute, La Jolla, California
92037; (858) 784-2939 Fax: (858) 784-2980; wilson@scripps.edu or Servé W. M. Kengen, Ph.D., Laboratory of Microbiology,
Wageningen University, 6703 HB, Wageningen, The Netherlands; 31 317 483737, Fax: 31 317-483829; serve.kengen@wur.nl.
*ML and GWH contributed equally to this work.
‡Current address: The Structural Genomics Consortium, Roosevelt Drive, Headington, Oxford OX3 7DQ, UK
NIH Public Access
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Published in final edited form as:
Proteins. 2012 June ; 80(6): 1545–1559. doi:10.1002/prot.24041.
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Keywords
Acetyl esterase; Thermotoga maritima; crystal structure; α/β hydrolase; inhibitor; serine rotation
INTRODUCTION
Thermotoga maritima is a hyperthermophilic bacterium that grows optimally at 80°C and is
able to metabolize a variety of simple and complex carbohydrates, including glucose,
sucrose, starch, cellulose, and xylan 1. Its carbohydrate utilization potential was confirmed
by analysis of its sequenced genome 2. The xylan degrading pathway of T. maritima has
been studied using microarrays 2–4, and several genes encoding transporters, xylanases, and
a β-xylosidase have been identified. Among the enzymes with a differential expression
pattern in the microarray was a predicted acetyl xylan esterase (locus tag TM0077,
axeA) 3,5. Depending on the source, the xylan backbone may contain a varying degree of
acetylated xylose residues. Therefore, in addition to xylanases and xylosidases, the complete
degradation of xylan requires esterases/deacetylases 6.
Presently, esterases and deacetylases that are active on carbohydrate substrates have been
classified into 16 families by Henrissat and coworkers (Carbohydrate-Active enZymes
Server (CAZy)) 7. According to this classification, the predicted acetyl xylan esterase from
T. maritima would be a member of family 7 of the carbohydrate esterases (CE7). In addition
to the acetyl xylan esterase activity, enzymes in the CE7 family are rather unusual in that
they display a high specific activity towards the antibiotic cephalosporin C [(Fig. 1(a-b)] 8.
Cephalosporins belong to the β-lactam class of antibiotics, which also includes penicillin,
and affect bacterial cell growth by inhibiting the penicillin-binding-protein that cross-links
peptide glycans required for cell wall formation 9. The production of deacetylated
cephalosporins is of great interest because these compounds are valuable building blocks for
the production of semi-synthetic β-lactam antibiotics10,11.
To explore the catalytic capacity of the predicted acetyl xylan esterase from T. maritima and
gain a better insight into the structure and function of the family 7 carbohydrate esterases,
TM0077 was expressed and purified, and three-dimensional structures of the native enzyme
and its complexes with phenylmethylsulfonyl fluoride (PMSF) and paraoxon inhibitors,
were determined by x-ray crystallography. Furthermore, the enzyme was functionally
characterized, and various biochemical properties including the positional specificity of the
esterase were investigated.
MATERIALS AND METHODS
Gene cloning
TM0077 was selected as part of the Joint Center for Structural Genomics (JCSG) effort on
complete structural coverage of the T. maritima soluble proteome as a large-scale center for
high-throughput structure determination funded under the NIHGMS Protein Structure
Initiative (PSI) 12. The gene encoding TM0077 (GenBank: AAD35171.1, GI:4980565;
SwissProt: Q9WXT2) was amplified by polymerase chain reaction (PCR) from genomic
DNA using PfuTurbo DNA polymerase (Stratagene) and primers corresponding to the
predicted 5′ and 3′ ends. The PCR product was cloned into plasmid pMH1, which encodes
an expression and purification tag (MGSDKIHHHHHH) at the amino terminus of the
protein. The cloning junctions were confirmed by DNA sequencing.
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TM0077-SeMet protein production and purification
Protein production was performed in a selenomethionine-containing medium using the
Escherichia coli methionine auxotrophic strain DL41. Expression was induced by the
addition of 0.15% L-arabinose. At the end of fermentation, cells were harvested and
subjected to one freeze/thaw cycle, and subsequently sonicated in Lysis Buffer [50 mM Tris
pH 7.9, 50 mM NaCl, 1 mM MgCl2, 0.25 mM Tris(2-carboxyethyl)phosphine hydrochloride
(TCEP), 1 mg/ml lysozyme] and the lysate was centrifuged at 3,400 × g for one hour. The
soluble fraction was applied to nickel-chelating resin (GE Healthcare) pre-equilibrated with
Equilibration Buffer [50 mM potassium phosphate pH 7.8, 300 mM NaCl, 10% (v/v)
glycerol, 0.25 mM TCEP] containing 20 mM imidazole. The resin was washed with
Equilibration Buffer containing 40 mM imidazole, and the protein was eluted with Elution
Buffer [20 mM Tris pH 7.9, 300 mM imidazole, 10% (v/v) glycerol, 0.25 mM TCEP]. The
eluate was buffer exchanged with Buffer Q [20 mM Tris pH 7.9, 5% (v/v) glycerol, 0.25
mM TCEP] containing 50 mM NaCl and applied to a RESOURCE Q column (GE
Healthcare) pre-equilibrated with the same buffer. The protein was eluted using a linear
gradient of 50 to 500 mM NaCl in Buffer Q and purified further with a HiLoad 16/60
Superdex 200 column (GE Healthcare), using Crystallization Buffer [20 mM Tris pH 7.9,
150 mM NaCl, 0.25 mM TCEP] as the mobile phase. For crystallization trials, the peak
Superdex 200 fractions were concentrated to ~15 mg/mL by centrifugal ultrafiltration
(Millipore). Molecular weight and oligomeric state of TM0077 were determined using a 1
cm × 30 cm Superdex 200 column (GE Healthcare) coupled with miniDAWN static light
scattering (SEC/SLS) and Optilab differential refractive index detectors (Wyatt
Technology). The mobile phase consisted of 20 mM Tris pH 8.0, 150 mM NaCl, and 0.02%
(w/v) sodium azide.
Native TM0077 production and purification
For protein production, E. coli DL41 cells were grown in LB medium for 8 hours (an
OD600 well above 2.0 was reached). Subsequently, the culture was induced by adding
0.15% L-arabinose and incubated another 16 hours at 37°C. Cells were harvested by
centrifugation at 10,000 × g for 20 min. The cell pellet was resuspended in 30 ml of Lysis
Buffer 2 [50 mM Tris-HCl pH 8.0, 50 mM NaCl, 10 mM imidazole, 0.25 mM TCEP]. The
cells were disrupted by two passages through a French press at 110 MPa. The crude cell
extract was treated with DNAse I at room temperature for 30 min and subsequently
centrifuged at 43,000 × g for 30 min in order to remove cell debris. The supernatant was
heated at 70°C for 25 min and then centrifuged to remove the precipitated proteins. The
supernatant was filtered and loaded onto a nickel-chelating column packed with 20 ml of Ni-
NTA His-Bind Resin (Novagen) and equilibrated in 50 mM Tris-HCl pH 8.0, 300 mM
NaCl, 2% (v/v) glycerol, and 0.25 mM TCEP. The column was washed with 20 mM
imidazole in the same buffer, and proteins were subsequently eluted with a linear gradient of
20–500 mM imidazole in the same buffer. Fractions containing esterase activity were pooled
and loaded onto a HiPrep Desalting column (GE Healthcare) equilibrated with 20 mM Tris-
HCl pH 8.0, 150 mM NaCl, and 0.25 mM TCEP. The homogeneity of the protein was
checked by SDS-PAGE, and activity staining of the SDS-PAGE gel was performed using α-
napthyl acetate, as described previously 13. The protein concentration was determined at 280
nm using a NanoDrop ND-1000 Spectrophotometer.
Crystallization
Crystals of selenomethionine-substituted TM0077 were obtained by hanging drop vapor
diffusion against a 250 μl crystallization solution consisting of 20% (w/v) PEG-3000, 0.1 M
HEPES pH 7.5, 0.2 M NaCl. Drops consisted of 0.5 μl protein and 0.5 μl crystallization
solution. Native TM0077 was crystallized using nanodrop vapor diffusion techniques
against a crystallization solution consisting of 0.2 M calcium acetate hydrate, 20% (w/v)
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PEG 3350, pH 7.3 at 20°C. Protein was concentrated to 22.8 mg/ml. Drops consisted of 100
nl protein and 100 nl of crystallization solution and a 60 μl reservoir of crystallization
solution. Crystals of TM0077 in complex with inhibitors PMSF or paraoxon were obtained
at 4°C in the same conditions with the same reagents as the native crystals. PMSF or
paraoxon were added in a molar ratio of 1:3 (protein:inhibitor).
Data collection
For cryoprotection, the TM0077-SeMet crystal was transferred to crystallization solution
supplemented with 15% (v/v) glycerol. The crystal was mounted in a cryoloop and
subsequently flash-cooled in liquid nitrogen. X-ray data were collected at 100 K on
beamline BL9-2 at the Stanford Synchrotron Radiation Lightsource (SSRL, Menlo Park,
CA) using a Quantum 4 CCD detector (ADSC). A TM0077-SeMet MAD data set was
collected to 2.1 Å resolution and the data were indexed in monoclinic space group P21, with
unit cell parameters a = 152.6 Å, b = 131.0 Å, and c = 157.8 Å, and β=118.9°, and 12
molecules in the asymmetric unit. Data were indexed and integrated with DENZO and then
scaled with SCALEPACK 14.
Native TM0077, TM0077-PMSF complex (TM0077-PMS) and TM0077-paraoxon complex
(TM0077-DEP) crystals were transferred to crystallization solution supplemented with 10%
(v/v) ethylene glycol and flash-cooled to 100K. Data were collected at beamline 5.0.3 of the
Advanced Light Source (ALS, Berkeley, CA) and processed with the HKL2000 package 14.
The native data set was collected to 2.5 Å resolution, and TM0077-PMS and TM0077-DEP
data sets were collected to 2.4 and 2.1 Å, respectively. All data were indexed in
orthorhombic space group P212121, with unit cell parameters approximately a=103Å
b=104Å c=221Å (See Table 1), and six molecules in the asymmetric unit. Data reduction
and refinement statistics for TM0077-SeMet, TM0077-Native, TM0077-PMS and TM0077-
DEP are summarized in Table I.
Structure solution and refinement
The TM0077-SeMet structure was solved by MAD phasing method using a two-wavelength
MAD dataset. At the time of the initial data collection (2001), the structure determination of
Se-MAD TM0077 posed a significant challenge to crystallographic programs, which were
still under active development. As a result, modifications were made in various structure
determination and refinement programs to achieve success. For initial phasing, SHELXD 15
was used to find candidate SeMet substructure sites. Attempts to complete phasing were
unsuccessful due to the translational non-crystallographic symmetry (NCS) (not recognized
initially). Self-consistent sets (partial sets) were found using the CCP4 program
PROFESSS 16 and additional SeMet sites were found by SHELXD, and added to these
partial sets. The SHARP 17 run did not complete initially; however, updates of SHARP and
ARP/wARP eventually helped to resolve issues and an initial trace was obtained by ARP/
wARP. The structure was then refined with BUSTER 18 using tight NCS restraints to an
Rcryst and Rfree of 18.6% and 22.3%, respectively. Model building was performed using
O 19 and the structure was refined using Refmac5 20. Refinement statistics are summarized
in Table I. The final model contains 12 protein molecules (chains A-L) in the asymmetric
unit each consisting of residues 2-323. The MolProbity 21 Ramachandran plot analysis
showed that 97.4% of all residues are in favored regions with a single outlier, Gln120 of
chain B, which is supported by unambiguous electron density. Ramachandran outlier
Gln120 of chain B of TM0077-SeMet is due to crystal packing with chain C. The backbone
carbonyl oxygens of Gln120 and Gly119 of chain B makes hydrogen bonds with the
backbone nitrogen of Gln140 of chain C (3.19 and 3.11 Å, respectively).
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The native TM0077 structure, TM0077-PMS and TM0077-DEP structures were solved by
molecular replacement using PHASER 22,23 with the TM0077-SeMet hexamer coordinates
(pdb: 1vlq; A-F chains) as a search model. One hexamer was successfully located and the
structure was further refined with Refmac5 20 using tight NCS restraints to an Rcryst and
Rfree of 16.7% and 21.2% (native TM0077), 16.0% and 20.8% (TM0077-PMS) and 16.7%
and 20.5% (TM0077-DEP), respectively. Iterative cycles of refinement and building were
performed with Refmac5, Phenix 24,25 and Coot 26. All other crystallographic manipulations
were carried out with the CCP4 package 16. Refinement statistics are summarized in Table I.
The final model of native TM0077 contains residues 3-324 (chains A, B, C, D and F) and
3-323 (chain E) in the asymmetric unit. Analysis of main-chain torsion angles using
MolProbity 21 showed that 97.8% of the residues are in favored regions of the
Ramachandran plot with 0.2% outliers (Asn302 of chains B, C and D), which are supported
by unambiguous electron density. The final model of TM0077-PMS contains residues 3-323
for all chains in the asymmetric unit with 97.5% of the residues in favored regions with
0.2% outliers (Asn302 of chains A, B, D and F). The final model of TM0077-DEP contains
residues 0-324 for (chains A, B, C and F) and 0-323 (chains D and E) in the asymmetric
unit, respectively, with 97.6% of the residues in favored region of the Ramachandran plot
with 0.2% outliers (Asn302 of B, C, D and F chains). Ramachandran outlier Asn302 in the
TM0077-Native, TM0077-PMS and TM0077-DEP structures is a neighbor to the catalytic
triad residue His303 and may reflect a slightly different state for these structures compared
to the Se-Met structure.
Structure validation and deposition
The quality of the crystal structure was analyzed using the JCSG Quality Control server
(http://smb.slac.stanford.edu/jcsg/QC). This server processes the coordinates and data
through a variety of validation tools including AutoDepInputTool 27 MolProbity 21,
WHATIF 5.0 28, RESOLVE 29, MOLEMAN2 30 as well as several in-house scripts, and
summarizes the results. Protein quaternary structure analysis were performed using the PISA
server 30. Figures were prepared with PyMOL (DeLano Scientific) 31. RMSD values were
calculated using the ProCKSI-Server 32. The structural data have been deposited in the
RCSB Protein Data Bank (PDB) with accession codes 1vlq for TM0077-SeMet, 3m81 for
TM0077-native, 3m83 for TM0077-DEP and 3m82 for TM0077-PMS.
Enzyme assays
Esterase activity was measured using p-nitrophenyl esters as described previously 13.
Briefly, the standard assay consisted of activity measurements with 0.2 mM p-nitrophenyl
acetate as substrate in 50 mM citrate-phosphate (pH 6) at 70°C. The p-nitrophenol liberated
was measured continuously at 405 nm on a Hitachi U-2001 spectrophotometer with a
temperature-controlled cuvette holder. Extinction coefficients of p-nitrophenol were
determined prior to each measurement. Kinetic parameters were determined by direct fitting
the data, obtained from multiple measurements, to the Michaelis–Menten curve (Tablecurve
2d, version 5.0).
The effect of pH on esterase activity was studied in the pH range from 5 to 10. The buffers
used were 50 mM citrate-phosphate (pH 5–8) and 50 mM CAPS (3-(cyclohexylamino) 1-
propanesulphonic acid) (pH 9.5–10). The pH of the buffers was set at room temperature, and
temperature corrections were made using their temperature coefficients: −0.0028 pH/°C for
citrate-phosphate buffer and −0.018 pH/°C for CAPS buffer. The effect of temperature on
esterase activity was studied in the range of 40–100°C using 0.2 mM p-nitrophenyl acetate
as substrate. Enzyme thermostability was determined by incubating the enzyme in a 50 mM
Tris-HCl, 150 mM NaCl (pH 7.8) buffer at 90°C and 100°C for various time intervals.
Residual activity was determined in the standard assay.
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Inhibition kinetics of PMSF and paraoxon were determined as described for the
acetylcholinesterase from electric eel 33. All experiments were performed at 70°C in 50 mM
citrate-phosphate (pH 6) buffer and 0.2 mM p-nitrophenyl acetate as substrate. The kinetic
constants for the inhibition of TM0077 with PMSF and paraoxon were measured in the
concentration range of 1.0–10.0 mM and 0.2–1.0 mM, respectively.
Deacetylase activity was determined using high-performance liquid chromatography
(HPLC) by measuring the amount of acetic acid released from the substrates cephalosporin
C, 7-aminocephalosporanic acid, glucose-pentaacetate and acetylated xylan. Xylan was
acetylated by the method described by Johnson 34. The reaction mixture contained 0.9 ml of
substrate solution (dissolved in 50 mM Tris-HCl, pH 7.5) and 0.1 ml of enzyme solution,
and was incubated at 37°C for various time intervals (0–10 min). The reaction was stopped
by adding 0.2 ml of stop solution (100 mN H2SO4 and 30 mM crotonate) and placing the
sample on ice. The conditions for HPLC were as follows: column, KC811 Shodex;
detection, RI and UV detectors; solvent, 3 mN H2SO4; flow rate, 1.5 ml/min; temperature,
30°C; internal standard, crotonate. One unit of enzyme activity was defined as the amount of
enzyme that releases one μmol of acetic acid per minute.
Activity on xylan was measured quantitatively using DMSO-extracted xylan (1%
polysaccharide solution in 0.1 M sodium phosphate buffer pH 6) at 60°C 35. Xylan will
precipitate as a consequence of deacetylation, resulting in a rapid turbidity of the solution.
Positional specificity assay
The positional specificity of TM0077 was investigated using an enzyme-coupled assay on
monoacetylated 4-nitrophenyl β-D-xylopyranosides (pNP-Xyl) as described 36. The β-
xylosidase XloA (locus tag: TM0076) from T. maritima was cloned into the vector pET24d
in frame with a C-terminal His6-tag. The enzyme was expressed and purified as described
above for native TM0077. Activity of XloA was confirmed by measuring the release of p-
nitrophenol at 405 nm from the substrate 4-nitrophenyl β-D-xylopyranoside.
The enzyme-coupled assay was performed at 60°C in a total volume of 125 μl, which
contained 0.1 M sodium phosphate (pH 6 or 7), 2-O-, 3-O-, or 4-O-acetyl pNP-Xyl, the β-
xylosidase XloA, and TM0077. Stable 50x-concentrated stock solutions of the substrates
were prepared in DMSO. The reaction was started by addition of 2.5 μl of a stock solution
to a preheated reaction mixture consisting of phosphate buffer, auxiliary β-xylosidase XloA
in excess, and TM0077. The reaction was terminated by addition of 800 μl of a 2% solution
of Na2CO3. Liberated p-nitrophenol was determined at 405 nm against substrate and
enzyme blanks. A short incubation time for activity determination was used to suppress
acetyl migration on the xylopyranosyl-ring, which is significant at pH 6 or 7 37. The kinetic
constants were determined at pH 7 and 60°C with reaction times of 2 or 5 minutes.
RESULTS and DISCUSSION
In silico analysis
TM0077 consists of 325 amino acids with a calculated molecular mass of 37 kDa. Sequence
analysis, using the SignalP 3.0 server, revealed that TM0077 has no predicted signal
sequence and is, therefore, believed to be an intracellular enzyme. Analysis of the gene
organization indicates that the TM0077 gene co-localizes with genes encoding a xylanase
(TM0070) 38, ABC transporter components (TM0071-TM0075), and a β-xylosidase
(TM0076) 39. BLAST-P analysis showed that TM0077 has highest similarity to putative
acetyl esterases, acetyl xylan esterases and cephalosporin C deacetylases. Among the
BLAST results, a predicted acetyl xylan esterase-related protein from T. maritima (locus
tag: TM0435) was also identified. TM0077 was compared with other members of the CE7
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family using structure-based, multi-sequence alignment and the putative catalytic triad,
Ser188, Asp274, and His303, was identified from conservation throughout the analyzed
sequences. The putative nucleophilic serine (Ser188) is located within a conserved
pentapeptide consensus sequence, Gly-Xaa-Ser-Gln-Gly, typical of this family. Previously, a
signature sequence motif, [RGQ]-(x:~70)-[GxSQG]-(x:~115)-[HE] (where x indicates any
amino acid), had been suggested for the CE7 family based on an aminoacid alignment of 12
sequences 40. In an updated alignment consisting now of 50 sequences, we observed many
sequences that have this signature motif, but it is not conserved throughout the entire family
(See Supporting Information and Fig. S1 for the multi-sequence alignment).
Overall structure
The crystal structure of seleno-methionine incorporated TM0077 (TM0077-SeMet) was
determined to 2.1 Å resolution by multi-wavelength anomalous dispersion (MAD) (Table I)
with twelve molecules per asymmetric unit. A native apo structure (TM0077-Native) was
determined in a different space group (see Methods) to 2.5 Å by molecular replacement,
using TM0077-SeMet as a search model, with six molecules in the asymmetric unit (Table
I). Each monomer of the native hexamer contained a calcium ion (see below) bound by
Lys22, Glu26, and Asp25 via a bridging water molecule. Superposition of the TM0077-
SeMet and the TM0077-Native structures gave a root-mean-square difference (RMSD) of
0.12 Å over 321 Cα atoms, which indicates that these structures are nearly identical as
expected.
In general, the TM0077 structure resembles the canonical α/β-hydrolase fold, which
consists of a central, twisted, eight-stranded β-sheet surrounded by α-helices on both sides,
with β2 antiparallel to the other strands. TM0077 deviates slightly from the canonical α/β-
hydrolase fold at two locations: a three-helix insertion after strand β6 and an extension of
the N-terminus (Fig. 2). Insertions after β6 or β7 are common for α/β-hydrolases and are
proposed to help shape the substrate-binding site 41. The N-terminus is extended by two
helices (αA-1 and αA-2) and an antiparallel β-strand (β-1) that aligns with the other eight β-
strands (β1-β8) and extends the central β-sheet. This nine-stranded β-sheet is highly twisted,
and β-1 and β8 at the extreme edges are rotated approximately 130° relative to each other.
Helices αA and αB both contain a short 310-helix segment at their N-terminus. Helices
αA-1, αA-2, αB, αC, αD, αD1, αD2, αD3, αE, and the 310-helix η2 are located on one side
of the central β-sheet, and helices αA, αF and the 310-helix η1 are on the other side.
A structural similarity search was performed using the program DALI 42. Monomer A of the
TM0077-SeMet structure was used as a search model and similarity was found with
cephalosporin C deacetylases, acetyl xylan esterases, acylamino-releasing enzymes,
dipeptidyl peptidases and some esterases and lipases. TM0077 is structurally most similar to
cephalosporin C deacetylase (CAH) from B. subtilis (PDB: 1ods) 40, acetyl xylan esterase
(AXE) from B. pumilus (PDB: 3fvr and 2xlb), acetyl xylan esterase (AXE1) from
Thermoanaerobacterium sp. JW/SL YS485 (PDB: 3fcy), and acylpeptide hydrolase/esterase
apAPH from Aeropyrum pernix K1 (PDB: 1ve6) 43. The sequence identity between
TM0077 and CAH is 41% and the two structures align with a Z-score of 46 and an RMSD
of 1.5 Å over 312 Cα atoms. The sequence identity with apAPH is 17% with a Z-score of
23.3 and an RMSD of 2.3 Å over 230 Cα atoms. Superpositions of TM0077 with CAH and
with apAPH are shown in Fig. 3.
Quaternary structure
The crystal structure of TM0077-SeMet contains two hexamers in the asymmetric unit that
are related by a non-crystallographic two-fold axis. Each hexamer contains a dimer of
trimers with a back-to-back arrangement (Fig. 4). The apo and the complex crystals
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contained one hexamer in the asymmetric unit. Crystallographic packing analysis using
PISA (EBI) 44 indicated that the relevant physiological oligomeric state of TM0077 is a
hexamer, which was confirmed by size exclusion chromatography coupled with static light
scattering. Further analyses of the hexameric assembly indicated that two main interfaces
play an essential role in complex formation. The first interface between subunit A and B
(green and cyan in Fig. 4) (identical to C/D and E/F) is stabilized by seven hydrogen bonds
on average and has a buried surface area of 1024 Å2 contributed by each chain. The second
interface between A and F (green and purple in Fig. 4) (B/C and D/E) is stabilized by 17
hydrogen bonds on average with a buried surface area of 1079 Å2 contributed by each chain
However, a multiple sequence alignment of TM0077 with other CE7 esterases showed that
the residues involved in these two main interfaces are not conserved. Other secondary
interfaces bury around 514 Å2 contributed by each chain. The hexamer has a total buried
surface area of 18,860 Å2, which is approximately 30% of the total surface area.
Approximately 3,143 Å2 per monomer is, therefore, buried upon complex formation.
The TM0077 hexamer has a doughnut-shape when viewed from the side, with the six active
sites located in the interior of the complex, where they line an oval-shaped cavity [Fig. 4(a)].
This cavity is accessible via two entrances, one on each side of the flat hexamer. Each of
these entrances is approximately 13 Å wide and connects to a short tunnel or pore spanning
approximately 10 Å to reach the inner cavity. Interestingly, in the TM0077-SeMet hexamer,
the entrance to the internal cavity is blocked by three phenylalanine residues (Phe4), one for
each of three monomers that compose half of the hexamer [Fig. 4(b)]. Residue Phe4 is
located in the mobile N-terminus (high B-values), which may indicate some flexibility or
multiple conformations.
Calcium ions were identified, by the electron density and coordination geometry, supported
by their presence in the crystallization reagents, in the native TM0077, TM0077-PMS and
TM0077-DEP structures, but not in the TM0077-SeMet structure. The SeMet protein was
crystallized without any calcium in the crystallization reagents. In each subunit of the
hexamer, one calcium ion is located at the N-terminal region of helix αA-1, and is
coordinated by the backbone carbonyl of Lys22 and the Glu26 carboxylate. The remainder
of the calcium coordination sphere is filled with waters from a neighboring solvent channel
present in all molecules in the asymmetric unit. The Asp25 carboxylate contributes to the
calcium binding via one of the coordinating water molecules. Another calcium ion is bound
in a crystal packing interface between chain A and chain C′ of a crystallographic symmetry-
related hexamer. This calcium is coordinated by the carboxylates of GluA45 and AspA58
from one chain and the carboxylate from Glu C’45 (bidentate coordination) of the
symmetry-related chain with three water molecules completing a capped-octahedral
coordination sphere. An equivalent calcium binding site is also observed in the crystal
packing interface between chains D and B′. No significant increase or reduction of activity
of TM0077 was observed in the presence of calcium ions or EDTA. Therefore, it seems that
these calcium ions are not important for activity. On the other hand, calcium may help
stabilize the structure. No calcium was present in the B. subtilis CAH structure 40; however,
Lys22, Glu26 and Ser25 are conserved and may also act as a calcium binding site.
Enzyme activity
The activity of TM0077 was investigated using p-nitrophenol esters with varying acyl-chain
length, ranging from C2 to C18. TM0077 is only active on the short-chain p-nitrophenol
esters of acetate and propionate and does not hydrolyze esters with acyl chains longer than
four carbons. No significant difference was found in the catalytic efficiency (kcat/Km) for the
hydrolysis of p-nitrophenyl with acyl chains containing 2 to 3 carbons (Table II) [Fig. 1(c)].
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The effect of temperature on activity was studied using p-nitrophenyl acetate as substrate.
The esterase activity increased from 40°C upwards until 100°C [Fig. 5(a)]. An Arrhenius
analysis resulted in linear plots in the temperature ranges of 40–60°C and 60–100°C [Fig.
5(a); inset], with calculated activation energies for the formation of the enzyme substrate-
enzyme complex of 33.7 and 21.9 kJ/mol, respectively. The transition or break in linearity
of the Arrhenius plot at 60°C (1000/T (K) = 3.0) could indicate some conformational change
of the enzyme. TM0077 is fairly resistant to thermal inactivation. An approximate 50%
transient increase in activity is seen during the first 10 to 20 minutes when the enzyme is
incubated at 90°C. After 30 minutes, inactivation of function occurs by first order kinetics
with a half-life of approximately 120 minutes [Fig. 5(b)]. A transient activation has also
been observed for other thermophilic esterases, such from Sulfolobus shibatae 45, and it is
believed that a high temperature is needed in order to obtain the optimal conformation for
catalysis. TM0077 was not stable at 100°C, resulting in a half-life of less than 5 minutes.
However, the optimum temperature and thermal stability of TM0077 are still considerably
higher than those reported for other characterized CE7 esterases, including the
Thermoanaerobacterium enzyme that has a temperature optimum of 80°C and a half-life of
1h at 75°C 46.
The effect of pH on activity was measured in the pH range of 4.8 to 9.2 using the substrate
p-nitrophenyl acetate. TM0077 displayed maximum activity at approximately pH 7.5 [Fig.
5(c)], which is comparable to other CE7 esterases, such as the acetyl xylan esterases from
Thermoanaerobacterium sp. strain JW/SL-YS485 46.
Positional specificity
The positional specificity of TM0077 was tested on three monoacetates of 4-nitrophenyl β-
D-xylopyranoside (pNP-Xyl). To determine the enzyme activity, the β-xylosidase XloA 39
(TM0076) from T. maritima is required as an auxiliary enzyme. This thermostable XloA
enzyme was, therefore, cloned, heterologously expressed, purified to homogeneity, and its
activity was confirmed by measuring release of p-nitrophenol from the substrate pNP-Xyl
(data not shown). The β-xylosidase was not active on the three monoacetates of pNP-Xyl. In
the XloA-coupled assay, TM0077 hydrolyzed acetate from positions 2, 3 and 4 of pNP-Xyl
with similar catalytic efficiency. The results are summarized in Table II.
In addition, TM0077 was investigated for its ability to remove acetyl groups from 7-
aminocephalosporanic acid (7-ACA), cephalosporin C, glucose penta-acetate, N-acetyl-D-
glucosamine, xylan and acetylated xylan. TM0077 has no activity for acetylated and non-
acetylated xylan polymers, indicating that it is, indeed, an acetyl esterase and not an acetyl
xylan esterase. As expected for an acetyl esterase, TM0077 displayed high activity on
glucose penta-acetate with a turnover number of 2680 s−1. Like other members of CE7,
TM0077 was also able to hydrolyze the acetyl groups from both cephalosporin C and 7-
ACA with a turnover number of 376 s−1 and 1140 s−1, respectively. TM0077 was not able to
hydrolyze the acetyl group from N-acetyl-D-glucosamine, indicating that it is specific for
ester bonds and unable to hydrolyze amide bonds.
Inhibitor assays and TM0077 structures complexed with PMSF and paraoxon
PMSF and paraoxon [Fig. 1(d,e)] are competitive irreversible inhibitors of esterases.
Inhibition proceeds by the formation of a reversible Michaelis complex, followed by an
irreversible step and inhibition can, therefore, be characterized by two parameters: a
dissociation constant and a binding rate constant. The inhibition kinetics for paraoxon and
PMSF were investigated in the presence of p-nitrophenyl acetate, as described previously 47,
and the dissociation and rate constants were 0.5 ± 0.1 mM and 0.13 ± 0.02 s−1 for paraoxon,
and 1.1 ± 0.2 mM and 0.020 ± 0.001 s−1 for PMSF, respectively. The acetyl xylan esterase
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from Bacillus pumilus (BpAxe) has slightly reduced sensitivity to paraoxon (dissociation
and rate constants of respectively 5.4 mM and 0.012 s−1), likely due to steric hindrance of
two tyrosine residues (Tyr91 and Tyr206) that hamper the binding of paraoxon. Although
these residues are essentially conserved in TM0077 (Tyr92 and Phe213), TM0077 is more
sensitive to paraoxon than BpAxe48. In comparison to EST2 of Alicyclobacillus
acidocaldarius 49 and EstA of T. maritima 47, the TM0077 dissociation constant is slightly
higher, but the rate constant is comparable. No significant stimulation or reduction of
activity of TM0077 was observed in the presence of divalent metal ions or
ethylenediaminetetraacetic acid (EDTA).
To obtain more information about inhibitor binding and any possible conformational
changes during catalysis, TM0077 was co-crystallized with the inhibitors PMSF and
paraoxon and the PMSF (TM0077-PMS) and paraoxon (TM0077-DEP) structures were
determined to 2.4 Å and 2.1 Å, respectively (Table I). The electron density map of TM0077
with PMSF showed clear density for the PMSF covalent modification. The fluorine was
cleaved from the PMSF molecule during the binding reaction and the phenylmethyl sulfonyl
(PMS) moiety is covalently bound to the Oγ atom of Ser188. The native apo and PMS-
bound structures superimpose well with RMSD’s of 0.09–0.11 Å over 320–321 Cα atoms.
Electron density maps of the paraoxon-bound structure displayed clear density for a diethyl-
phosphate moiety covalently bound to the Oγ atom of Ser188. This covalent modification
indicates that the p-nitrophenol group of paraoxon was cleaved off during co-crystallization,
and a tetrahedral product reminiscent of the first transition state was formed during carboxyl
ester hydrolysis. The native apo and paraoxon-bound structures superimpose with RMSD’s
of 0.12–0.32 Å over 320–322 Cα atoms. Attempts to obtain co-crystals of TM0077 with
cephalosporin C, even at a low temperature of 4°C, were unsuccessful.
Analysis of the active site
TM0077 has a classic catalytic triad, consisting of Ser188 as the nucleophile, His303 as the
proton acceptor/donor, and Asp274 as the acidic residue stabilizing the histidine (Fig. 6).
The catalytic serine Ser188 is located within a conserved pentapeptide sequence, Gly-X-Ser-
X-Gly (GGSQG), characteristic of esterases and lipases. The positions of Ser188, Asp274,
and His303 are consistent with their expected locations in the canonical fold of the α/β-
hydrolase family. Ser188 is located at the nucleophile elbow in a sharp turn between β5 and
helix αC. The presence of three glycine residues (Gly186, Gly187, and Gly190) in close
proximity to Ser188 prevents steric hindrance and facilitates access to the nucleophile
elbow. Asp274 and His303 are located in loops between β7 and helix αE, and between β8
and helix αF, respectively. The oxyanion hole is formed by the backbone amide groups of
Tyr92 and Gln189. The catalytic triad and oxyanion hole are located in a depression on the
surface of TM0077. This ellipsoid pocket (S1), which is approximately 12 Å wide, extends
15 Å from the catalytic serine. A smaller pocket (S2), approximately 5 Å long, extends to
the other side of the catalytic serine [Fig. 6(a)]. The volume of both pockets combined (S1 +
S2) is 1082 Å3 (CASTp analysis; 50). The substrate-binding pocket is bordered by residues
from helices αA and αF, and its base is formed by residues from β-strands 4, 5, 6, and their
adjacent C-terminal loops. The overall pocket is hydrophobic, although it does have some
polar residues (Gln88, Asp210, and Gln314), which may interact with the substrate.
In the native apo structure, the Ser188 hydroxyl makes a hydrogen bond with the imidazole
of His303 [Fig. 6(b)]. Extra density was observed near the side chain of Ser188 and was
interpreted as a chloride ion based on electron density size and shape as well as the
geometry of the interactions with surrounding residues. This chloride ion is bound at the
entrance of the oxyanion hole, forming hydrogen bonds with the backbone amides of Tyr92
and Gln189. In the PMSF-bound structure, the phenyl ring of the inhibitor is located in the
small active site groove surrounded by hydrophobic residues Tyr92, Trp124, Pro228, Ile276,
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and His303 [Fig. 6(c)]. The sulfonyl group of PMSF makes hydrogen bonds with the
backbone amides of Tyr92 and Gln189. In the paraoxon-bound structure, the diethyl-
phosphate (DEP) moiety is stabilized by hydrogen-bonding interactions with the oxyanion
hole. One of the two ethyl arms of bound paraoxon points toward the larger pocket in the
protein, while the other follows the groove of the small pocket. The two ethyl arms are
stabilized by packing against Tyr92, Trp124, Pro228, Ile276, and His303 [Fig. 6(d)].
Two rotamers of the catalytic serine
Although no large conformational changes were observed upon binding of PMSF or
paraoxon, a different rotamer of the catalytic serine side chain was observed compared to
native TM0077 [Fig. 7(a,b)]. Similar changes have been observed in several other esterases
and have been shown to play a key role in the catalytic mechanism (see CONCLUSION for
more details). In the native structure, the catalytic Ser188 Oγ is in the plane of the imidazole
ring of His303, as most commonly observed in the resting state of esterases and lipases 51.
The Ser188 Oγ forms a hydrogen bond (2.6 Å) with His303 Nε2. In the PMSF- and
paraoxon-bound structures, the conformation of the catalytic serine changes; the Ser188 Oγ
rotates about 110°, increasing the distance (3.1 Å and 2.8 Å for PMSF and paraoxon bound
structures, respectively) to the His303 imidazole ring. In the TM0077-SeMet structure, the
catalytic serine is also rotated over ~110°, with a distance to the imidazole ring of 3.0 Å
[Fig. 7(c.)]. A probable explanation for this observation could be the protonation of His303,
since TM0077-SeMet was crystallized at a lower pH (pH 4.2) compared to the native
TM0077 (pH 7.3). Furthermore, extra electron density was identified in the TM0077-SeMet
structure, suggesting a partially occupied acyl intermediate on Ser188. However, as this
density is not sufficiently clear and interpretable to fit an acyl intermediate, water molecules
were modeled instead. No rearrangements of any other residues in the active site were
observed.
CONCLUSION
TM0077 from the hyperthermophilic bacterium T. maritima was predicted from its gene
sequence to be an acetyl xylan esterase. We have expressed and purified TM0077 and
experimentally demonstrated that it has ester-hydrolyzing activity. The TM0077 activity was
restricted to short acyl chain esters (C2 and C3) when artificial p-nitrophenyl-esters were
used as substrates. In addition, the enzyme has high specific activity on glucose penta-
acetate. However, no activity was detected on xylan or acetylated xylan. Thus, TM0077
should be reclassified as an acetyl esterase, and not as an acetyl xylan esterase as currently
annotated 52. Furthermore, the lack of any apparent signal sequence suggests that the protein
is not secreted. Thus, the predicted intracellular location of TM0077 is compatible with a
role other than the deacetylation of extracellular xylan. Based on these results, we conclude
that the likely biological function of TM0077 is removal of the remaining acetyl groups
from the short, end products of xylan degradation that are imported into the cytoplasm. The
resulting deacetylated xylose oligomers are the substrates for a β-xylosidase. This role for
TM0077 is in good agreement with the clustering of the TM0077 gene with other genes
involved in xylose metabolism. However, it cannot be ruled out that TM0077 may also act
on other small, acetylated compounds.
TM0077 is the first esterase from the CE7 family to be tested for its positional specificity for
the deacetylation of 4-nitrophenyl-β-D-xylopyranoside. TM0077 hydrolyzes acetate at the 2,
3 and 4 positions of 4-nitrophenyl-β-D-xylopyranoside with similar efficiency. Conversely,
the CtAxe esterase from Clostridium thermocellum in the CE4 family shows a clear
preference for hydrolyzing acetate at the 2 position 53, and Penicillium purpurogenum AXE
II esterase, a member of the CE5 family, also has a preference for acetate at position 2 54.
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This lack of preference for a specific position of the acetate group correlates with the
relative broad substrate specificity of the CE7 esterases.
Esterases and deacetylases in the CE7 family are unusual in that they are active towards both
acetylated xylo-oligosaccharides and the antibiotic cephalosporin C [Fig. 1(a,b)]. Therefore,
TM0077 was investigated for activity towards the substrates 7-ACA and cephalosporin C.
The activity of TM0077 on these substrates is approximately ten-fold higher than that of the
acetyl xylan esterase from B. pumilus 55 or the acetyl esterase from Thermoanaerobacterium
sp. strain JW/SL YS485 56. TM0077 has a higher hydrolytic activity on 7-ACA compared to
cephalosporin C, as described for other CE7 esterases 40,55,56. Nonetheless, it is unlikely that
both compounds are natural substrates, because the stability of these compounds at the
optimal growth temperature (80°C) of T. maritima is very low.
Crystal structures of TM0077 in complex with inhibitors PMSF and paraoxon revealed that,
upon binding of PMSF or paraoxon, the reaction is trapped at the acylation step via the
formation of a covalent tetrahedral reaction product. In the complexed structures, the
negatively charged oxygen of the tetrahedral intermediate, derived from the substrate
oxyanion, is stabilized by hydrogen bonds with the backbone amide groups of Tyr92 and
Gln189. Comparison of the TM0077 complexed structures with the native structure shows
that the catalytic serine (Ser188) Oγ rotates about 110°, thereby increasing the distance
between Ser188 Oγ and His303 Nε2. Such a conformational change of the catalytic serine
has been observed in several other esterases, including Fusarium solani cutinase 57,
Penicillium purpurogenum acetyl xylan esterase 51, Rhodococcus sp. strain MB1 cocaine
esterase 58, Bacillus subtilis lipase 59, Rhodococcus sp. strain H1 heroine esterase 60, and
Aspergillus niger feruloyl esterase 61. The classical model for the catalytic mechanism of
esterases consists of a sequential two-step hydrolysis. The first reaction involves
nucleophilic attack by the catalytic serine on the substrate carbonyl carbon, resulting in an
acyl-enzyme and the liberation of an alcohol. In the second reaction, a water molecule
performs a nucleophilic attack on the acyl-enzyme, the acyl-enzyme bond breaks and the
carboxylate is released 62. Although the catalytic mechanism of esterases is well established,
it is unclear why the initially generated tetrahedral intermediate does not collapse back to the
reactant complex during the nucleophilic attack of the substrate. A previously proposed
mechanism that would prevent this collapse is the spatial reorganization of the catalytic
residues during the initial catalytic step, causing the residues to separate and thereby drive
the reaction forward 62–64. The apo and inhibitor bound structures of TM0077, presented
herein, support this proposed mechanism. Moreover, in a recent study of the serine protease
mechanism, it was suggested that subtle atomic motions of the catalytic serine and histidine
residues during the catalytic cycle favor the forward reaction 65. Thus, rotation of Ser188 Oγ
of TM0077 may be required to inhibit reversal of the reaction. In addition, such changes
may facilitate the access of water to the catalytic histidine so that the second step of the
reaction can go to completion.
Deacetyl cephalosporins are valuable building blocks for the production of semisynthetic β-
lactam antibiotics. These compounds are derived from cephalosporin C or 7-
aminocephalosporanic acid via enzymatic or chemical processes 10. The thermostable
TM0077 esterase may be valuable in the preparation of derivatives of β-lactam antibiotics.
Recently, the substrate specificity of the acetyl xylan esterase from P. purpurogenum was
engineered to accept a range of fatty acid esters of up to 14 carbons compared to its wild-
type preference for acetate54. It might also be possible to engineer TM0077 and enable the
(de)acetylation of cephalosporins at the C10 position with various acyl chains. Because of
its high stability and activity on 7-ACA and cephalosporin C, TM0077 presents an attractive
candidate for the production of new semi-synthetic antibiotics.
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Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We gratefully acknowledge contributions from George Sheldrick for modifications of the SHELXD program, and
for Global Phasing Ltd. that made significant improvements in the automation of autoSHARP. We also thank
Victor Lamzin for updates of chain docking of the ARP/wARP program, and Gerard Bricogne and Eleanor Dodson
for helpful discussion on phasing for the large TM0077-SeMet structure, and Willem J. van Berkel for valuable
discussion on the catalytic mechanism of TM0077. Portions of this research were carried out at the Stanford
Synchrotron Radiation Lightsource (SSRL) and the Advanced Light Source (ALS). The SSRL is a Directorate of
SLAC National Accelerator Laboratory and an Office of Science User Facility operated for the U.S. Department of
Energy Office of Science by Stanford University. The SSRL Structural Molecular Biology Program is supported by
the DOE Office of Biological and Environmental Research, and by the National Institutes of Health, National
Center for Research Resources, Biomedical Technology Program (P41RR001209), and the National Institute of
General Medical Sciences. The ALS is supported by the Director, Office of Science, Office of Basic Energy
Sciences, Materials Sciences Division, of the U.S. Department of Energy under Contract No. DE-
AC02-05CH11231 at Lawrence Berkeley National Laboratory. Genomic DNA from Thermotoga maritima MSB8
(DSM3109) (ATCC #43589D-5) was obtained from the American Type Culture Collection (ATCC). The content is
solely the responsibility of the authors and does not necessarily represent the official views of the National Institute
of General Medical Sciences or the National Institutes of Health.
Grant sponsor: NIH Grant numbers U54 GM094586 and U54 GM074898 (Protein Structure Initiative); Grant
sponsor: The Graduate School VLAG Wageningen, the Netherlands (ML).
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Figure 1.
Substrates and inhibitors of the CE7 family of enzymes. Structures of (A) acetylated
xylooligosaccharide, (B) cephalosporin C, (C) p-nitrophenyl-acetate, (D)
phenylmethylsulfonyl fluoride (PMSF), and (E) paraoxon.
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Figure 2.
Overall fold and topology of TM0077. (A) Stereo view of a TM0077 protomer. The β-
strands are labeled numerically (-1 to 8) with the core strands in red, α-helices are labeled
alphabetically (A-2 to F) and 310-helices are labeled with an Eta (η1 and η2) with the core
helices in cyan. The three-helix insertion after β6 is colored green and the N-terminal
extension is colored sky blue. The figure was generated using Pymol 31. (B) Topology
diagram of TM0077, with the helices displayed as cylinders and the strands displayed as
arrows following the color and label scheme of (A). The location of residues forming the
catalytic triad is also indicated.
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Figure 3.
Structural superposition of TM0077 with structurally related esterases. Superposition of
TM0077 (yellow) with (A) the cephalosporin C deacetylase (CAH) from B. subtilis (PDB:
1ods; blue) 40 and (B) the α/β-hydrolase domain of the acylpeptide hydrolase/esterase
apAPH from A. pernix K1 (PDB: 1ve6; grey) 43.
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Figure 4.
TM0077 oligomeric assembly. (A) Surface representation of the biological unit of the
TM0077-Native hexamer with each monomer in a different color (left). The “cross section”
shows the entrances on either side of the assembly and the internal cavity (center), and a 90°
rotated view of the TM0077-Native hexamer, with a close-up view of the open central hole
(right). (B) Surface representation of the biological hexamer unit of CAH from B. subtilis 40
(left) and the TM0077-SeMet hexamer with a close-up view of the blocked central hole
(right).
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Figure 5.
Effect of temperature and pH on esterase activity. (A) The esterase activity was studied
using pNP-C2 as a substrate at temperatures ranging from 40–100°C. The inset shows the
temperature dependence as an Arrhenius plot. (B) Thermal stability of TM0077 at 90°C. (C)
The effect of pH on esterase activity studied using pNP-C2 as a substrate at pH values in the
range of 4.8–9.2.
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Figure 6.
TM0077 catalytic site. (A) Surface representation of the TM0077 catalytic site, with His303,
Asp274 and the intermediate DEP-modified Ser188 shown as sticks. The two binding
pockets are indicated with S1 and S2. (B) Apo TM0077 with a bound chloride ion (green
sphere), (C) TM0077 with PMS-modified Ser188 and (D) TM0077 with DEP-modified
Ser188. The catalytic residues are shown as sticks, with the hydrogen bonds shown as
dashed lines. Carbon atoms are in green (apo), cyan (PMS) or blue (DEP), oxygen atoms in
red, sulfur atoms in yellow and phosphate in orange. Electron density omit maps shown for
inhibitor modified Ser188 contoured at 1σ show that the PMS and DEP are covalently
bonded to Ser188 in (C) and (D), respectively. Distances are shown in Ångströms.
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Figure 7.
Conformational change of Ser188 Oγ. The Oγ atom of the Ser188 is rotated ~110° between
the native apo structure (cyan) and (A) the complexed PMS-modified Ser188 structure
(pink), (B) the DEP-modified Ser188 structure (light blue) and (C) the SeMet structure
(purple). The different hydrogen bonds made for the Ser Oγ in the native versus complexed
structures are shown as dashed black lines with distances in Ångströms.
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Table I
Summary of crystal parameters, data collection, and refinement statistics
TM0077-SeMet
TM0077-Native
TM0077-PMS
TM0077-DEP
Space group
P 21
P 21 21 21
P 21 21 21
P 21 21 21
Unit cell parameters
a=152.64Å b=130.95Å
c=157.82Å β=118.90°
a=103.46Å
b=103.79Å
c=221.02Å
a=103.57Å
b=104.50Å
c=221.61Å
a=103.80Å
b=104.43Å
c=221.64Å
Data collection
λ1 MAD-Se
λ2 MAD-Se
Wavelength (Å)
0.9791
0.9183
0.9765
0.9765
0.9765
Resolution range (Å)
29.6 – 2.10
29.6 – 2.10
48.8 – 2.50
49.0 – 2.40
49.0 – 2.12
No. observations
1,119,236
1,100,249
1,222,016
765,546
989,949
No. unique reflections
293,140
291,757
83,045
94,681
123,070
Completeness (%)
93.0 (61.8)a
92.6 (60.8)
100 (100)
100 (100)
89.8 (53.5)
Mean I/σ(I)
9.1 (2.4)a
9.6 (2.2)
14.4 (2.9)
11.5 (3.4)
15.3 (2.2)
Rmerge on I (%)
12.3 (52.5) a
11.9 (57.9)
20.7 (109.7) c
18.0 (67.4)
9.5 (51.9)
Rmeas on I (%)
14.3 (62.2) a
13.9 (68.7)
21.4 (113.6)
19.2 (71.9)
10.2 (60.2)
Rpim on I (%)
7.2 (32.7) a
7.1 (36.2)
5.5 (29.2)
6.7 (24.9)
3.5 (29.2)
Highest resolution shell (Å)
2.15 – 2.10
2.15 – 2.10
2.56 – 2.50
2.46 – 2.40
2.18 – 2.12
Model and refinement statistics
Resolution range (Å)
29.6 – 2.10
48.8 – 2.50
49.0–2.40
49.0 – 2.12
No. reflections (total)
293,097 b
83,045
94,680
122,994
No. reflections (test)
14,726
4,200
4,742
6,188
Completeness (% total)
92.8
100.0
100.0
89.8
Data set used in refinement
λ1 MAD-Se
Cutoff criteria
|F| > 0
|F| > 0
|F| > 0
|F| > 0
Rcryst
0.186
0.167
0.160
0.167
Rfree
0.223
0.212
0.208
0.205
Stereochemical parameters
Restraints (RMSD observed)
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TM0077-SeMet
TM0077-Native
TM0077-PMS
TM0077-DEP
Bond angle (°)
1.48
1.47
1.53
1.44
Bond length (Å)
0.018
0.017
0.017
0.015
Av. isotropic B-value (Å2)
27.9
24.7
19.4
19.6
ESU based on Rfree
0.17
0.25
0.22
0.18
Water molecules/other solvent molecules
2,464/1
507/24
946/17
987/23
PDB ID
1vlq
3m81
3m82
3m83
aHighest resolution shell
ESU = Estimated overall coordinate error 16,66.
Rmerge=ΣhklΣi|Ii(hkl)-<I(hkl)>|/ΣhklΣiIi(hkl), Rmeas(redundancy-independent Rmerge)=Σhkl[Nhkl/(Nhkl-1)]1/2Σi|Ii(hkl)-<I(hkl)>|/ΣhklΣiIi(hkl), and Rpim(precision-indicating Rmerge)=Σhkl[1/
(Nhkl-1)]1/2Σi|Ii(hkl)-<I(hkl)>|/ΣhklΣiIi(hkl) 67–69.
Rcryst = Σ| |Fobs|-|Fcalc| |/Σ|Fobs| where Fcalc and Fobs are the calculated and observed structure factor amplitudes, respectively.
Rfree = as for Rcryst, but for 5.0 % of the total reflections chosen at random and omitted from refinement.
bTypically, the number of unique reflections used in refinement is slightly less than the total number that were integrated and scaled. Reflections are excluded due to systematic absences, negative
intensities, and rounding errors in the resolution limits and cell parameters.
cRmerge of the highest resolution shell is high due to high redundancy (14.7). However, the completeness and mean I/σ of the highest resolution shell are reasonable, and these data were included in the
refinement.
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Table II
Kinetic parameters for hydrolysis of various esters
Ester
Km (mM)
kcat (s−1)
kcat/Km (s−1 mM−1)
pNP-Acetate
0.185 ± 0.026
57.5 ± 2.2
310.8 ± 45.3
pNP-Propionate
0.137 ± 0.013
41.3 ± 1.1
301.5 ± 29.7
2-O-acetyl pNP-Xyl
3.6 ± 0.5
76.1 ± 19.2
21.1 ± 6.1
3-O-acetyl pNP-Xyl
4.2 ± 0.4
70.1 ± 7.7
16.7 ± 2.4
4-O-acetyl pNP-Xyl
4.0 ± 0.1
78.6 ± 12.9
19.7 ± 3.3
Proteins. Author manuscript; available in PMC 2013 June 01.
|
3M83
|
Crystal structure of Acetyl xylan esterase (TM0077) from THERMOTOGA MARITIMA at 2.12 A resolution (paraoxon inhibitor complex structure)
|
Functional and structural characterization of a thermostable
acetyl esterase from Thermotoga maritima
Mark Levisson1,*, Gye Won Han2,3,*, Marc C. Deller2,3, Qingping Xu2,4, Peter Biely5, Sjon
Hendriks1, Lynn F. Ten Eyck6,7, Claus Flensburg8, Pietro Roversi8, Mitchell D. Miller2,4,
Daniel McMullan9, Frank von Delft2,3,‡, Andreas Kreusch10, Ashley M. Deacon2,4, John van
der Oost1, Scott A. Lesley2,3,10, Marc-André Elsliger2,3, Servé W. M. Kengen1,†, and Ian A.
Wilson2,3,†
1Laboratory of Microbiology, Wageningen University, 6703 HB, Wageningen, The Netherlands
2Joint Center for Structural Genomics, http://www.jcsg.org 3Department of Molecular Biology, The
Scripps Research Institute, La Jolla, California 92037 4Stanford Synchrotron Radiation
Lightsource, SLAC National Accelerator Laboratory, Stanford University, Menlo Park, California
92045 5Institute of Chemistry, Slovak Academy of Sciences, 845 38 Bratislava, Slovakia
6Department of Chemistry and Biochemistry, University of California at San Diego, La Jolla,
California 92093-0505 7San Diego Supercomputer Center, University of California at San Diego,
La Jolla, California 92093-0505 8Global Phasing Ltd. Sheraton House, Castle Park, Cambridge
CB3 0AX, United Kingdom 9Protein Therapeutics Department, Genomics Institute of the Novartis
Research Foundation, San Diego, California 92121 10Protein Sciences Department, Genomics
Institute of the Novartis Research Foundation, San Diego, California 92121
Abstract
TM0077 from Thermotoga maritima is a member of the carbohydrate esterase family 7 and is
active on a variety of acetylated compounds, including cephalosporin C. TM0077 esterase activity
is confined to short-chain acyl esters (C2-C3), and is optimal around 100°C and pH 7.5. The
positional specificity of TM0077 was investigated using 4-nitrophenyl-β-D-xylopyranoside
monoacetates as substrates in a β-xylosidase-coupled assay. TM0077 hydrolyzes acetate at
positions 2, 3 and 4 with equal efficiency. No activity was detected on xylan or acetylated xylan,
which implies that TM0077 is an acetyl esterase and not an acetyl xylan esterase as currently
annotated. Selenomethionine-substituted and native structures of TM0077 were determined at 2.1
Å and 2.5 Å resolution, respectively, revealing a classic α/β-hydrolase fold. TM0077 assembles
into a doughnut-shaped hexamer with small tunnels on either side leading to an inner cavity,
which contains the six catalytic centers. Structures of TM0077 with covalently bound
phenylmethylsulfonyl fluoride (PMSF) and paraoxon were determined to 2.4 Å and 2.1 Å,
respectively, and confirmed that both inhibitors bind covalently to the catalytic serine (Ser188).
Upon binding of inhibitor, the catalytic serine adopts an altered conformation, as observed in other
esterase and lipases, and supports a previously proposed catalytic mechanism in which this Ser
hydroxyl rotation prevents reversal of the reaction and allows access of a water molecule for
completion of the reaction.
†Correspondence to: Ian A. Wilson, Ph.D., Department of Molecular Biology, The Scripps Research Institute, La Jolla, California
92037; (858) 784-2939 Fax: (858) 784-2980; wilson@scripps.edu or Servé W. M. Kengen, Ph.D., Laboratory of Microbiology,
Wageningen University, 6703 HB, Wageningen, The Netherlands; 31 317 483737, Fax: 31 317-483829; serve.kengen@wur.nl.
*ML and GWH contributed equally to this work.
‡Current address: The Structural Genomics Consortium, Roosevelt Drive, Headington, Oxford OX3 7DQ, UK
NIH Public Access
Author Manuscript
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Published in final edited form as:
Proteins. 2012 June ; 80(6): 1545–1559. doi:10.1002/prot.24041.
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Keywords
Acetyl esterase; Thermotoga maritima; crystal structure; α/β hydrolase; inhibitor; serine rotation
INTRODUCTION
Thermotoga maritima is a hyperthermophilic bacterium that grows optimally at 80°C and is
able to metabolize a variety of simple and complex carbohydrates, including glucose,
sucrose, starch, cellulose, and xylan 1. Its carbohydrate utilization potential was confirmed
by analysis of its sequenced genome 2. The xylan degrading pathway of T. maritima has
been studied using microarrays 2–4, and several genes encoding transporters, xylanases, and
a β-xylosidase have been identified. Among the enzymes with a differential expression
pattern in the microarray was a predicted acetyl xylan esterase (locus tag TM0077,
axeA) 3,5. Depending on the source, the xylan backbone may contain a varying degree of
acetylated xylose residues. Therefore, in addition to xylanases and xylosidases, the complete
degradation of xylan requires esterases/deacetylases 6.
Presently, esterases and deacetylases that are active on carbohydrate substrates have been
classified into 16 families by Henrissat and coworkers (Carbohydrate-Active enZymes
Server (CAZy)) 7. According to this classification, the predicted acetyl xylan esterase from
T. maritima would be a member of family 7 of the carbohydrate esterases (CE7). In addition
to the acetyl xylan esterase activity, enzymes in the CE7 family are rather unusual in that
they display a high specific activity towards the antibiotic cephalosporin C [(Fig. 1(a-b)] 8.
Cephalosporins belong to the β-lactam class of antibiotics, which also includes penicillin,
and affect bacterial cell growth by inhibiting the penicillin-binding-protein that cross-links
peptide glycans required for cell wall formation 9. The production of deacetylated
cephalosporins is of great interest because these compounds are valuable building blocks for
the production of semi-synthetic β-lactam antibiotics10,11.
To explore the catalytic capacity of the predicted acetyl xylan esterase from T. maritima and
gain a better insight into the structure and function of the family 7 carbohydrate esterases,
TM0077 was expressed and purified, and three-dimensional structures of the native enzyme
and its complexes with phenylmethylsulfonyl fluoride (PMSF) and paraoxon inhibitors,
were determined by x-ray crystallography. Furthermore, the enzyme was functionally
characterized, and various biochemical properties including the positional specificity of the
esterase were investigated.
MATERIALS AND METHODS
Gene cloning
TM0077 was selected as part of the Joint Center for Structural Genomics (JCSG) effort on
complete structural coverage of the T. maritima soluble proteome as a large-scale center for
high-throughput structure determination funded under the NIHGMS Protein Structure
Initiative (PSI) 12. The gene encoding TM0077 (GenBank: AAD35171.1, GI:4980565;
SwissProt: Q9WXT2) was amplified by polymerase chain reaction (PCR) from genomic
DNA using PfuTurbo DNA polymerase (Stratagene) and primers corresponding to the
predicted 5′ and 3′ ends. The PCR product was cloned into plasmid pMH1, which encodes
an expression and purification tag (MGSDKIHHHHHH) at the amino terminus of the
protein. The cloning junctions were confirmed by DNA sequencing.
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TM0077-SeMet protein production and purification
Protein production was performed in a selenomethionine-containing medium using the
Escherichia coli methionine auxotrophic strain DL41. Expression was induced by the
addition of 0.15% L-arabinose. At the end of fermentation, cells were harvested and
subjected to one freeze/thaw cycle, and subsequently sonicated in Lysis Buffer [50 mM Tris
pH 7.9, 50 mM NaCl, 1 mM MgCl2, 0.25 mM Tris(2-carboxyethyl)phosphine hydrochloride
(TCEP), 1 mg/ml lysozyme] and the lysate was centrifuged at 3,400 × g for one hour. The
soluble fraction was applied to nickel-chelating resin (GE Healthcare) pre-equilibrated with
Equilibration Buffer [50 mM potassium phosphate pH 7.8, 300 mM NaCl, 10% (v/v)
glycerol, 0.25 mM TCEP] containing 20 mM imidazole. The resin was washed with
Equilibration Buffer containing 40 mM imidazole, and the protein was eluted with Elution
Buffer [20 mM Tris pH 7.9, 300 mM imidazole, 10% (v/v) glycerol, 0.25 mM TCEP]. The
eluate was buffer exchanged with Buffer Q [20 mM Tris pH 7.9, 5% (v/v) glycerol, 0.25
mM TCEP] containing 50 mM NaCl and applied to a RESOURCE Q column (GE
Healthcare) pre-equilibrated with the same buffer. The protein was eluted using a linear
gradient of 50 to 500 mM NaCl in Buffer Q and purified further with a HiLoad 16/60
Superdex 200 column (GE Healthcare), using Crystallization Buffer [20 mM Tris pH 7.9,
150 mM NaCl, 0.25 mM TCEP] as the mobile phase. For crystallization trials, the peak
Superdex 200 fractions were concentrated to ~15 mg/mL by centrifugal ultrafiltration
(Millipore). Molecular weight and oligomeric state of TM0077 were determined using a 1
cm × 30 cm Superdex 200 column (GE Healthcare) coupled with miniDAWN static light
scattering (SEC/SLS) and Optilab differential refractive index detectors (Wyatt
Technology). The mobile phase consisted of 20 mM Tris pH 8.0, 150 mM NaCl, and 0.02%
(w/v) sodium azide.
Native TM0077 production and purification
For protein production, E. coli DL41 cells were grown in LB medium for 8 hours (an
OD600 well above 2.0 was reached). Subsequently, the culture was induced by adding
0.15% L-arabinose and incubated another 16 hours at 37°C. Cells were harvested by
centrifugation at 10,000 × g for 20 min. The cell pellet was resuspended in 30 ml of Lysis
Buffer 2 [50 mM Tris-HCl pH 8.0, 50 mM NaCl, 10 mM imidazole, 0.25 mM TCEP]. The
cells were disrupted by two passages through a French press at 110 MPa. The crude cell
extract was treated with DNAse I at room temperature for 30 min and subsequently
centrifuged at 43,000 × g for 30 min in order to remove cell debris. The supernatant was
heated at 70°C for 25 min and then centrifuged to remove the precipitated proteins. The
supernatant was filtered and loaded onto a nickel-chelating column packed with 20 ml of Ni-
NTA His-Bind Resin (Novagen) and equilibrated in 50 mM Tris-HCl pH 8.0, 300 mM
NaCl, 2% (v/v) glycerol, and 0.25 mM TCEP. The column was washed with 20 mM
imidazole in the same buffer, and proteins were subsequently eluted with a linear gradient of
20–500 mM imidazole in the same buffer. Fractions containing esterase activity were pooled
and loaded onto a HiPrep Desalting column (GE Healthcare) equilibrated with 20 mM Tris-
HCl pH 8.0, 150 mM NaCl, and 0.25 mM TCEP. The homogeneity of the protein was
checked by SDS-PAGE, and activity staining of the SDS-PAGE gel was performed using α-
napthyl acetate, as described previously 13. The protein concentration was determined at 280
nm using a NanoDrop ND-1000 Spectrophotometer.
Crystallization
Crystals of selenomethionine-substituted TM0077 were obtained by hanging drop vapor
diffusion against a 250 μl crystallization solution consisting of 20% (w/v) PEG-3000, 0.1 M
HEPES pH 7.5, 0.2 M NaCl. Drops consisted of 0.5 μl protein and 0.5 μl crystallization
solution. Native TM0077 was crystallized using nanodrop vapor diffusion techniques
against a crystallization solution consisting of 0.2 M calcium acetate hydrate, 20% (w/v)
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PEG 3350, pH 7.3 at 20°C. Protein was concentrated to 22.8 mg/ml. Drops consisted of 100
nl protein and 100 nl of crystallization solution and a 60 μl reservoir of crystallization
solution. Crystals of TM0077 in complex with inhibitors PMSF or paraoxon were obtained
at 4°C in the same conditions with the same reagents as the native crystals. PMSF or
paraoxon were added in a molar ratio of 1:3 (protein:inhibitor).
Data collection
For cryoprotection, the TM0077-SeMet crystal was transferred to crystallization solution
supplemented with 15% (v/v) glycerol. The crystal was mounted in a cryoloop and
subsequently flash-cooled in liquid nitrogen. X-ray data were collected at 100 K on
beamline BL9-2 at the Stanford Synchrotron Radiation Lightsource (SSRL, Menlo Park,
CA) using a Quantum 4 CCD detector (ADSC). A TM0077-SeMet MAD data set was
collected to 2.1 Å resolution and the data were indexed in monoclinic space group P21, with
unit cell parameters a = 152.6 Å, b = 131.0 Å, and c = 157.8 Å, and β=118.9°, and 12
molecules in the asymmetric unit. Data were indexed and integrated with DENZO and then
scaled with SCALEPACK 14.
Native TM0077, TM0077-PMSF complex (TM0077-PMS) and TM0077-paraoxon complex
(TM0077-DEP) crystals were transferred to crystallization solution supplemented with 10%
(v/v) ethylene glycol and flash-cooled to 100K. Data were collected at beamline 5.0.3 of the
Advanced Light Source (ALS, Berkeley, CA) and processed with the HKL2000 package 14.
The native data set was collected to 2.5 Å resolution, and TM0077-PMS and TM0077-DEP
data sets were collected to 2.4 and 2.1 Å, respectively. All data were indexed in
orthorhombic space group P212121, with unit cell parameters approximately a=103Å
b=104Å c=221Å (See Table 1), and six molecules in the asymmetric unit. Data reduction
and refinement statistics for TM0077-SeMet, TM0077-Native, TM0077-PMS and TM0077-
DEP are summarized in Table I.
Structure solution and refinement
The TM0077-SeMet structure was solved by MAD phasing method using a two-wavelength
MAD dataset. At the time of the initial data collection (2001), the structure determination of
Se-MAD TM0077 posed a significant challenge to crystallographic programs, which were
still under active development. As a result, modifications were made in various structure
determination and refinement programs to achieve success. For initial phasing, SHELXD 15
was used to find candidate SeMet substructure sites. Attempts to complete phasing were
unsuccessful due to the translational non-crystallographic symmetry (NCS) (not recognized
initially). Self-consistent sets (partial sets) were found using the CCP4 program
PROFESSS 16 and additional SeMet sites were found by SHELXD, and added to these
partial sets. The SHARP 17 run did not complete initially; however, updates of SHARP and
ARP/wARP eventually helped to resolve issues and an initial trace was obtained by ARP/
wARP. The structure was then refined with BUSTER 18 using tight NCS restraints to an
Rcryst and Rfree of 18.6% and 22.3%, respectively. Model building was performed using
O 19 and the structure was refined using Refmac5 20. Refinement statistics are summarized
in Table I. The final model contains 12 protein molecules (chains A-L) in the asymmetric
unit each consisting of residues 2-323. The MolProbity 21 Ramachandran plot analysis
showed that 97.4% of all residues are in favored regions with a single outlier, Gln120 of
chain B, which is supported by unambiguous electron density. Ramachandran outlier
Gln120 of chain B of TM0077-SeMet is due to crystal packing with chain C. The backbone
carbonyl oxygens of Gln120 and Gly119 of chain B makes hydrogen bonds with the
backbone nitrogen of Gln140 of chain C (3.19 and 3.11 Å, respectively).
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The native TM0077 structure, TM0077-PMS and TM0077-DEP structures were solved by
molecular replacement using PHASER 22,23 with the TM0077-SeMet hexamer coordinates
(pdb: 1vlq; A-F chains) as a search model. One hexamer was successfully located and the
structure was further refined with Refmac5 20 using tight NCS restraints to an Rcryst and
Rfree of 16.7% and 21.2% (native TM0077), 16.0% and 20.8% (TM0077-PMS) and 16.7%
and 20.5% (TM0077-DEP), respectively. Iterative cycles of refinement and building were
performed with Refmac5, Phenix 24,25 and Coot 26. All other crystallographic manipulations
were carried out with the CCP4 package 16. Refinement statistics are summarized in Table I.
The final model of native TM0077 contains residues 3-324 (chains A, B, C, D and F) and
3-323 (chain E) in the asymmetric unit. Analysis of main-chain torsion angles using
MolProbity 21 showed that 97.8% of the residues are in favored regions of the
Ramachandran plot with 0.2% outliers (Asn302 of chains B, C and D), which are supported
by unambiguous electron density. The final model of TM0077-PMS contains residues 3-323
for all chains in the asymmetric unit with 97.5% of the residues in favored regions with
0.2% outliers (Asn302 of chains A, B, D and F). The final model of TM0077-DEP contains
residues 0-324 for (chains A, B, C and F) and 0-323 (chains D and E) in the asymmetric
unit, respectively, with 97.6% of the residues in favored region of the Ramachandran plot
with 0.2% outliers (Asn302 of B, C, D and F chains). Ramachandran outlier Asn302 in the
TM0077-Native, TM0077-PMS and TM0077-DEP structures is a neighbor to the catalytic
triad residue His303 and may reflect a slightly different state for these structures compared
to the Se-Met structure.
Structure validation and deposition
The quality of the crystal structure was analyzed using the JCSG Quality Control server
(http://smb.slac.stanford.edu/jcsg/QC). This server processes the coordinates and data
through a variety of validation tools including AutoDepInputTool 27 MolProbity 21,
WHATIF 5.0 28, RESOLVE 29, MOLEMAN2 30 as well as several in-house scripts, and
summarizes the results. Protein quaternary structure analysis were performed using the PISA
server 30. Figures were prepared with PyMOL (DeLano Scientific) 31. RMSD values were
calculated using the ProCKSI-Server 32. The structural data have been deposited in the
RCSB Protein Data Bank (PDB) with accession codes 1vlq for TM0077-SeMet, 3m81 for
TM0077-native, 3m83 for TM0077-DEP and 3m82 for TM0077-PMS.
Enzyme assays
Esterase activity was measured using p-nitrophenyl esters as described previously 13.
Briefly, the standard assay consisted of activity measurements with 0.2 mM p-nitrophenyl
acetate as substrate in 50 mM citrate-phosphate (pH 6) at 70°C. The p-nitrophenol liberated
was measured continuously at 405 nm on a Hitachi U-2001 spectrophotometer with a
temperature-controlled cuvette holder. Extinction coefficients of p-nitrophenol were
determined prior to each measurement. Kinetic parameters were determined by direct fitting
the data, obtained from multiple measurements, to the Michaelis–Menten curve (Tablecurve
2d, version 5.0).
The effect of pH on esterase activity was studied in the pH range from 5 to 10. The buffers
used were 50 mM citrate-phosphate (pH 5–8) and 50 mM CAPS (3-(cyclohexylamino) 1-
propanesulphonic acid) (pH 9.5–10). The pH of the buffers was set at room temperature, and
temperature corrections were made using their temperature coefficients: −0.0028 pH/°C for
citrate-phosphate buffer and −0.018 pH/°C for CAPS buffer. The effect of temperature on
esterase activity was studied in the range of 40–100°C using 0.2 mM p-nitrophenyl acetate
as substrate. Enzyme thermostability was determined by incubating the enzyme in a 50 mM
Tris-HCl, 150 mM NaCl (pH 7.8) buffer at 90°C and 100°C for various time intervals.
Residual activity was determined in the standard assay.
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Inhibition kinetics of PMSF and paraoxon were determined as described for the
acetylcholinesterase from electric eel 33. All experiments were performed at 70°C in 50 mM
citrate-phosphate (pH 6) buffer and 0.2 mM p-nitrophenyl acetate as substrate. The kinetic
constants for the inhibition of TM0077 with PMSF and paraoxon were measured in the
concentration range of 1.0–10.0 mM and 0.2–1.0 mM, respectively.
Deacetylase activity was determined using high-performance liquid chromatography
(HPLC) by measuring the amount of acetic acid released from the substrates cephalosporin
C, 7-aminocephalosporanic acid, glucose-pentaacetate and acetylated xylan. Xylan was
acetylated by the method described by Johnson 34. The reaction mixture contained 0.9 ml of
substrate solution (dissolved in 50 mM Tris-HCl, pH 7.5) and 0.1 ml of enzyme solution,
and was incubated at 37°C for various time intervals (0–10 min). The reaction was stopped
by adding 0.2 ml of stop solution (100 mN H2SO4 and 30 mM crotonate) and placing the
sample on ice. The conditions for HPLC were as follows: column, KC811 Shodex;
detection, RI and UV detectors; solvent, 3 mN H2SO4; flow rate, 1.5 ml/min; temperature,
30°C; internal standard, crotonate. One unit of enzyme activity was defined as the amount of
enzyme that releases one μmol of acetic acid per minute.
Activity on xylan was measured quantitatively using DMSO-extracted xylan (1%
polysaccharide solution in 0.1 M sodium phosphate buffer pH 6) at 60°C 35. Xylan will
precipitate as a consequence of deacetylation, resulting in a rapid turbidity of the solution.
Positional specificity assay
The positional specificity of TM0077 was investigated using an enzyme-coupled assay on
monoacetylated 4-nitrophenyl β-D-xylopyranosides (pNP-Xyl) as described 36. The β-
xylosidase XloA (locus tag: TM0076) from T. maritima was cloned into the vector pET24d
in frame with a C-terminal His6-tag. The enzyme was expressed and purified as described
above for native TM0077. Activity of XloA was confirmed by measuring the release of p-
nitrophenol at 405 nm from the substrate 4-nitrophenyl β-D-xylopyranoside.
The enzyme-coupled assay was performed at 60°C in a total volume of 125 μl, which
contained 0.1 M sodium phosphate (pH 6 or 7), 2-O-, 3-O-, or 4-O-acetyl pNP-Xyl, the β-
xylosidase XloA, and TM0077. Stable 50x-concentrated stock solutions of the substrates
were prepared in DMSO. The reaction was started by addition of 2.5 μl of a stock solution
to a preheated reaction mixture consisting of phosphate buffer, auxiliary β-xylosidase XloA
in excess, and TM0077. The reaction was terminated by addition of 800 μl of a 2% solution
of Na2CO3. Liberated p-nitrophenol was determined at 405 nm against substrate and
enzyme blanks. A short incubation time for activity determination was used to suppress
acetyl migration on the xylopyranosyl-ring, which is significant at pH 6 or 7 37. The kinetic
constants were determined at pH 7 and 60°C with reaction times of 2 or 5 minutes.
RESULTS and DISCUSSION
In silico analysis
TM0077 consists of 325 amino acids with a calculated molecular mass of 37 kDa. Sequence
analysis, using the SignalP 3.0 server, revealed that TM0077 has no predicted signal
sequence and is, therefore, believed to be an intracellular enzyme. Analysis of the gene
organization indicates that the TM0077 gene co-localizes with genes encoding a xylanase
(TM0070) 38, ABC transporter components (TM0071-TM0075), and a β-xylosidase
(TM0076) 39. BLAST-P analysis showed that TM0077 has highest similarity to putative
acetyl esterases, acetyl xylan esterases and cephalosporin C deacetylases. Among the
BLAST results, a predicted acetyl xylan esterase-related protein from T. maritima (locus
tag: TM0435) was also identified. TM0077 was compared with other members of the CE7
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family using structure-based, multi-sequence alignment and the putative catalytic triad,
Ser188, Asp274, and His303, was identified from conservation throughout the analyzed
sequences. The putative nucleophilic serine (Ser188) is located within a conserved
pentapeptide consensus sequence, Gly-Xaa-Ser-Gln-Gly, typical of this family. Previously, a
signature sequence motif, [RGQ]-(x:~70)-[GxSQG]-(x:~115)-[HE] (where x indicates any
amino acid), had been suggested for the CE7 family based on an aminoacid alignment of 12
sequences 40. In an updated alignment consisting now of 50 sequences, we observed many
sequences that have this signature motif, but it is not conserved throughout the entire family
(See Supporting Information and Fig. S1 for the multi-sequence alignment).
Overall structure
The crystal structure of seleno-methionine incorporated TM0077 (TM0077-SeMet) was
determined to 2.1 Å resolution by multi-wavelength anomalous dispersion (MAD) (Table I)
with twelve molecules per asymmetric unit. A native apo structure (TM0077-Native) was
determined in a different space group (see Methods) to 2.5 Å by molecular replacement,
using TM0077-SeMet as a search model, with six molecules in the asymmetric unit (Table
I). Each monomer of the native hexamer contained a calcium ion (see below) bound by
Lys22, Glu26, and Asp25 via a bridging water molecule. Superposition of the TM0077-
SeMet and the TM0077-Native structures gave a root-mean-square difference (RMSD) of
0.12 Å over 321 Cα atoms, which indicates that these structures are nearly identical as
expected.
In general, the TM0077 structure resembles the canonical α/β-hydrolase fold, which
consists of a central, twisted, eight-stranded β-sheet surrounded by α-helices on both sides,
with β2 antiparallel to the other strands. TM0077 deviates slightly from the canonical α/β-
hydrolase fold at two locations: a three-helix insertion after strand β6 and an extension of
the N-terminus (Fig. 2). Insertions after β6 or β7 are common for α/β-hydrolases and are
proposed to help shape the substrate-binding site 41. The N-terminus is extended by two
helices (αA-1 and αA-2) and an antiparallel β-strand (β-1) that aligns with the other eight β-
strands (β1-β8) and extends the central β-sheet. This nine-stranded β-sheet is highly twisted,
and β-1 and β8 at the extreme edges are rotated approximately 130° relative to each other.
Helices αA and αB both contain a short 310-helix segment at their N-terminus. Helices
αA-1, αA-2, αB, αC, αD, αD1, αD2, αD3, αE, and the 310-helix η2 are located on one side
of the central β-sheet, and helices αA, αF and the 310-helix η1 are on the other side.
A structural similarity search was performed using the program DALI 42. Monomer A of the
TM0077-SeMet structure was used as a search model and similarity was found with
cephalosporin C deacetylases, acetyl xylan esterases, acylamino-releasing enzymes,
dipeptidyl peptidases and some esterases and lipases. TM0077 is structurally most similar to
cephalosporin C deacetylase (CAH) from B. subtilis (PDB: 1ods) 40, acetyl xylan esterase
(AXE) from B. pumilus (PDB: 3fvr and 2xlb), acetyl xylan esterase (AXE1) from
Thermoanaerobacterium sp. JW/SL YS485 (PDB: 3fcy), and acylpeptide hydrolase/esterase
apAPH from Aeropyrum pernix K1 (PDB: 1ve6) 43. The sequence identity between
TM0077 and CAH is 41% and the two structures align with a Z-score of 46 and an RMSD
of 1.5 Å over 312 Cα atoms. The sequence identity with apAPH is 17% with a Z-score of
23.3 and an RMSD of 2.3 Å over 230 Cα atoms. Superpositions of TM0077 with CAH and
with apAPH are shown in Fig. 3.
Quaternary structure
The crystal structure of TM0077-SeMet contains two hexamers in the asymmetric unit that
are related by a non-crystallographic two-fold axis. Each hexamer contains a dimer of
trimers with a back-to-back arrangement (Fig. 4). The apo and the complex crystals
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contained one hexamer in the asymmetric unit. Crystallographic packing analysis using
PISA (EBI) 44 indicated that the relevant physiological oligomeric state of TM0077 is a
hexamer, which was confirmed by size exclusion chromatography coupled with static light
scattering. Further analyses of the hexameric assembly indicated that two main interfaces
play an essential role in complex formation. The first interface between subunit A and B
(green and cyan in Fig. 4) (identical to C/D and E/F) is stabilized by seven hydrogen bonds
on average and has a buried surface area of 1024 Å2 contributed by each chain. The second
interface between A and F (green and purple in Fig. 4) (B/C and D/E) is stabilized by 17
hydrogen bonds on average with a buried surface area of 1079 Å2 contributed by each chain
However, a multiple sequence alignment of TM0077 with other CE7 esterases showed that
the residues involved in these two main interfaces are not conserved. Other secondary
interfaces bury around 514 Å2 contributed by each chain. The hexamer has a total buried
surface area of 18,860 Å2, which is approximately 30% of the total surface area.
Approximately 3,143 Å2 per monomer is, therefore, buried upon complex formation.
The TM0077 hexamer has a doughnut-shape when viewed from the side, with the six active
sites located in the interior of the complex, where they line an oval-shaped cavity [Fig. 4(a)].
This cavity is accessible via two entrances, one on each side of the flat hexamer. Each of
these entrances is approximately 13 Å wide and connects to a short tunnel or pore spanning
approximately 10 Å to reach the inner cavity. Interestingly, in the TM0077-SeMet hexamer,
the entrance to the internal cavity is blocked by three phenylalanine residues (Phe4), one for
each of three monomers that compose half of the hexamer [Fig. 4(b)]. Residue Phe4 is
located in the mobile N-terminus (high B-values), which may indicate some flexibility or
multiple conformations.
Calcium ions were identified, by the electron density and coordination geometry, supported
by their presence in the crystallization reagents, in the native TM0077, TM0077-PMS and
TM0077-DEP structures, but not in the TM0077-SeMet structure. The SeMet protein was
crystallized without any calcium in the crystallization reagents. In each subunit of the
hexamer, one calcium ion is located at the N-terminal region of helix αA-1, and is
coordinated by the backbone carbonyl of Lys22 and the Glu26 carboxylate. The remainder
of the calcium coordination sphere is filled with waters from a neighboring solvent channel
present in all molecules in the asymmetric unit. The Asp25 carboxylate contributes to the
calcium binding via one of the coordinating water molecules. Another calcium ion is bound
in a crystal packing interface between chain A and chain C′ of a crystallographic symmetry-
related hexamer. This calcium is coordinated by the carboxylates of GluA45 and AspA58
from one chain and the carboxylate from Glu C’45 (bidentate coordination) of the
symmetry-related chain with three water molecules completing a capped-octahedral
coordination sphere. An equivalent calcium binding site is also observed in the crystal
packing interface between chains D and B′. No significant increase or reduction of activity
of TM0077 was observed in the presence of calcium ions or EDTA. Therefore, it seems that
these calcium ions are not important for activity. On the other hand, calcium may help
stabilize the structure. No calcium was present in the B. subtilis CAH structure 40; however,
Lys22, Glu26 and Ser25 are conserved and may also act as a calcium binding site.
Enzyme activity
The activity of TM0077 was investigated using p-nitrophenol esters with varying acyl-chain
length, ranging from C2 to C18. TM0077 is only active on the short-chain p-nitrophenol
esters of acetate and propionate and does not hydrolyze esters with acyl chains longer than
four carbons. No significant difference was found in the catalytic efficiency (kcat/Km) for the
hydrolysis of p-nitrophenyl with acyl chains containing 2 to 3 carbons (Table II) [Fig. 1(c)].
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The effect of temperature on activity was studied using p-nitrophenyl acetate as substrate.
The esterase activity increased from 40°C upwards until 100°C [Fig. 5(a)]. An Arrhenius
analysis resulted in linear plots in the temperature ranges of 40–60°C and 60–100°C [Fig.
5(a); inset], with calculated activation energies for the formation of the enzyme substrate-
enzyme complex of 33.7 and 21.9 kJ/mol, respectively. The transition or break in linearity
of the Arrhenius plot at 60°C (1000/T (K) = 3.0) could indicate some conformational change
of the enzyme. TM0077 is fairly resistant to thermal inactivation. An approximate 50%
transient increase in activity is seen during the first 10 to 20 minutes when the enzyme is
incubated at 90°C. After 30 minutes, inactivation of function occurs by first order kinetics
with a half-life of approximately 120 minutes [Fig. 5(b)]. A transient activation has also
been observed for other thermophilic esterases, such from Sulfolobus shibatae 45, and it is
believed that a high temperature is needed in order to obtain the optimal conformation for
catalysis. TM0077 was not stable at 100°C, resulting in a half-life of less than 5 minutes.
However, the optimum temperature and thermal stability of TM0077 are still considerably
higher than those reported for other characterized CE7 esterases, including the
Thermoanaerobacterium enzyme that has a temperature optimum of 80°C and a half-life of
1h at 75°C 46.
The effect of pH on activity was measured in the pH range of 4.8 to 9.2 using the substrate
p-nitrophenyl acetate. TM0077 displayed maximum activity at approximately pH 7.5 [Fig.
5(c)], which is comparable to other CE7 esterases, such as the acetyl xylan esterases from
Thermoanaerobacterium sp. strain JW/SL-YS485 46.
Positional specificity
The positional specificity of TM0077 was tested on three monoacetates of 4-nitrophenyl β-
D-xylopyranoside (pNP-Xyl). To determine the enzyme activity, the β-xylosidase XloA 39
(TM0076) from T. maritima is required as an auxiliary enzyme. This thermostable XloA
enzyme was, therefore, cloned, heterologously expressed, purified to homogeneity, and its
activity was confirmed by measuring release of p-nitrophenol from the substrate pNP-Xyl
(data not shown). The β-xylosidase was not active on the three monoacetates of pNP-Xyl. In
the XloA-coupled assay, TM0077 hydrolyzed acetate from positions 2, 3 and 4 of pNP-Xyl
with similar catalytic efficiency. The results are summarized in Table II.
In addition, TM0077 was investigated for its ability to remove acetyl groups from 7-
aminocephalosporanic acid (7-ACA), cephalosporin C, glucose penta-acetate, N-acetyl-D-
glucosamine, xylan and acetylated xylan. TM0077 has no activity for acetylated and non-
acetylated xylan polymers, indicating that it is, indeed, an acetyl esterase and not an acetyl
xylan esterase. As expected for an acetyl esterase, TM0077 displayed high activity on
glucose penta-acetate with a turnover number of 2680 s−1. Like other members of CE7,
TM0077 was also able to hydrolyze the acetyl groups from both cephalosporin C and 7-
ACA with a turnover number of 376 s−1 and 1140 s−1, respectively. TM0077 was not able to
hydrolyze the acetyl group from N-acetyl-D-glucosamine, indicating that it is specific for
ester bonds and unable to hydrolyze amide bonds.
Inhibitor assays and TM0077 structures complexed with PMSF and paraoxon
PMSF and paraoxon [Fig. 1(d,e)] are competitive irreversible inhibitors of esterases.
Inhibition proceeds by the formation of a reversible Michaelis complex, followed by an
irreversible step and inhibition can, therefore, be characterized by two parameters: a
dissociation constant and a binding rate constant. The inhibition kinetics for paraoxon and
PMSF were investigated in the presence of p-nitrophenyl acetate, as described previously 47,
and the dissociation and rate constants were 0.5 ± 0.1 mM and 0.13 ± 0.02 s−1 for paraoxon,
and 1.1 ± 0.2 mM and 0.020 ± 0.001 s−1 for PMSF, respectively. The acetyl xylan esterase
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from Bacillus pumilus (BpAxe) has slightly reduced sensitivity to paraoxon (dissociation
and rate constants of respectively 5.4 mM and 0.012 s−1), likely due to steric hindrance of
two tyrosine residues (Tyr91 and Tyr206) that hamper the binding of paraoxon. Although
these residues are essentially conserved in TM0077 (Tyr92 and Phe213), TM0077 is more
sensitive to paraoxon than BpAxe48. In comparison to EST2 of Alicyclobacillus
acidocaldarius 49 and EstA of T. maritima 47, the TM0077 dissociation constant is slightly
higher, but the rate constant is comparable. No significant stimulation or reduction of
activity of TM0077 was observed in the presence of divalent metal ions or
ethylenediaminetetraacetic acid (EDTA).
To obtain more information about inhibitor binding and any possible conformational
changes during catalysis, TM0077 was co-crystallized with the inhibitors PMSF and
paraoxon and the PMSF (TM0077-PMS) and paraoxon (TM0077-DEP) structures were
determined to 2.4 Å and 2.1 Å, respectively (Table I). The electron density map of TM0077
with PMSF showed clear density for the PMSF covalent modification. The fluorine was
cleaved from the PMSF molecule during the binding reaction and the phenylmethyl sulfonyl
(PMS) moiety is covalently bound to the Oγ atom of Ser188. The native apo and PMS-
bound structures superimpose well with RMSD’s of 0.09–0.11 Å over 320–321 Cα atoms.
Electron density maps of the paraoxon-bound structure displayed clear density for a diethyl-
phosphate moiety covalently bound to the Oγ atom of Ser188. This covalent modification
indicates that the p-nitrophenol group of paraoxon was cleaved off during co-crystallization,
and a tetrahedral product reminiscent of the first transition state was formed during carboxyl
ester hydrolysis. The native apo and paraoxon-bound structures superimpose with RMSD’s
of 0.12–0.32 Å over 320–322 Cα atoms. Attempts to obtain co-crystals of TM0077 with
cephalosporin C, even at a low temperature of 4°C, were unsuccessful.
Analysis of the active site
TM0077 has a classic catalytic triad, consisting of Ser188 as the nucleophile, His303 as the
proton acceptor/donor, and Asp274 as the acidic residue stabilizing the histidine (Fig. 6).
The catalytic serine Ser188 is located within a conserved pentapeptide sequence, Gly-X-Ser-
X-Gly (GGSQG), characteristic of esterases and lipases. The positions of Ser188, Asp274,
and His303 are consistent with their expected locations in the canonical fold of the α/β-
hydrolase family. Ser188 is located at the nucleophile elbow in a sharp turn between β5 and
helix αC. The presence of three glycine residues (Gly186, Gly187, and Gly190) in close
proximity to Ser188 prevents steric hindrance and facilitates access to the nucleophile
elbow. Asp274 and His303 are located in loops between β7 and helix αE, and between β8
and helix αF, respectively. The oxyanion hole is formed by the backbone amide groups of
Tyr92 and Gln189. The catalytic triad and oxyanion hole are located in a depression on the
surface of TM0077. This ellipsoid pocket (S1), which is approximately 12 Å wide, extends
15 Å from the catalytic serine. A smaller pocket (S2), approximately 5 Å long, extends to
the other side of the catalytic serine [Fig. 6(a)]. The volume of both pockets combined (S1 +
S2) is 1082 Å3 (CASTp analysis; 50). The substrate-binding pocket is bordered by residues
from helices αA and αF, and its base is formed by residues from β-strands 4, 5, 6, and their
adjacent C-terminal loops. The overall pocket is hydrophobic, although it does have some
polar residues (Gln88, Asp210, and Gln314), which may interact with the substrate.
In the native apo structure, the Ser188 hydroxyl makes a hydrogen bond with the imidazole
of His303 [Fig. 6(b)]. Extra density was observed near the side chain of Ser188 and was
interpreted as a chloride ion based on electron density size and shape as well as the
geometry of the interactions with surrounding residues. This chloride ion is bound at the
entrance of the oxyanion hole, forming hydrogen bonds with the backbone amides of Tyr92
and Gln189. In the PMSF-bound structure, the phenyl ring of the inhibitor is located in the
small active site groove surrounded by hydrophobic residues Tyr92, Trp124, Pro228, Ile276,
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and His303 [Fig. 6(c)]. The sulfonyl group of PMSF makes hydrogen bonds with the
backbone amides of Tyr92 and Gln189. In the paraoxon-bound structure, the diethyl-
phosphate (DEP) moiety is stabilized by hydrogen-bonding interactions with the oxyanion
hole. One of the two ethyl arms of bound paraoxon points toward the larger pocket in the
protein, while the other follows the groove of the small pocket. The two ethyl arms are
stabilized by packing against Tyr92, Trp124, Pro228, Ile276, and His303 [Fig. 6(d)].
Two rotamers of the catalytic serine
Although no large conformational changes were observed upon binding of PMSF or
paraoxon, a different rotamer of the catalytic serine side chain was observed compared to
native TM0077 [Fig. 7(a,b)]. Similar changes have been observed in several other esterases
and have been shown to play a key role in the catalytic mechanism (see CONCLUSION for
more details). In the native structure, the catalytic Ser188 Oγ is in the plane of the imidazole
ring of His303, as most commonly observed in the resting state of esterases and lipases 51.
The Ser188 Oγ forms a hydrogen bond (2.6 Å) with His303 Nε2. In the PMSF- and
paraoxon-bound structures, the conformation of the catalytic serine changes; the Ser188 Oγ
rotates about 110°, increasing the distance (3.1 Å and 2.8 Å for PMSF and paraoxon bound
structures, respectively) to the His303 imidazole ring. In the TM0077-SeMet structure, the
catalytic serine is also rotated over ~110°, with a distance to the imidazole ring of 3.0 Å
[Fig. 7(c.)]. A probable explanation for this observation could be the protonation of His303,
since TM0077-SeMet was crystallized at a lower pH (pH 4.2) compared to the native
TM0077 (pH 7.3). Furthermore, extra electron density was identified in the TM0077-SeMet
structure, suggesting a partially occupied acyl intermediate on Ser188. However, as this
density is not sufficiently clear and interpretable to fit an acyl intermediate, water molecules
were modeled instead. No rearrangements of any other residues in the active site were
observed.
CONCLUSION
TM0077 from the hyperthermophilic bacterium T. maritima was predicted from its gene
sequence to be an acetyl xylan esterase. We have expressed and purified TM0077 and
experimentally demonstrated that it has ester-hydrolyzing activity. The TM0077 activity was
restricted to short acyl chain esters (C2 and C3) when artificial p-nitrophenyl-esters were
used as substrates. In addition, the enzyme has high specific activity on glucose penta-
acetate. However, no activity was detected on xylan or acetylated xylan. Thus, TM0077
should be reclassified as an acetyl esterase, and not as an acetyl xylan esterase as currently
annotated 52. Furthermore, the lack of any apparent signal sequence suggests that the protein
is not secreted. Thus, the predicted intracellular location of TM0077 is compatible with a
role other than the deacetylation of extracellular xylan. Based on these results, we conclude
that the likely biological function of TM0077 is removal of the remaining acetyl groups
from the short, end products of xylan degradation that are imported into the cytoplasm. The
resulting deacetylated xylose oligomers are the substrates for a β-xylosidase. This role for
TM0077 is in good agreement with the clustering of the TM0077 gene with other genes
involved in xylose metabolism. However, it cannot be ruled out that TM0077 may also act
on other small, acetylated compounds.
TM0077 is the first esterase from the CE7 family to be tested for its positional specificity for
the deacetylation of 4-nitrophenyl-β-D-xylopyranoside. TM0077 hydrolyzes acetate at the 2,
3 and 4 positions of 4-nitrophenyl-β-D-xylopyranoside with similar efficiency. Conversely,
the CtAxe esterase from Clostridium thermocellum in the CE4 family shows a clear
preference for hydrolyzing acetate at the 2 position 53, and Penicillium purpurogenum AXE
II esterase, a member of the CE5 family, also has a preference for acetate at position 2 54.
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This lack of preference for a specific position of the acetate group correlates with the
relative broad substrate specificity of the CE7 esterases.
Esterases and deacetylases in the CE7 family are unusual in that they are active towards both
acetylated xylo-oligosaccharides and the antibiotic cephalosporin C [Fig. 1(a,b)]. Therefore,
TM0077 was investigated for activity towards the substrates 7-ACA and cephalosporin C.
The activity of TM0077 on these substrates is approximately ten-fold higher than that of the
acetyl xylan esterase from B. pumilus 55 or the acetyl esterase from Thermoanaerobacterium
sp. strain JW/SL YS485 56. TM0077 has a higher hydrolytic activity on 7-ACA compared to
cephalosporin C, as described for other CE7 esterases 40,55,56. Nonetheless, it is unlikely that
both compounds are natural substrates, because the stability of these compounds at the
optimal growth temperature (80°C) of T. maritima is very low.
Crystal structures of TM0077 in complex with inhibitors PMSF and paraoxon revealed that,
upon binding of PMSF or paraoxon, the reaction is trapped at the acylation step via the
formation of a covalent tetrahedral reaction product. In the complexed structures, the
negatively charged oxygen of the tetrahedral intermediate, derived from the substrate
oxyanion, is stabilized by hydrogen bonds with the backbone amide groups of Tyr92 and
Gln189. Comparison of the TM0077 complexed structures with the native structure shows
that the catalytic serine (Ser188) Oγ rotates about 110°, thereby increasing the distance
between Ser188 Oγ and His303 Nε2. Such a conformational change of the catalytic serine
has been observed in several other esterases, including Fusarium solani cutinase 57,
Penicillium purpurogenum acetyl xylan esterase 51, Rhodococcus sp. strain MB1 cocaine
esterase 58, Bacillus subtilis lipase 59, Rhodococcus sp. strain H1 heroine esterase 60, and
Aspergillus niger feruloyl esterase 61. The classical model for the catalytic mechanism of
esterases consists of a sequential two-step hydrolysis. The first reaction involves
nucleophilic attack by the catalytic serine on the substrate carbonyl carbon, resulting in an
acyl-enzyme and the liberation of an alcohol. In the second reaction, a water molecule
performs a nucleophilic attack on the acyl-enzyme, the acyl-enzyme bond breaks and the
carboxylate is released 62. Although the catalytic mechanism of esterases is well established,
it is unclear why the initially generated tetrahedral intermediate does not collapse back to the
reactant complex during the nucleophilic attack of the substrate. A previously proposed
mechanism that would prevent this collapse is the spatial reorganization of the catalytic
residues during the initial catalytic step, causing the residues to separate and thereby drive
the reaction forward 62–64. The apo and inhibitor bound structures of TM0077, presented
herein, support this proposed mechanism. Moreover, in a recent study of the serine protease
mechanism, it was suggested that subtle atomic motions of the catalytic serine and histidine
residues during the catalytic cycle favor the forward reaction 65. Thus, rotation of Ser188 Oγ
of TM0077 may be required to inhibit reversal of the reaction. In addition, such changes
may facilitate the access of water to the catalytic histidine so that the second step of the
reaction can go to completion.
Deacetyl cephalosporins are valuable building blocks for the production of semisynthetic β-
lactam antibiotics. These compounds are derived from cephalosporin C or 7-
aminocephalosporanic acid via enzymatic or chemical processes 10. The thermostable
TM0077 esterase may be valuable in the preparation of derivatives of β-lactam antibiotics.
Recently, the substrate specificity of the acetyl xylan esterase from P. purpurogenum was
engineered to accept a range of fatty acid esters of up to 14 carbons compared to its wild-
type preference for acetate54. It might also be possible to engineer TM0077 and enable the
(de)acetylation of cephalosporins at the C10 position with various acyl chains. Because of
its high stability and activity on 7-ACA and cephalosporin C, TM0077 presents an attractive
candidate for the production of new semi-synthetic antibiotics.
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Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We gratefully acknowledge contributions from George Sheldrick for modifications of the SHELXD program, and
for Global Phasing Ltd. that made significant improvements in the automation of autoSHARP. We also thank
Victor Lamzin for updates of chain docking of the ARP/wARP program, and Gerard Bricogne and Eleanor Dodson
for helpful discussion on phasing for the large TM0077-SeMet structure, and Willem J. van Berkel for valuable
discussion on the catalytic mechanism of TM0077. Portions of this research were carried out at the Stanford
Synchrotron Radiation Lightsource (SSRL) and the Advanced Light Source (ALS). The SSRL is a Directorate of
SLAC National Accelerator Laboratory and an Office of Science User Facility operated for the U.S. Department of
Energy Office of Science by Stanford University. The SSRL Structural Molecular Biology Program is supported by
the DOE Office of Biological and Environmental Research, and by the National Institutes of Health, National
Center for Research Resources, Biomedical Technology Program (P41RR001209), and the National Institute of
General Medical Sciences. The ALS is supported by the Director, Office of Science, Office of Basic Energy
Sciences, Materials Sciences Division, of the U.S. Department of Energy under Contract No. DE-
AC02-05CH11231 at Lawrence Berkeley National Laboratory. Genomic DNA from Thermotoga maritima MSB8
(DSM3109) (ATCC #43589D-5) was obtained from the American Type Culture Collection (ATCC). The content is
solely the responsibility of the authors and does not necessarily represent the official views of the National Institute
of General Medical Sciences or the National Institutes of Health.
Grant sponsor: NIH Grant numbers U54 GM094586 and U54 GM074898 (Protein Structure Initiative); Grant
sponsor: The Graduate School VLAG Wageningen, the Netherlands (ML).
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Figure 1.
Substrates and inhibitors of the CE7 family of enzymes. Structures of (A) acetylated
xylooligosaccharide, (B) cephalosporin C, (C) p-nitrophenyl-acetate, (D)
phenylmethylsulfonyl fluoride (PMSF), and (E) paraoxon.
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Figure 2.
Overall fold and topology of TM0077. (A) Stereo view of a TM0077 protomer. The β-
strands are labeled numerically (-1 to 8) with the core strands in red, α-helices are labeled
alphabetically (A-2 to F) and 310-helices are labeled with an Eta (η1 and η2) with the core
helices in cyan. The three-helix insertion after β6 is colored green and the N-terminal
extension is colored sky blue. The figure was generated using Pymol 31. (B) Topology
diagram of TM0077, with the helices displayed as cylinders and the strands displayed as
arrows following the color and label scheme of (A). The location of residues forming the
catalytic triad is also indicated.
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Figure 3.
Structural superposition of TM0077 with structurally related esterases. Superposition of
TM0077 (yellow) with (A) the cephalosporin C deacetylase (CAH) from B. subtilis (PDB:
1ods; blue) 40 and (B) the α/β-hydrolase domain of the acylpeptide hydrolase/esterase
apAPH from A. pernix K1 (PDB: 1ve6; grey) 43.
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Figure 4.
TM0077 oligomeric assembly. (A) Surface representation of the biological unit of the
TM0077-Native hexamer with each monomer in a different color (left). The “cross section”
shows the entrances on either side of the assembly and the internal cavity (center), and a 90°
rotated view of the TM0077-Native hexamer, with a close-up view of the open central hole
(right). (B) Surface representation of the biological hexamer unit of CAH from B. subtilis 40
(left) and the TM0077-SeMet hexamer with a close-up view of the blocked central hole
(right).
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Figure 5.
Effect of temperature and pH on esterase activity. (A) The esterase activity was studied
using pNP-C2 as a substrate at temperatures ranging from 40–100°C. The inset shows the
temperature dependence as an Arrhenius plot. (B) Thermal stability of TM0077 at 90°C. (C)
The effect of pH on esterase activity studied using pNP-C2 as a substrate at pH values in the
range of 4.8–9.2.
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Figure 6.
TM0077 catalytic site. (A) Surface representation of the TM0077 catalytic site, with His303,
Asp274 and the intermediate DEP-modified Ser188 shown as sticks. The two binding
pockets are indicated with S1 and S2. (B) Apo TM0077 with a bound chloride ion (green
sphere), (C) TM0077 with PMS-modified Ser188 and (D) TM0077 with DEP-modified
Ser188. The catalytic residues are shown as sticks, with the hydrogen bonds shown as
dashed lines. Carbon atoms are in green (apo), cyan (PMS) or blue (DEP), oxygen atoms in
red, sulfur atoms in yellow and phosphate in orange. Electron density omit maps shown for
inhibitor modified Ser188 contoured at 1σ show that the PMS and DEP are covalently
bonded to Ser188 in (C) and (D), respectively. Distances are shown in Ångströms.
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Figure 7.
Conformational change of Ser188 Oγ. The Oγ atom of the Ser188 is rotated ~110° between
the native apo structure (cyan) and (A) the complexed PMS-modified Ser188 structure
(pink), (B) the DEP-modified Ser188 structure (light blue) and (C) the SeMet structure
(purple). The different hydrogen bonds made for the Ser Oγ in the native versus complexed
structures are shown as dashed black lines with distances in Ångströms.
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Table I
Summary of crystal parameters, data collection, and refinement statistics
TM0077-SeMet
TM0077-Native
TM0077-PMS
TM0077-DEP
Space group
P 21
P 21 21 21
P 21 21 21
P 21 21 21
Unit cell parameters
a=152.64Å b=130.95Å
c=157.82Å β=118.90°
a=103.46Å
b=103.79Å
c=221.02Å
a=103.57Å
b=104.50Å
c=221.61Å
a=103.80Å
b=104.43Å
c=221.64Å
Data collection
λ1 MAD-Se
λ2 MAD-Se
Wavelength (Å)
0.9791
0.9183
0.9765
0.9765
0.9765
Resolution range (Å)
29.6 – 2.10
29.6 – 2.10
48.8 – 2.50
49.0 – 2.40
49.0 – 2.12
No. observations
1,119,236
1,100,249
1,222,016
765,546
989,949
No. unique reflections
293,140
291,757
83,045
94,681
123,070
Completeness (%)
93.0 (61.8)a
92.6 (60.8)
100 (100)
100 (100)
89.8 (53.5)
Mean I/σ(I)
9.1 (2.4)a
9.6 (2.2)
14.4 (2.9)
11.5 (3.4)
15.3 (2.2)
Rmerge on I (%)
12.3 (52.5) a
11.9 (57.9)
20.7 (109.7) c
18.0 (67.4)
9.5 (51.9)
Rmeas on I (%)
14.3 (62.2) a
13.9 (68.7)
21.4 (113.6)
19.2 (71.9)
10.2 (60.2)
Rpim on I (%)
7.2 (32.7) a
7.1 (36.2)
5.5 (29.2)
6.7 (24.9)
3.5 (29.2)
Highest resolution shell (Å)
2.15 – 2.10
2.15 – 2.10
2.56 – 2.50
2.46 – 2.40
2.18 – 2.12
Model and refinement statistics
Resolution range (Å)
29.6 – 2.10
48.8 – 2.50
49.0–2.40
49.0 – 2.12
No. reflections (total)
293,097 b
83,045
94,680
122,994
No. reflections (test)
14,726
4,200
4,742
6,188
Completeness (% total)
92.8
100.0
100.0
89.8
Data set used in refinement
λ1 MAD-Se
Cutoff criteria
|F| > 0
|F| > 0
|F| > 0
|F| > 0
Rcryst
0.186
0.167
0.160
0.167
Rfree
0.223
0.212
0.208
0.205
Stereochemical parameters
Restraints (RMSD observed)
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Page 25
TM0077-SeMet
TM0077-Native
TM0077-PMS
TM0077-DEP
Bond angle (°)
1.48
1.47
1.53
1.44
Bond length (Å)
0.018
0.017
0.017
0.015
Av. isotropic B-value (Å2)
27.9
24.7
19.4
19.6
ESU based on Rfree
0.17
0.25
0.22
0.18
Water molecules/other solvent molecules
2,464/1
507/24
946/17
987/23
PDB ID
1vlq
3m81
3m82
3m83
aHighest resolution shell
ESU = Estimated overall coordinate error 16,66.
Rmerge=ΣhklΣi|Ii(hkl)-<I(hkl)>|/ΣhklΣiIi(hkl), Rmeas(redundancy-independent Rmerge)=Σhkl[Nhkl/(Nhkl-1)]1/2Σi|Ii(hkl)-<I(hkl)>|/ΣhklΣiIi(hkl), and Rpim(precision-indicating Rmerge)=Σhkl[1/
(Nhkl-1)]1/2Σi|Ii(hkl)-<I(hkl)>|/ΣhklΣiIi(hkl) 67–69.
Rcryst = Σ| |Fobs|-|Fcalc| |/Σ|Fobs| where Fcalc and Fobs are the calculated and observed structure factor amplitudes, respectively.
Rfree = as for Rcryst, but for 5.0 % of the total reflections chosen at random and omitted from refinement.
bTypically, the number of unique reflections used in refinement is slightly less than the total number that were integrated and scaled. Reflections are excluded due to systematic absences, negative
intensities, and rounding errors in the resolution limits and cell parameters.
cRmerge of the highest resolution shell is high due to high redundancy (14.7). However, the completeness and mean I/σ of the highest resolution shell are reasonable, and these data were included in the
refinement.
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Table II
Kinetic parameters for hydrolysis of various esters
Ester
Km (mM)
kcat (s−1)
kcat/Km (s−1 mM−1)
pNP-Acetate
0.185 ± 0.026
57.5 ± 2.2
310.8 ± 45.3
pNP-Propionate
0.137 ± 0.013
41.3 ± 1.1
301.5 ± 29.7
2-O-acetyl pNP-Xyl
3.6 ± 0.5
76.1 ± 19.2
21.1 ± 6.1
3-O-acetyl pNP-Xyl
4.2 ± 0.4
70.1 ± 7.7
16.7 ± 2.4
4-O-acetyl pNP-Xyl
4.0 ± 0.1
78.6 ± 12.9
19.7 ± 3.3
Proteins. Author manuscript; available in PMC 2013 June 01.
|
3M85
|
Archaeoglobus fulgidus exosome y70a with RNA bound to the active site
|
Quantitative analysis of processive RNA degradation
by the archaeal RNA exosome
Sophia Hartung1, Theresa Niederberger1, Marianne Hartung2, Achim Tresch1 and
Karl-Peter Hopfner1,*
1Center for Integrated Protein Sciences and Munich Center for Advanced Photonics at the Gene Center,
Department of Biochemistry, Ludwig-Maximilians-University Munich, Feodor-Lynen-Strasse 25, 81377 Munich
and 2General Electric - Global Research, Freisinger Landstrasse 50, 85748 Munich, Germany
Received February 9, 2010; Revised March 18, 2010; Accepted March 21, 2010
ABSTRACT
RNA exosomes are large multisubunit assemblies
involved
in
controlled
RNA
processing.
The
archaeal exosome possesses a heterohexameric
processing
chamber
with
three
RNase-PH-like
active sites, capped by Rrp4- or Csl4-type subunits
containing RNA-binding domains. RNA degradation
by RNA exosomes has not been studied in a quan-
titative manner because of the complex kinetics
involved, and exosome features contributing to effi-
cient RNA degradation remain unclear. Here we
derive a quantitative kinetic model for degradation
of a model substrate by the archaeal exosome.
Markov Chain Monte Carlo methods for parameter
estimation allow for the comparison of reaction
kinetics between different exosome variants and
substrates.
We
show
that
long substrates
are
degraded in a processive and short RNA in a more
distributive manner and that the cap proteins influ-
ence degradation speed. Our results, supported by
small angle X-ray scattering, suggest that the
Rrp4-type cap efficiently recruits RNA but prevents
fast RNA degradation of longer RNAs by molecular
friction,
likely
by
RNA
contacts
to
its
unique
KH-domain. We also show that formation of the
RNase-PH like ring with entrapped RNA is not
required for high catalytic efficiency, suggesting
that the exosome chamber evolved for controlled
processivity, rather than for catalytic chemistry in
RNA decay.
INTRODUCTION
The eukaryotic and archaeal RNA exosomes and their
distant relative, the bacterial degradosome, are large
multiprotein assemblies that function as central cellular
RNA
processing
and
degradation
machineries.
The
RNA exosome was originally found in yeast as an essen-
tial protein complex with 30 ! 50 exonuclease activity.
First, identified for the 30-processing of the yeast 5.8S ribo-
somal RNA (1), the yeast RNA exosome subsequently
turned out to be important for the trimming and degrad-
ation of the 30-end of several nuclear RNA precursors (2).
In addition, the exosome was shown to be also active in
the cytoplasm by controlling mRNA turnover (3), and by
its implication in various mRNA surveillance pathways
like the non-sense-mediated and the non-stop decay
pathways (4–7). Due to its involvement in all the different
RNA processing and surveillance pathways the exosome is
apparently one of the central exonucleases of a yeast cell
[for reviews see for instance (8,9)].
Structural homologues of the yeast exosome were sub-
sequently identified in humans, previously known as the
PM-Scl
(polymyositis–scleroderma
overlap
syndrome)
complex, and in archaea (10–12). A variety of structural
studies revealed a conserved architecture of exosome like
complexes (13–18): exosomes consist of nine conserved
core subunits, six RNase PH type subunits and three
subunits with S1 and KH or zinc-ribbon domains. The
six RNase-PH like domains form a ring, arranged as
trimers of pseudo-dimers. In archaea, the ring is formed
by three (archaeal)aRrp41:aRrp42 dimers, while human
and yeast exosomes contain six different RNase PH type
subunits.
The archaeal exosome possesses a central chamber
within the RNase PH ring which contains three phos-
phorolytic active sites. The actual active site is located in
the aRrp41 subunits, but the whole aRrp41:aRrp42 dimer
is involved in positioning the RNA. These sites degrade
single-stranded RNA (ssRNA) in a phosphate dependent
manner in 30 ! 50 direction. They also catalyse the reverse
reaction of adding nucleoside diphosphates to the 30-end of
RNA (13), liberating inorganic phosphate. In archaea, this
*To whom correspondence should be addressed. Tel: +49 89 2180 76953; Fax: +49 89 2180 76999; Email: hopfner@lmb.uni-muenchen.de
5166–5176
Nucleic Acids Research, 2010, Vol. 38, No. 15
Published online 14 April 2010
doi:10.1093/nar/gkq238
The Author(s) 2010. Published by Oxford University Press.
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/
by-nc/2.5), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.
activity has been attributed to formation of poly-A-rich
tails on RNA (19). A proposed RNA entry pore at one
side of the chamber restricts entry to mostly unstructured
ssRNA, providing an explanation for controlled RNA deg-
radation. Furthermore, three Csl4 or Rrp4 type putative
RNA recognition subunits are located on top of the
(Rrp41:Rrp42)3 ring and frame the proposed RNA entry
pore. Current models suggest that these domains recognize
RNA substrates and help to funnel them into the process-
ing chamber.
Although the human exosome is structurally related to
the archaeal complex, including S1 and KH domain con-
taining subunits (Csl4, Rrp4 and Rrp40), it has lost
phosphorolytic activity (14). Instead, it gained additional
ectopic subunits: the hydrolytic RNase Rrp44 (20,21)
has both exonucleolytic and endonucleolytic activities,
located in RNB and PIN domain subunits, respectively
(14,20,22–24). A second hydrolytic RNase (Rrp6) was
identified as transient part of the nuclear complex (10).
Recent results indicate that despite the ectoptic placement
of the nuclease active sites, RNA is still threaded through
the nuclease deficient RNase PH type ring (25).
A variety of groups have biochemically observed
processive
RNA
degradation,
in
particular
for
the
archaeal
exosome.
From
structural
studies,
it
was
proposed that RNA is channelled through an entry pore
between the S1 domains of aCsl4 or aRrp4 trimers into the
processing chamber, where the 30-end of the RNA is pos-
itioned in one of the three phosphorolytic active sites,
subsequently degrading RNA base-per-base (16,26,27).
The presence of the cap proteins Csl4 and Rrp4 in
general
increases
the
degradation
efficiency
of
the
exosome, but it is unclear how they do so. For instance,
if the cap proteins recruit RNA, one would expect an
increase in the general binding affinity. However, once
RNA has entered the processing chamber, high affinity
binding to the ectopic domains should slow down
processive degradation. Another mechanism that is not
yet understood is why processivity depends on the length
of the RNA molecules (28). To address these questions
and to develop means to quantitatively analyse processive
degradation, we performed quantitative high-resolution
RNase degradation activity assays with different variants
of the Archaeoglobus fulgidus exosome. We evaluated dif-
ferent kinetic models and developed a Markov Chain
Monte Carlo (MCMC) analysis to fit the model to the
data and derive appropriate rate constants of individual
RNA degradation steps. Our data identify different struc-
tural contributors to processivity, suggesting that ectopic
RNA-binding domains, the entry pore and the active site
are different contributors to processive degradation. The
methods
should
be
easily
applicable
also
to
other
processive enzymes, including the hydrolytic nucleases of
the eukaryotic exosomes.
MATERIALS AND METHODS
Protein expression and purification
The Archaeaoglobus fulgidus RNA exosome with different
Rrp4 and Csl4 caps, derivatives and mutants were
expressed and purified as described (29). Site directed mu-
tations
were
introduced
using
the
QuickChange
Site-directed Mutagenesis Kit (Stratagene) and verified
by sequencing. Oligonucleotide sequences are provided
in the Supplementary Table S2.
Crystallization and structure determination
An amount of 120 mM Csl4-exosome (Csl4:Rrp41:Rrp42)3
or its Y70ARrp42 mutant (= 27 g/l) were incubated with
400 mM RNA (3.3-fold excess, 6-mer CCCCUC) for
10 min on ice. Protein:RNA complexes were crystallized
by sitting drop vapour diffusion technique by mixing 1 ml
protein and 1 ml of reservoir solution (0.1 M NaAcetate,
pH 4.6, 30% 3-Methyl-1,5-pentadiol (MPD), 100 mM
NaCl) at 20C. Datasets were recorded at the ID-14-2
beamline (ESRF, Grenoble, France) to 2.4 A˚
(wild-type
exosome) and at the PX I beamline (SLS, Villigen,
Switzerland) to 3.0 A˚ (Y70ARrp42 mutant) and processed
with X-ray Detector Software (XDS) (30). A model of the
apo-Csl4-exosome complex (29) was used as a search
model for molecular replacement using PHASER (31).
Refinement to 2.4 A˚ and 3.0 A˚ , respectively was performed
with CNS (32) and PHENIX (33). In the additional
electron density RNA nucleotides were positioned using
COOT (34). Refinement of the complete complexes was
followed by iterative cycles of manual model completion
with COOT and positional and B-factor refinement with
CNS (Supplementary Table S1).
Small angle X-ray scattering
For small angle X-ray scattering (SAXS) studies, the
(Rrp41:Rrp42:Rrp4)3 complex was purified as described
above. To purify the exosome with endogenously bound
Escherichia coli RNA the protocol was modified as
follows: RNA was not washed offwith high salt, and in
all buffers the salt concentration was 250 mM or lower.
After the Ni-NTA affinity chromatography, the complex
was loaded on an anion exchange column to remove
unbound nucleic acids and the procedure was repeated
to assure the total removal of free RNA. Not until only
one distinct peak was eluted, the fractions were pooled,
concentrated and flash frozen. The apo-complex was
measured at 5, 10 and 15 mg/ml and the RNA complex
was
concentrated
to
an
absorption
at
280 nm
of
A280 = 55 and measured in a 1: 0, 1: 1 and 1: 2 dilution
to evaluate the concentration dependency of scattering.
Both complexes did not show concentration dependent
aggregation and were not affected by long exposure to
high-energy X-rays. SAXS data collection was performed
in 20 mM Tris pH 7.4 and 200 mM NaCl buffer at the
SIBYLS beamline (Advanced Light Source, Berkeley,
CA, USA) (35). The radius of gyration was calculated
using the Guinier plot in the linear region (constraint: s
Rg <1.3) and the calculation of the pair distribution
function was done with GNOM within PRIMUS (36).
Ab initio modelling of the solution structures was done
with GASBORp (37) and more than 10 identically
calculated models were aligned and averaged using
DAMAVER and SUPCOMB (38). For analysis of the
Nucleic Acids Research, 2010, Vol. 38, No. 15
5167
bound RNA, the protein was separated from the RNA by
running the complex on a denaturing 6 M urea and 20%
polyacrylamide gel and elution of the RNA from the gel.
The pelleted RNA was sent to Vertis Biotechnologie AG,
where the sample was poly(A)-tailed using poly(A) poly-
merase followed by ligation of an RNA adapter to the
50-phosphate of the small RNAs. First-strand cDNA syn-
thesis was then performed using an oligo(dT)-adapter
primer and M-MLV H- reverse transcriptase. The result-
ing cDNAs were PCR-amplified to 20–30 ng/ml in 19
cycles using standard Taq DNA polymerase. We cloned
the cDNA products with EcoRV into pET21 vectors,
transformed and amplified the plasmids and isolated and
sequenced single clones.
Cross-linking
Site-specific crosslinking of the K37CRrp41:D143CRrp42
mutant was performed with a HBVS (1,6-Hexane-bis-
vinylsulfon) crosslinker. The crosslinking reaction was
performed with a 100-fold excess of crosslinker under
oxygen-free conditions in a glove-box. We removed
crosslinked
protein
from
non-crosslinked
protein
complexes using a Superose 6 size-exclusion column,
equilibrated
with
a
running
buffer
containing
4 M
guanidinium chloride. Protein from the peak correspond-
ing to a crosslinked Rrp41/Rrp42 dimer was refolded in
50 mM Tris (pH 7.4), 200 mM NaCl, 500 mM arginine and
5% glycerol by dilution. The refolded protein was again
applied onto a Superose 6 column in 50 mM Tris (pH 7.4)
and 200 mM NaCl. Only the correctly refolded protein,
verified by the formation of a hexamer in size exclusion
chromatography, was used for further experiments. As a
control sample, the same complex without crosslinker was
partly denatured, purified and refolded in the same way as
the crosslinked protein.
RNase activity assays
We carried out RNase activity assays using 32P-labelled
poly(rA)-oligoribonucleotides with different lengths as
substrate (26). RNA was incubated with [g-32P] ATP
(Hartmann Analytics) and T4 polynucleotide kinase
(NEB)
for
45 min
at
37C
and
purified
by
using
MicroSpin G-25 columns (GE Healthcare). For each
reaction, the protein (30 nM for the Csl4- and the
Rrp4-capped
wild-type
exosome
and
the
interface
mutant; 60 nM for the cap-less exosome and the single
site mutants R65ERrp41 and Y70ARrp42; 120 nM for the
crosslinked cap-less exosome) was incubated with RNA
in buffer containing 20 mM Tris (pH 7.8), 60 mM KCl,
10 mM MgCl2, 10% glycerol, 2 mM DTT, 0.1% PEG
8000, 10 mM NaH2PO4 (pH 7.8) and 0.8 U/ml RNasin
(Promega) at 50C. Different time points were taken and
the reaction was stopped by adding one volume of loading
dye [0.75 g/l bromphenol blue, 0.75 g/l xylene cyanol, 25%
(v/v) glycerol, 50% formamide]. The reaction products
were
resolved
on
a
20%
polyacrylamide/6 M
urea
sequencing gel running at 50C and were analysed
by phosphorimaging (GE Healthcare). The gel bands
were
quantified
using
the
ImageQuant
Software
(GE Healthcare) and data analysis, simulation and
fitting was done with MatLab (Mathworks).
Models and kinetic data analysis
Kinetic models are shown in Figure 3A. They are
described by four parameters: association rate ka,i, dissoci-
ation rate kd,i, cleavage rate kc,i and polymerization rate
kp,i, one for each RNA of length i = 4, 5, . . ., 30. The cor-
responding set of differential equations that quantitatively
describe RNA degradation is shown in the supplement
Data (Chapter 1). Since the reaction takes place in an
excess
of
inorganic
phosphate
(10 mM
phosphate
compared to only 3.6 mM ADP at the time all RNA mol-
ecules are totally degraded), we may assume no polymer-
ization takes place, i.e. kp,i = 0 for all i. Consistently, we
saw no synthesis of longer RNAs in our reactions. To
obtain empirical estimates of the posterior parameter dis-
tribution, we implemented a MCMC approach based on
the Metropolis–Hastings algorithm. The key ingredients
are the likelihood function, the prior, and the proposal
distribution. The likelihood function penalizes the estima-
tion error produced by a given model. More precisely, it
penalizes the residuals, i.e. the deviation of the measured
RNA amounts at each time point from the amounts that
have been predicted from the current parameter set. We
assume that the residuals are independent realizations of
Gaussian distributions with zero mean. Since the vari-
ances of these Gaussians are not known a priori, we
assume a two-parameter error model with an additive
and a multiplicative error component which has been
proposed (39) in the context of spot quantification on
arrays. We initialize the error model very conservatively
(presuming
large
measurement
errors).
During
the
MCMC run, the error model is updated continuously by
replacing it with an empirical estimate derived from the
residuals that occurred in the Markov chain so far.
The prior encodes prior knowledge/assumptions on
the distribution of the parameters. It is sensible to
require the kinetic parameters to vary smoothly with the
RNA length i. This is made explicit by penalizing the
difference of two successive kinetic parameters kx,i+1 and
kx,i using a Gaussian prior on these differences. We em-
phasize that this does not impose any restrictions on the
absolute level of the parameter values. The comparison of
the parameter levels obtained by different experiments is
virtually unaffected by our prior choice and therefore
practically unbiased. The proposal distribution generates
a new parameter set as a candidate for the next MCMC
step that is based on the current parameter set. We simply
use a multivariate log-normal distribution with fixed
diagonal covariance matrix, which is centred at the
current parameter set.
It turns out that the parameters of the model as stated
above are not identifiable. We therefore fixed kd,i to
one global constant kd, whereas the association param-
eters ka,i are sampled individually. The parameters kc,i
are set equal to one length-independent parameter kc.
The details of this approach and its justification through
extensive simulations are given in the supplementary Data
(Chapters 2–4).
5168
Nucleic Acids Research, 2010, Vol. 38, No. 15
RESULTS
RNA is not degraded with constant velocity
Despite intense structural and biochemical research on
RNA exosomes, a kinetic model, quantitative analysis of
processive RNA degradation and a biochemical identifi-
cation of elements that contribute to processive degrad-
ation have not been studied, due to the complex kinetics
involved. To address these issues, we performed RNase
assays with 50-radioactively labelled 30-mer oligo(A)
RNAs and the A. fulgidus Csl4- (Csl4:Rr41:Rrp42)3 and
Rrp4-exosomes
(Rrp4:Rr41:Rrp42)3.
The
reaction
products and their time evolution were resolved on
a
denaturing
sequencing
gel
and
quantified
by
phosphorimaging (Figure 1), controls are shown in the
supplemental material (Supplementary Figures S4 and
S5). Several characteristic features of substrate degrad-
ation by exosomes are revealed:
First, RNA is not degraded at a constant speed, but
the degradation of substrate has several phases and is
distinct in different isoforms. In the Csl4 exosome
(Figure 1A), after a slower first processing step, longer
RNAs (>12–13 nt) are degraded very fast, seen by
the low amount of intermediates in this range; shorter
RNAs (<12–13 nt) are degraded slower and accumulate
first before they are further degraded. On the contrary,
the first processing step is faster in the Rrp4- than in the
Csl4-exosome (Figure 1B). However, oligo-rA substrates
>24 nt are degraded slower, intermediate substrates
(24–13 nts) faster, and RNAs <13 nt slower again.
This result is astonishing, considering homooligomeric se-
quences are used and the effect is consequently not
sequence
dependent.
In
addition,
the
unexpected
slow-fast-slow kinetics of the Rrp4 isoform reveals a
quite complex length dependency of RNA degradation
speeds.
Second, the final degradation product is a 3-mer.
Further
degradation
is
extremely
slow,
comparable
to spontaneous background hydrolytic cleavage under
the present conditions. We hypothesized that features
of the active site might interact specifically with the
fourth base at the 30-end. Previous structural analysis
with the Sulfolobus solfataricus exosome has shown that
at least 4 nt are stably bound in the phosphoropytic active
sites (26), but in the case of the Pyrococcus furiosus
exosome some nucleotides were recognized (16). To get
direct
structural
information
for
the
A.
fulgidus
exosome:RNA
interaction,
used
in
this
study,
we
crystallized
our
Csl4-exosome
with
a
6-mer
RNA
molecule (Figure 2; Supplementary Table S1). Four nu-
cleotides from the 30-end are clearly visible in the unbiased
Fo–Fc electron density, with weaker density for the two
additional nucleotides. Interestingly, the side chain of
Y70Rrp42
shows
p-stacking
with
the
fourth
base
(counting from the active site) and this seems to be a
conserved feature among archaeal exosomes (16,26).
This interaction specifically stabilizes the first 4 nt, while
RNA positions+5 and+6 behind Y70Rrp42 appear not to
be specifically recognized. To test the role of Y70Rrp42, we
determined
the
co-crystal
structure
of
the
Csl4-
exosome-Y70A mutant with a CCCCUC oligonucleotide.
In fact, we only see clear electron density for 4 nt in the
active site and the electron density at position+4 is weaker
and less defined compared to the wild-type. Thus,
the 3-mer as degradation end-product is likely the
cause of inefficient recognition of RNA’s with <4 nt at
the active site.
Figure 1. Visualization of RNase activity of the archaeal exosome on denaturing polyacrylamide gels: the input (I) is a 30-mer polyA RNA
radioactively labelled at the 50-end that is degraded from the 30-end to a final product (FP) of a 3-mer. Time points were taken in increasing
intervals [in minutes: 0:10; 0:20; 0:30; 0:40; 0:50; 1:00; 1:10; 1:20; 1:40; 2:00; 2:20; 2:40; 3:00; 3:30; 4:00; 4:30; 5:00; 5:50; 6:00; 6:30; 7:00; 7:30; 8:00;
9:00; 10:00; 12:00; 14:00; 16:00; 18:00; 20:00; 25:00; 30:00; 35:00; 40:00; (B) ends at 8:00 min]. RNA degradation does clearly not occur with constant
speed and the (Csl4:Rrp41:Rrp42)3 exosome (A) degrades RNA with a different time dependency than the (Rrp4:Rrp41:Rrp42)3 exosome (B).
Nucleic Acids Research, 2010, Vol. 38, No. 15
5169
A kinetic model for processive RNA degradation
by exosomes
To obtain a comprehensive picture of the exosomal RNA
decay, we need to analyse the reaction speeds in a quan-
titative manner. The amount of RNA as function of time
of intermediate i of an rA n-mer may be described by
several rate constants (Figure 3A): an association rate
constant ka,i of the 30-end of RNA to the active site; a
corresponding dissociation rate constant kd,i; a rate of for-
mation of intermediate i by cleavage of intermediate i+1,
Figure 3. Three different models to describe the kinetics of RNA degradation by the exosome were tested: (A) scheme for the general kinetic
model, which includes cleavage and polymerization rates kc and kp as well as association and dissociation rates ka and kd for all RNAs from 30–4 nt.
(B–D) Quantified concentrations of RNA intermediates from Figure 1A, along with least square fits to different kinetic models. (B) Strict processivity
considers only 27 different cleavage rates kc,30 –kc,4. (C) cleavage-and-polymerization considers 27 different cleavage rates kc,30 –kc,4, 27 different
polymerization rates kp,30–kp,4 and one initial association rate ka,30 (=55 rates). With models (C) and (B), no reasonable fit could be obtained. (D) By
including association, dissociation and cleavage and making rational simplifications (see text) we can convincingly fit the data with a model con-
taining 28 different rate constants.
Figure 2. Crystal structure of 6-mer RNA bound to the active site of the archaeal exosome. Rrp41 is shown in light and Rrp42 in dark green. The
2Fo–Fc electron density is contoured at 1.0s and only shown for the RNA and the side chain of Y70Rrp41. (A) In the wild-type exosome Y70 is
stacking with the fourth base of the bound RNA, and only weak density can be seen for the fifth and sixth base. (B) Electron density for the fourth
base of the RNA is much weaker in the Y70ARrp41 mutant compared to the wild-type and no density can be detected at this contour level for
additional nucleotides.
5170
Nucleic Acids Research, 2010, Vol. 38, No. 15
kc,i+1; and by adenylation (polymerization) of intermedi-
ate i1, kp,i-1; a rate of disappearance of intermediate i by
cleavage of i, kc,i and by adenylation of i, kp,i. The system
kinetics is then given by a set of differential equations
(Supplementary Data).
However, it is possible that this more general model can
be further simplified. For instance, we likely can neglect
adenylation (kp,i = 0) because our reaction conditions
contain 10 mM phosphate compared to only 3.6 mM
ADP
at
the
time
all
RNA
molecules
are
totally
degraded,
strongly
shifting
the
reversible
reaction
towards degradation. In addition, the exosome might
be strictly processive, i.e. association and dissociation
rates of RNA intermediates are negligible compared to
the
cleavage
rates
(ka,i = kd,i = 0).
Furthermore,
all
cleavage rates may be independent of the length of
RNA, because they could be a local active site property
(kc,i = kc,j). Hence we analysed three simplified models
(Figure 3B–D). Once initial values for the rate constants,
enzyme concentration and RNA substrate concentration
(rA 30-mer) are provided, this corresponding set of differ-
ential equations can be used to calculate the concentra-
tions of all RNA intermediates over time. We then
minimized the resulting least square deviations between
the
calculated
and
experimental
concentrations
of
reaction intermediates by optimizing the rate constants
using the ‘fminsearch’ parameter optimization procedure
as implemented in Matlab.
Using the ‘strict processivity’ model with 27 independ-
ent
variables
(ka,i = kd,i = kp,i = 0)
(Figure
3B),
we
obtained no reasonable fit of the experimental data. A
second model including the adenylation reaction (55 inde-
pendent variables, ka,i = kd,i = 0) could also not properly
interpret the data (Figure 3C). Thus, simply adding more
parameter does not automatically lead to reasonable fits
and the RNA degradation activity cannot be convincingly
explained by strict processivity. Consequently, we added
association and dissociation of RNA intermediates to the
equations and used the following alternative simplifica-
tions: (i) adenylation is omitted (kp,i = 0; see above); (ii)
the same cleavage rate is used for all RNA molecules
(kc,i = kc,j),
i.e.
cleavage
rate
is
a
local
active
site
property
and
not
dependent
on
RNA
length.
We
estimated starting values for kc and validated this simpli-
fication from analysis of the initial exponential decay of
RNAs substrates with different initial lengths (data not
shown). (iii) Due to our experimental approach, we
cannot experimentally distinguish between bound and
free RNA since the gel bands represent the sum of free
and exosome-bound RNA intermediates of length i. For
that reason, we cannot reconstruct dissociation-, associ-
ation- and cleavage rate constants independently of each
other. Consequently, we do not treat the association and
the dissociation rate constants independently, but analyse
the ratio of ka,i/kd,i by setting kd,i’s to a constant low value,
leaving ka,i free to vary. Variation of the value for kd did
not
result
in
significant
changes
in
the
analysis
(Supplementary Data). These three reasonable simplifica-
tions leave essentially one free parameter per intermediate
plus one overall cleavage rate constant. Although this
model has less degrees of freedom than the second
model (55 versus 28), it can convincingly interpret the ex-
perimental data for the both Csl4 and Rrp4 exosome
variants and most mutants (Figure 3D).
MCMC analysis of degradation
To address the problem of multidimensional parameter
fitting and to assess the variance in parameter esti-
mation, we established MCMC simulations. Because of
the difficulty in determination of separate values for
the single rate constants, we defined an RNA length-
dependent quantity vi
vi ¼ kc,i
Km,i
ð1Þ
with Km,i the Michaelis–Menten constant
Km,i ¼ kc,i+kd,i
ka,i
ð2Þ
vi is called ‘catalytic efficiency’ or ‘specificity constant’, as
it is a measure of the velocity of RNA intermediate i deg-
radation by the exosome. We are now in a position to test
exosome features important for vi. We observe that for the
Csl4 exosome, vi is highly dependent on the RNA length:
the initial RNA processing step, likely determined by the
initial association of RNA with the exosome, is generally
slow. Once RNA is bound, vi is large and relatively
constant for RNA lengths >13 nt. vi then progressively
decreases for RNA lengths <13 nt until the final 3-mer
appears (Figures 4B and 5A). This length dependency may
be explained by the exosome structure: RNA molecules
longer than 13 nt might still reach through the ‘neck’,
and this topological interaction will induce a higher
‘local concentration’ of RNA at the active site with
increased vi. Short RNAs, on the contrary, will lose this
contact and due to their smaller size more easily diffuse
out of the processing chamber, therefore decreasing vi.
To test this idea, we analysed the Y70A mutant of the
Csl4-exosome. The length profile of the catalytic efficiency
has a similar shape than for the wild-type, although the
catalytic efficiency is lower for all RNA intermediates
(Figure 5B). For RNAs >13 nt, the difference in vi is
2- to 3-fold (about one log unit). However, the drop in
vi for RNAs <13 nt is progressively more pronounced
compared to the wild-type and towards short RNAs
(<8 nt), the mutant is 20- to 150-fold (three to five log
units) slower than the wild-type. This is consistent with the
idea that for long RNAs the neck provides additional
interaction and thus overcomes in part the destabilizing
effect of Y70A. For shorter RNAs, the active site becomes
the sole attachment, leading to a rapid drop of catalytic
efficiency in the Y70A mutant.
We also analysed the ‘neck’ mutant R65E, which has
been shown to severely reduce exosome activity (16,26,27).
This mutant exhibited a substantially delayed onset of
degradation, presumably because RNA is unable to effi-
ciently enter the active site (data not shown). A likely
reason is the formation of non-productive RNA:protein
complexes with RNA trapped on the outside of the
exosome (29). At present, our model cannot deal with
Nucleic Acids Research, 2010, Vol. 38, No. 15
5171
this scenario and we could not convincingly include—as
only variant—the R65E mutant in the analysis. However,
the data of the analysis of R65E are provided in the sup-
plementary Figure S15 and following.
Role of exosome ring formation and ring dynamics for
RNA binding
Although the initial binding of RNA appears slow, it
seems unlikely that the 30-end directly finds its way
through the small hole in the neck. It is perhaps more
likely that the ring structure ‘breathes’—as observed e.g.
in hexameric helicases—and allows some lateral entry at
the neck. To explore this idea we analysed a crosslinked
exosome, where the ring is rigidified by three site specific
crosslinks, and a mutant that disrupts the ring structure
into Rrp41:Rrp42 pairs. We compared these isoforms with
the
corresponding
wild-type,
the
cap-less
hexameric
(Rrp41:Rrp42)3 ring (Figure 5C). From the structural
analyses, it was observed that the Rrp41 and Rrp42
subunits possess two interfaces. One interface is larger,
and characterized by contiguous b-sheets between Rrp41
and Rrp42 (40). The other interface is smaller, presumably
more dynamic, and was chosen for the crosslinking and
mutagenesis
analysis
(Supplementary
Figure
S2).
(Rrp41:Rrp42)3 exhibit a biphasic length dependence of
vi, similar to the Csl4-exosome. However, the catalytic
efficiency of (Rrp41:Rrp42)3 is 5- to 10-fold higher
Figure 5. Comparisons
of
the
catalytic
efficiency
vi
of
different
exosome variants versus RNA lengths: (A) differences in the cap
proteins influence catalytic activity. This is shown by comparison of
vi from the cap-less exosome (Rrp41:Rrp42)3 in magenta, the Csl4
capped exosome (Csl4:Rrp41:Rrp42)3 in red and the Rrp4 capped
exosome (Rrp4:Rrp41:Rrp42)3 in blue. (B) Tyr70Rrp42 close to the
active site is especially important to efficiently degrade small RNAs.
The wild-typ Csl4 exosome is shown in red and the Y70ARrp42 mutant
in green. (C) The role of the ring architecture and dynamics for cata-
lytic activity is shown by comparing wild-type cap-less exosome
(Rrp41:Rrp42)3 in magenta with the dimeric and open interface
mutant (Rrp41:Rrp42)1 and a rigidified crosslinked variant that is less
dynamic in yellow. A total of 1000 parameter sets have been randomly
drawn from the stationary phase of the Markov chain. Thus for each
RNA length and each timepoint, we obtained 1000 estimates whose
distribution is displayed by boxplots.
Figure 4. Catalytic efficiency vi for all RNA intermediates present
during the degradation of a 30-mer RNA by the Csl4-Rrp41-Rrp42
exosome was determined with MCMC simulations. (A) shows the
traceplot and (B) the final parameter set (burnin = 150 000). It can be
seen that the MCMC chains vary in convergence speed as well as in
variability. The boxplots in (B) illustrate the main advantage of the
MCMC approach: it not only offers a set of parameters that best
describe the measured data, but it also yields a posterior distribution
for each catalytic efficiency parameter and thus provides a more com-
prehensive summary of the data.
5172
Nucleic Acids Research, 2010, Vol. 38, No. 15
across the RNA spectrum than that of the Csl4-exosome,
indicating that the Csl4 cap subunits do not substantially
promote
degradation
of
this
model
substrate.
In
addition,
vi
drops
for
RNAs
<13 nt
even
for
the
(Rrp41:Rrp42)3 particle, indicating that the neck not
e.g. the S1 domains of caps are responsible for the
higher catalytic efficiency on longer RNAs.
To explore the effect of the hexameric ring formation,
we mutated Lys51 to Glu, located in the ‘smaller’ interface
between alternating Rrp41 and Rrp42 pairs. This resulted
in stable Rrp41:Rrp42 dimers that do not assemble into
hexamers
anymore
(Supplementary
Figure
S3).
The
Rrp41:Rrp42 dimers exhibit catalytic efficiencies that are
only slightly lower than the (Rrp41:Rrp42)3 particle for
RNAs >13 nt, and almost identical to the corresponding
hexamers for RNAs <13 nt. As a result, the drop around
13 nt from a faster to a slower degradation is much less
pronounced in the Rrp41:Rrp42 dimer, further supporting
the idea that encapsulation in the neck is responsible for
higher degradation speeds. The relatively high activity of
the dimers is possibly also a result of the effective ‘tripli-
cation’ of active sites, i.e. only one RNA molecule can be
degraded by a (Rrp41:Rrp42)3, while three RNA mol-
ecules can be degraded by three Rrp41:Rrp42 dimers. In
addition, while RNA probably dissociates faster from the
dimers, this effect could be compensated by a faster asso-
ciation of RNA to the readily accessible active sites in the
open dimers.
The opposite is observed, when the ring structure is
crosslinked. We introduced cysteines on the outside of
the
RNase-PH
ring
and
crosslinked
the
three
Rrp41:Rrp42 dimers via a thiol specific bifunctional
crosslinker. This procedure resulted in a hexameric
RNase PH ring with wild-type-like size and shape accord-
ing
to
gel
filtration
(Supplementary
Figure
S3).
Comparison of ni between the crosslinked isoform with
the (Rrp41:Rrp42)3 hexamer, revealed a dramatically
reduced ni (500- to 2000-fold) indicating that rigidifying
the exosome by the crosslink severely affects catalytic ef-
ficiency. We cannot formally rule out that the crosslinking
affects activity by other means, but considering that the
hexamer disrupting mutation at the same interface does
not severely reduce activity, a plausible scenario is that the
rigidified exosome does not allow efficient association with
RNA anymore.
Thus, taken together, the self-compartmentalization of
exosomes is probably not an evolution for high activity,
but rather for controlled RNA degradation.
Effect of the cap structures
To learn about the role of the cap proteins in exosome
activity, we compared the rate constants of the cap-less
exosome with the Csl4 and the Rrp4 exosome. The
initial rates for degradation of the 30-mer rA are similar
for the Csl4-exosome and the cap-less version, but consid-
erably faster for the Rrp4-exosome (Figure 5A). This in-
dicates that cap proteins can influence recognition and
recruitment of RNA substrates and that this step is
more efficient for the Rrp4 exosome. However, while
RNA degradation for medium and short RNAs is quite
comparable between the Csl4 and Rrp4 exosomes, there is
an interesting difference for long RNA molecules (>24 nt).
The Rrp4 exosome is quite slower for RNAs >24 nt, faster
for RNA between 24 and 13 nt, and then progressively
slower for RNAs <13 nt. This remarkable length depen-
dency is clearly evident in degradation profiles (Figure 1).
The most likely explanation is that long RNAs might still
have contacts with Rrp4, where a more specific binding
site holds them partially back from rapid degradation. In
principle, this could be viewed as molecular friction. When
RNAs are shorter, they loose contact to Rrp4 and degrad-
ation speed is increased. The Csl4 protein and the Rrp4
protein differ in their domain structures. While Csl4
contains
a
Zn-ribbon
domain,
Rrp4
possesses
a
KH-domain, which is a typical RNA-binding domain
and could recognize the oligo-rA. Such a binding could
be responsible for the faster first degradation step, because
it more efficiently sequesters RNAs on the exosome
surface, but may subsequently slow down degradation
until RNAs are too short to maintain simultaneous
contacts at the KH domain and active site.
However, the Rrp4 isoform is more efficient for smaller
RNA species than the Csl4 and capless isoforms. Since
these shorter RNAs cannot form dual contacts with the
active site and outside the caps, the Rrp4 could also influ-
ence the dynamics or other properties of the RNase-PH
ring, for instance to help in loading of RNA into the ring
structure.
SAXS structure of the Rrp4 exosome with endogenously
purified bacterial RNA
To explore the role of the Rrp4 cap in efficiently recruiting
RNAs further, we performed SAXS studies with a
nuclease deficient nine-subunit Rrp4 exosome bound to
RNA:
we
had
noticed
that
this
nuclease
deficient
Rrp4-exosome
(D180A
in
Rrp41)
very
efficiently
co-purifies with E. coli RNA. To determine the kind of
RNA that binds to the exosome we run it on a denaturing
gel together with RNAs with known sizes and could
estimate the size of the RNA to be between 55 and 65 nt
(Supplementary Figure S1). Cloning and sequencing of
bound RNA molecules revealed a set of much shorter in-
homogeneous mixed sequences (Supplementary Table S3).
It is possible that the bound RNAs are a mixture of
various mRNAs from E. coli, although the isolated
RNA is larger than the identified sequences and it is
possible that highly structured RNAs such as tRNAs are
underrepresented due to inefficient amplification and
cloning.
Comparison
of
the
SAXS
structure
of
apo-Rrp4-exosome with the RNA bound complex shows
an increase in the radius of gyration from 39.6 A˚ to 46.8 A˚
when RNA is bound and the corresponding pair distribu-
tion functions contains longer vectors (Figure 6A), likely
because
additional
scattering
elements
from
RNA
protrude from the compact protein core. The resulting
ab initio model of the complex overlaid with the crystal
structure of the apo-complex clearly indicates additional
mass from the bound RNA (Figure 6B and C). This clear
additional mass is distributed in the centre of the cap
structure on top of the neck region but also protrudes
Nucleic Acids Research, 2010, Vol. 38, No. 15
5173
away from the complex. When looking at the overlay with
the crystal structure it appears that the RNA is bound at
the KH and the S1-domains. The SAXS analysis supports
the model that RNA binds near the KH-domain on the
outside of the caps and reveals a low-resolution image of
trapped exosome–RNA complexes.
DISCUSSION
RNA exosomes are large, self-compartmentalized nucle-
ases, implicated in processive, controlled degradation of a
large variety of RNAs. While the archaeal exosome
possesses three phosphorolytic active sites within the com-
partment, the eukaryotic exosomes apparently have lost
this activity but adopted hydrolytic RNase subunits that
are bound at the outside of the evolutionary conserved
core. Nevertheless, recent data suggest that RNA is still
threaded through the eukaryotic core exosome before it is
degraded in ectopic hydrolytic active sites, suggesting that
the core particle retained critical ‘structural’ functions re-
garding RNA degradation such as increased processivity
or controlled RNA degradation (25).
To be able to quantitatively address RNA exosome
activities, we derived a kinetic model for the complex
RNA degradation of the archaeal RNA exosome using
Markov Chain Monte Carlo analysis. The kinetic model
gives a realistic assessment of the velocity of the exosome
and mutant variants during processive degradation of a
rA 30-mer oligonucleotide. The considerable effort we had
to put into the MCMC simulation pays offeventually. We
are now able to derive a realistic joint posterior distribu-
tion of kinetic parameters, enabling us to quantify the
relation of different parameters in either the same or in
distinct exosome mutants. This would have been impos-
sible with a conventional least squares fit of the data,
which produces very unstable parameter estimates (see
Supplementary Data for a comparison), although the
obtained fits are very good (Figure 3D).
With this in hand, we find several interesting and unex-
pected features of RNA degradation activities. First,
kinetic evaluation of RNA degradation of exosomes
needs to include association and dissociation rate con-
stants. Thus the kinetics cannot be treated as strictly
processive, at least for RNA species in the assessed
Figure 6. SAXS structure of the Rrp4 exosome with endogenously purified bacterial RNA. (A) SAXS data of the Rrp4 exosome (green) and the
Rrp4 exosome with RNA (orange) (curves show the scattering intensity I(q) as a function of the scattering angle 2y and X-ray wavelength , where
q = (4sin/y)) and the pair-distribution function describing intramolecular distances; in the presence of RNA longer distances occur and the radius of
gyration increases. (B) Average of 10 independent ab initio models for the apo exosome and the RNA-bound complex superimposed with the crystal
structure. The additional density for the RNA is clearly visible.
5174
Nucleic Acids Research, 2010, Vol. 38, No. 15
length range. This does not necessarily imply that RNA
dissociates and rebinds completely from the exosome.
Longer RNAs may be retained within the neck as well
as cap domains, while binding and dissociating from the
active site in the processing chamber. The association and
dissociation
constants
can
thus
be
understood
as
‘ratcheting’ constants that influence the rate of translation
along the RNA to and from the active site. For short
substrates that are unable to simultaneously bind neck
and active site this connection is lost, the dissociation in-
creases, degradation speeds drop and the exosome changes
from being fast and processive to a slower distributive
enzyme.
Our results also show how neck region and active site
features contribute to exosome activity. Although we
could
not
quantitatively
address
the
importance
of
Arg65 in the neck with the simplified model in hand,
this residue appears to be important for loading RNA
into the processing chamber, but not for efficient degrad-
ation once RNA is bound. This conclusion is derived from
the observation that while the initial degradation is sub-
stantially delayed, the appearance of smaller RNA species
is qualitatively similar to the wild-type Csl4 exosome.
Taken together with the observation that crosslinking
severely reduces processing and the RNase PH ring
needs
to
breath
or
display
some
conformational
dynamics, it is unlikely that RNA is simply threaded
into the processing chamber like a yarn through the eye
of a needle. Rather, we propose that initial RNA binding
includes some lateral entry near the neck.
We are also in the position now to address the influence
of the cap proteins Rrp4 and Csl4. These proteins possess
a variety of domains with unclear function in exosome
activity.
While
eukaryotic
exosomes
have
defined
heterotrimeric caps, the stoichiometry of cap proteins in
archaeal exosomes is not defined in vitro and perhaps
variable in vivo. For the archaeal exosome, the Csl4
capped isoform displays similar degradation kinetics to
that of the capless variant, and the function of this type
of cap remains to be shown. However, the Rrp4 isoform
substantially differs from the other two variants and our
analysis suggests that Rrp4 more efficiently recruits RNA
to the exosome. In fact, RNA from the heterologous
expression in E. coli is very tightly bound to the Rrp4
exosome. It must be noted that the gene coding for
Rrp4 is in the same operon as genes for Rrp41 and
Rrp42, indicating that this cap is perhaps a ‘default’
isoform of the exosome, while the Csl4 cap, located else-
where in the genome, might be differentially regulated.
The cap structures, however, also influence the degrad-
ation of short RNAs. This is to some extent surprising,
since short RNAs (<13 nt) cannot bind to the caps and the
active site at the same time. However, the Rrp4 subunit
more intimately interacts with the RNase PH ring than the
Csl4 protein and might influence also the dynamics of
the RNase PH type ring. Likewise, binding of RNA to
the KH domains, consistent with the lateral density of
RNA in the SAXS models, may position it better for
loading into the processing chamber.
In sum, we present here a robust method to analyse the
complex degradation kinetics of a partially processive
degradation enzyme in a quantitative manner, with esti-
mates of the posterior distribution of the model param-
eters. We applied this analysis to degradation of RNA by
several isoforms and variants of the archaeal exosome and
could reveal a variety of features that are important for
catalytic efficiency. The objective of this manuscript is to
derive a general method that can now be used to unravel
the biochemistry of exosomes in a more quantitative
manner. The method can now form a basis for compre-
hensive analysis of different substrates, other RNA se-
quences, as well as mutants of this system or related
systems such as the eukaryotic exosome.
ACCESSION NUMBERS
3M7N, 3M85.
SUPPLEMENTARY DATA
Supplementary Data are available at NAR Online.
ACKNOWLEDGEMENTS
The authors thank Christian Luginsland for help in
protein purification, Katharina Bu¨ ttner for exosome con-
structs and Katja Lammens and Gregor Witte for helpful
discussions. The authors thank the staffof the European
Synchrotron Radiation Facility (beamline 14–2) and the
Swiss Light Source (beamline PX I) for help with diffrac-
tion data collection and Michal Hammel from the
Advanced Light Source (SIBYLS beamline) for help
with scattering data collection.
FUNDING
Deutsche
Forschungsgemeinschaft
(HO2489/3
and
SFB646);
Center
for
Integrated
Protein
Science
Munich. Funding for open access charge: Deutsche
Forschungsgemeinschaft.
Conflict of interest statement. None declared.
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|
3M89
|
Structure of TubZ-GTP-g-S
|
Plasmid protein TubR uses a distinct mode of HTH-
DNA binding and recruits the prokaryotic tubulin
homolog TubZ to effect DNA partition
Lisheng Ni, Weijun Xu, Muthiah Kumaraswami, and Maria A. Schumacher1
Department of Biochemistry and Molecular Biology, University of Texas, M. D. Anderson Cancer Center, Unit 1000, Houston, TX 77030
Edited by Robert T. Sauer, Massachusetts Institute of Technology, Cambridge, MA, and approved May 11, 2010 (received for review March 22, 2010)
The segregation of plasmid DNA typically requires three elements:
a DNA centromere site, an NTPase, and a centromere-binding
protein. Because of their simplicity, plasmid partition systems
represent tractable models to study the molecular basis of DNA
segregation. Unlike eukaryotes, which utilize the GTPase tubulin
to segregate DNA, the most common plasmid-encoded NTPases
contain Walker-box and actin-like folds. Recently, a plasmid stabi-
lity cassette on Bacillus thuringiensis pBtoxis encoding a putative
FtsZ/tubulin-like NTPase called TubZ and DNA-binding protein
called TubR has been described. How these proteins collaborate
to impart plasmid stability, however, is unknown. Here we show
that the TubR structure consists of an intertwined dimer with a
winged helix-turn-helix (HTH) motif. Strikingly, however, the TubR
recognition helices mediate dimerization, making canonical HTH–
DNA interactions impossible. Mutagenesis data indicate that a
basic patch, encompassing the two wing regions and the N termini
of the recognition helices, mediates DNA binding, which indicates
an unusual HTH–DNA interaction mode in which the N termini of
the recognition helices insert into a single DNA groove and the
wings into adjacent DNA grooves. The TubZ structure shows that
it is as similar structurally to eukaryotic tubulin as it is to bacterial
FtsZ. TubZ forms polymers with guanine nucleotide-binding
characteristics and polymer dynamics similar to tubulin. Finally,
we show that the exposed TubZ C-terminal region interacts with
TubR-DNA, linking the TubR-bound pBtoxis to TubZ polymerization.
The combined data suggest a mechanism for TubZ-polymer pow-
ered plasmid movement.
T
he cytoskeletons of eukaryotic cells are constructed of three
primary elements: actin, tubulin, and intermediate filaments.
Although it had long been presumed that the proteins forming
these elements were absent in prokaryotes, it is now known that
prokaryotes contain structural homologs to all three components.
These prokaryotic proteins appear to carry out distinct functions
compared to their eukaryotic counterparts; however, their roles
are similar enough to indicate a likely common ancestor. The best
known prokaryotic homolog of tubulin is FtsZ. Both FtsZ and
tubulin form long filamentous structures by head to tail associa-
tion in a manner regulated by GTP, which binds between adjacent
subunits (1–4). However, unlike tubulin, FtsZ does not function
in DNA segregation but rather cell division. Specifically, it forms
a cytokinetic ring called the Z ring at midcell, which mediates
septation (5, 6). Recently, however, prokaryotic proteins encoded
on large plasmids harbored in bacilli showing 15–20% sequence
similarity to both FtsZ and tubulin have been identified and
dubbed TubZ (7–12). Studies showed that the Bacillus thuringien-
sis TubZ protein from the pBtoxis plasmid is essential for plasmid
DNA segregation.
DNA segregation of most low copy number plasmids is carried
out by specific partition (par) systems. These systems require only
three elements: a centromere DNA site, a centromere-binding
protein, and a partition NTPase (13, 14). Partition systems have
been classified into two main types on the basis of the kind
of NTPase present (15). Type I systems contain NTPases with
deviant Walker A-type ATPase folds, whereas type II systems uti-
lize actin-like NTPases. Interestingly, both types of NTPases form
polymers in NTP-dependent manners that are implicated to play
a role in plasmid DNA separation (16–19). The recent discovery
of TubZ NTPases has led to the designation of “type III” par sys-
tems (13, 14). The best studied of these systems is that found on
the pBtoxis plasmid in B. thuringiensis. This plasmid stability sys-
tem is represented by an operon encoding two proteins: ORF156
(TubZ) and ORF157 (TubR) (7–9, 11). TubR is a 11.6 kDa DNA-
binding protein that shows no sequence homology to any known
protein. Studies showed that TubR binds a 48-bp centromere con-
taining four repeat sites in the pBtoxis plasmid and also autore-
gulates tubRZ transcription (8, 9). TubZ is a 54.4 kDa protein that
can assemble into filaments in a GTP-dependent manner (12).
Both proteins were found to be required for plasmid stability
(9). However, how the TubR and TubZ proteins work together
to effect pBtoxis plasmid segregation is not known. To gain insight
into the molecular mechanism utilized by these proteins in DNA
segregation, we carried out structural and biochemical studies on
the pBtoxis TubR and TubZ proteins. The TubR structure reveals
that it employs a helix-turn-helix (HTH) motif in a previously un-
described manner to bind DNA. TubZ contains a tubulin/FtsZ
fold but has structural distinctions from these proteins indicating
that it forms distinct protofilaments. TubR binds the flexible
C-terminal region of TubZ, thus attaching the TubZ filament
to the pBtoxis plasmid, providing a mechanism for plasmid move-
ment and, ultimately, segregation.
Results and Discussion
Overall Structure of pBtoxis TubR. The crystal structure of the 107-
residue pBtoxis TubR protein was solved to 2.0-Å resolution by
selenomethionine multiple wavelength anomalous diffraction
(MAD) methods (Table S1). The structure contains two TubR
molecules in the crystallographic asymmetric unit and consists
of residues 6–102 of one subunit and 4–100 of the second subunit,
and has Rwork∕Rfree ¼ 23.8%∕27.0%. The TubR structure forms a
highly
intertwined
dimer
with
dimensions
30 × 30 × 60 Å3
(Fig. 1A). Each TubR subunit has the topology β1-α1-α2-α3-α4-β2-
β3-α5, which is similar to winged HTH motifs found in a number
of DNA-binding proteins in both prokaryotes and eukaryotes
(20). In TubR, α3-α4 forms the HTH motif and the loop between
Author contributions: M.A.S. designed research; L.N., W.X., M.K., and M.A.S. performed
research; M.A.S. analyzed data; and M.A.S. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Freely available online through the PNAS open access option.
Data deposition: The atomic coordinates and structure factor amplitudes for the WT TubR
(C2), WT TubR (I222), TubR(S63W), TubZ, and TubZ-GTP-γ-S structures have been
deposited with the Protein Data Bank, www.pdb.org (PDB ID codes 3M8E, 3M9A,
3M8F, 3M8K, and 3M89).
1To whom correspondence may be addressed. E-mail: maschuma@mdanderson.org.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1003817107/-/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.1003817107
PNAS ∣June 29, 2010 ∣vol. 107 ∣no. 26 ∣11763–11768
BIOCHEMISTRY
β2 and β3, the wing. Indeed, each TubR subunit shows the stron-
gest structural similarity to members of the ArsR family of
prokaryotic winged helix transcription regulators, in particular
the Staphylococcus aureus CzrA protein (21, 22). Superposition
of one subunit of TubR onto that of CzrA results in a rmsd of
2.7 Å. This similarity includes the core regions of the winged
HTH, because the loops and N-terminal regions of the proteins
are structurally distinct. For example, TubR contains a β-strand in
its N-terminal region compared to a long helix in the CzrA struc-
ture (Fig. 1B). This structural similarity initially suggested that
TubR may be a member of the ArsR family of proteins. However,
the arrangement of the TubR dimer was found to be strikingly
different from the dimer organization exhibited by the ArsR
proteins (Fig. 1C).
ArsR family members are involved in metal-regulated tran-
scription processes whereby they act as repressors in their apo
forms and are induced off their DNA sites upon metal binding
(22). The specific dimer structures of the ArsR proteins are cri-
tical for creation of their metal-binding motifs. Not only does
TubR form a very different dimer from the ArsR proteins, it also
does not harbor any of their metal-binding signatures nor does it
contain any other characterized metal-binding motif. Consistent
with this, we find that the addition of metals has no effect on TubR
DNA binding. Dimerization of the ArsR proteins is imparted by
residues from the N-terminal regions, α1 and α5, which, impor-
tantly, leaves its recognition helices exposed for DNA interaction
(22). By contrast, the TubR dimer is formed primarily by contacts
between its twofold related “recognition helices” α4 and α4·. This
interaction results in a near complete burial of these helices, leav-
ing only the N-terminal residues exposed to solvent. Whereas the
α4 and α4· interaction creates the dimer core, the dimer is further
stabilized by interactions between the twofold related β1 strands,
which swap to form an antiparallel β-sheet. Residues from α1 and
α5 interact with β1 to further seal the top of the dimer. The dimer
interface formed by these interactions is predominantly hydropho-
bic and buries a large 1;200 Å2 of subunit surface from solvent.
TubR Forms a “Recognition Helix Dimer”: Implications for DNA Bind-
ing. Gel filtration studies on TubR confirmed that it is a dimer in
solution. However, the finding from the structure that the TubR
α4 recognition helices are buried in the dimer core has important
implications in terms of its DNA-binding mechanism. Indeed, it
suggests that, although TubR contains a structurally canonical
HTH, it is not utilized for DNA binding in a manner typical
of HTH proteins. A second crystal form (I222) of TubR, which
was solved to 2.5-Å resolution, revealed the same TubR dimer.
The presence of the identical dimer in two different crystal forms
and its large buried surface area supports that the dimer observed
in the crystal structures is physiologically relevant. However, to
test this, we mutated residues within the recognition helices that
the structure indicates are critical for dimerization and assayed
the ability of the mutant proteins to dimerize via gel filtration.
Specifically, we mutated Ser-63 and Ala-67 individually to tryp-
tophan and arginine.
The structure shows that residues occupying positions 63 and
67 must be small and largely hydrophobic to permit the proper
packing of the α4 helices in the dimer (Fig. S1A). Hence, the in-
troduction of the bulky side chain of tryptophan and, in particu-
lar, the large as well as charged side chain of arginine would be
predicted to be highly disruptive to dimerization. Gel filtration
analyses on purified mutant proteins clearly showed that the ar-
ginine mutants exist primarily as monomers in solution (>80%),
whereas the tryptophan mutants were able to maintain the di-
meric state (Fig. 2A and Fig. S1B). However, all mutant proteins
showed reduced or loss of DNA-binding activity as ascertained by
fluorescence polarization (FP) studies, which examined TubR
protein binding to its centromere site (Fig. 2B and Fig. S1 C
and D) (9). The fact that the monomeric mutants were severely
impaired in DNA binding was not surprising. However, the
finding that the tryptophan mutants, which were largely dimeric,
displayed reduced DNA-binding activity suggested that their oli-
gomer structures might be altered. To address this issue, the struc-
ture of S63W TubR was solved to 2.8-Å resolution, resulting in
Rwork∕Rfree values of 20.4%∕26.6% (Table S1). The subunit struc-
ture of S63W TubR is essentially identical to that of WT TubR as
revealed by their superimposition (rmsd of 1.2 Å) (Fig. 2 C and
D). However, this single subunit overlay shows that the S63W
TubR dimer, although the same as the WT in general arrange-
ment, is forced into a more open oligomer conformation in which
one subunit is rotated 20° away from its dimer mate compared to
WT. This rotation is required to accommodate the bulky S63W
side chains (Fig. 2D). The N-terminal β1–β1′-sheet interaction
appears to play a key role in holding the TubR subunits together.
In addition, the tight stacking of the twofold related Trp63 indole
groups (3.5 Å) provides a compensatory interaction that, com-
bined with the β1–β1′ interaction, apparently permits retention
of the dimer state, indicating why the TubR tryptophan mutants
were able to maintain the dimer state, albeit an altered dimer
state relative to WT (Fig. 2 C and D). By contrast, the TubR
arginine mutations, which introduced both bulk and charge with-
in the predominantly hydrophobic dimer interface, were highly
destabilizing for dimerization. In addition, the finding that the
S63W TubR mutant forms an altered dimer explains the severe
effect on DNA binding because a correct dimer orientation is
likely essential for binding to its palindromic DNA sites (9).
TubR-DNA Model. Because all but the N-terminal residues of the
TubR recognition helices are buried in the dimer interface, TubR
must use a different mode of DNA binding than the ArsR or other
HTH containing proteins (23). Examination of surface electro-
statics of TubR reveals that one face of the protein is electro-
negative, whereas the other is strongly electropositive (Fig. 3A).
Notably, the positive region is composed of one large and contig-
uous basic patch. Basic residues in this region correspond to Arg-
74, Arg-77, and Lys-79, in the wing and Lys-43, which is on α3, the
helix preceding α4 in the HTH motif. These residues were mutated
singly to alanine to examine their roles in DNA binding (Fig. 3B).
FP experiments showed that mutation of the basic wing residues
resulted in either a complete (R74A and R77A) or nearly com-
plete (K79A) abrogation of DNA binding, indicating that the
wings play a major role in TubR DNA binding. Residue Lys-43
is surface exposed and located at the center of the basic region
on the TubR dimer (Fig. 3A). The K43A mutant also showed
no binding to TubR, supporting the notion that the continuous
basic patch of TubR represents its DNA-binding surface.
To gain insight into the structural mechanism of DNA binding,
a DNA duplex was docked onto the basic patch of the TubR di-
Fig. 1.
B. thuringiensis pBtoxis TubR structure. (A) One TubR subunit is red
and the other cyan. Secondary structural elements and N and C termini are
labeled. (B) Superimposition of one subunit of TubR (Red) onto a S. aureus
CzrA subunit (Yellow). Regions with different structures are labeled. (C) Same
superimposition as B showing the location of the other subunit in the TubR
and CzrA dimers after one subunit is overlaid. A–C are in the same orienta-
tion to highlight differences. Figs. 1 A–C, 2 C and D, 3 A, C, and E, 4B, and 5B
were made by using PyMOL (31).
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mer by using the location of the mutations that affected DNA
binding as a guide (Fig. 3C). This model revealed that the wings
are positioned to interact with successive minor grooves, with
either the bases or the phosphate backbone depending on the
ability of the DNA to deform. In the model TubR interacts with
a minimum of 14 bp of DNA. However, the centromere bound by
Fig. 2.
The TubR “recognition helix” dimer. (A) Gel fil-
tration studies on TubR mutants S63W and S63R show-
ing that the S63W mutant remains dimeric whereas
S63R is >80% monomer. (B) Fluorescence polarization
studies examining the ability of WT TubR, S63R, and
S63W TubR mutants to bind iteronic DNA. Fluorescence
polarization units (millipolarization) and TubR concen-
tration (nM) are along the y and x axes, respectively.
The Kd of WT TubR for the centromere DNA is
8 2 nm. (C) Superimposition of WT TubR (Green) onto
the TubR S63W mutant structure (Tan). (D) Close-up of
the site of the S63W mutation in the expanded TubR
S63W dimer showing stacking interactions between
the twofold related tryptophans.
Fig. 3.
TubR-DNA binding. (A) Electrostatic surface potential of the TubR dimer. Blue and red represent electropositive and electronegative regions, respec-
tively. (Left) The electronegative side of the TubR dimer, and the side on the right is the electropositive side. Labeled on the left side are the locations of the
mutated residues. (B) FP binding isotherms showing the DNA binding of WT TubR and the K43A, R74A, R77A, and K79A mutants. Fluorescence polarization
units (millipolarization) and TubR concentration (nM) are along the y and x axes, respectively. (C) TubR-DNA model showing TubR electrostatic potential.
(D) Stoichiometry of TubR (subunit) binding showing titration curve of TubR into the 48-mer iteron resulting in a molar ratio of TubR subunit to DNA of
eight (or four) dimers. (E) Left: Ribbon diagram of the TubR-DNA model with the recognition helices colored yellow. Right: Ribbon diagram showing a canonical
HTH–DNA interaction (the λ repressor-DNA complex) with the recognition helices colored yellow (32).
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TubR consists of four 12-bp sites with the consensus T(T/A)(T/A)
(C/A)(G/A)GTTTA(A/C)(A/C) (9). Thus, we used FP to ascer-
tain the binding stoichiometry of TubR for its 48-bp centromere.
As shown in Fig. 3D, eight TubR subunits or four TubR dimers
bind the 48-mer centromere, consistent with a dimer of TubR
binding each palindrome. Thus, either TubR distorts its DNA
or the TubR dimers bind with some degree of overlap on their
DNA sites perhaps imparting cooperativity, as observed in other
centromere-binding protein–DNA interactions (13, 14). In addi-
tion to the insertion of the wings, a striking outcome of the mod-
eling was the finding that the N termini of the recognition helices,
which interact with each other in a parallel, coiled-coil-like man-
ner, are in position to insert into a single major groove. Structures
of HTH proteins bound to DNA have thus far shown that the
recognition helices insert singly into successive major grooves
by using residues in the first few turns or the central portion
of the recognition helix to contact the DNA bases (Fig. 3E). Thus,
in the TubR-DNA model, the HTH–DNA interaction is drama-
tically different from any displayed previously by a HTH protein.
TubZ Binds TubR-DNA. A characteristic feature exhibited by
partition centromere-binding proteins is the ability to bind their
partner NTPase (13, 14). To determine if TubR binds TubZ, we
utilized a FP assay and found that full length (FL) TubZ bound
avidly to the TubR-centromere complex (Fig. 4A). However,
unlike other partition systems in which the NTPase must be com-
plexed with nucleotide to bind its centromere-binding protein,
the interaction of TubR with TubZ did not require the presence
of GTP-Mg2þ. Previous studies have shown that the C-terminal
regions of tubulin and FtsZ mediate key binding events with their
target proteins (24–26). We noted that the terminal region of
TubZ, consisting of residues 407–484, is the most divergent region
between TubZ proteins and between TubZ and tubulin/FtsZ pro-
teins, suggesting that it may be similarly utilized and bind TubR.
To test this hypothesis, we constructed various TubZ truncations,
TubZ(1-407), TubZ(1-442), TubZ(1-460), and TubZ(1-470), and
examined the ability of each protein to bind TubR-DNA. TubZ
(1-407) showed no binding to TubR, whereas the remaining trun-
cation mutants bound weakly to TubR-DNA (Fig. 4A). Thus, the
data indicate that the last 14 amino acids of TubZ are critical for
the ability of TubZ to form a tight interaction with TubR but that
residues 408–470 also play an important role in this interaction.
These data demonstrate that TubR acts as a partition partner for
TubZ, linking it to pBtoxis plasmid DNA. Although TubZ has
been shown to form polymers in a GTP-dependent manner,
the TubZ protein displays limited sequence similarity to tubu-
lin/FtsZ, suggesting potential differences in TubZ and tubulin/
FtsZ structures (7–9). To gain insight into TubZ function, we next
determined structures of B. thuringiensis pBtoxis TubZ.
Structure of TubZ. Crystallization of FLTubZ was not successful, in
either its apo form or bound to guanine nucleotides. We noted that
FL TubZ degraded over time whereby C-terminal residues were
proteolyzed. Therefore, truncated TubZ proteins were utilized
in crystallization trials, and crystals were obtained of apo TubZ
(1-428) and the structure solved by MAD (Table S2). The model
consists of residues 1–79 and 91–404 and has an Rwork∕Rfree of
21.4∕24.9% to 2.3-Å resolution (Fig. 4B). No discernible oligomer-
ization of apo TubZ(1-428) was observed in the crystal packing,
and gel filtration analyses confirmed that it is monomeric
(Fig. S2). The overall TubZ structure can be divided into two main
domains: an N domain (residues 25–235) and a C domain (resi-
dues 258–377). These domains are connected by a long, core helix,
H7. The TubZ N domain has a Rossman fold and consists of six
parallel β-strands with topology 3-2-1-4-5-6. The resulting β-sheet
is sandwiched by five α-helices, with two helices on one side and
three on the other. The C domain consists of four β-strands with
the topology 1-4-2-3. The C-domain β-strands are arranged nearly
perpendicular to those in the N domain. In addition to these main
protein domains, there are two helices: one at the N terminus, H0,
and a long helix at the C terminus, H11 (Fig. 4B). Database
searches showed that TubZ indeed belongs to the tubulin/FtsZ fa-
mily of proteins and is similar to both eukaryotic and prokaryotic
members of the family; TubZ can be optimally superimposed with
rmsds of 3.4 Å onto both bovine α tubulin and Pseudomonas
aeruginosa FtsZ (1, 5). Whereas the two-domain architectures
of tubulin, FtsZ, and TubZ are similar in overall structure, the
extreme N- and C-terminal regions of these proteins are very di-
vergent (Fig. S3 A–D).
N-Terminal and C-Terminal Differences in TubZ, FtsZ, and Tubulin: Im-
plications for Polymer Formation and Target Protein Binding. Tubulin
proteins do not contain significant N-terminal extensions,
whereas FtsZ proteins from different organisms show structural
variability within their N-terminal regions. For instance, in the
Escherichia coli FtsZ structure the N-terminal residues are disor-
dered, whereas Methanococcus jannaschii FtsZ has an extra
N-terminal helix, H0, which is flexibly attached to the body of
the protein and has been captured in multiple orientations (5).
Although H0 is not conserved in FtsZ proteins, one M. jannaschii
FtsZ structure revealed a semicontinuous polymer in the crystal,
thought to closely represent in vivo protofilaments, which utilizes
H0 in subunit-subunit contacts (4). This finding suggests that
the flexibly attached H0 is stabilized in a specific orientation
by protofilament formation, at least in the M. jannaschii protein.
The TubZ H0 helix extends in the opposite direction compared to
that of the protofilament stabilized FtsZ H0 helix. Moreover, in
TubZ, H0 is not flexibly attached to the N domain but is tightly
anchored to the C domain through numerous interactions with
the core helix and C-domain residues. The large number of inter-
actions involving H0, and the fact that it covers what would other-
wise be a surface exposed hydrophobic patch, indicate that the
TubZ H0 does not undergo conformational changes during
protofilament formation and is important for the general fold
of TubZ (Fig. S3 A and C).
Data suggest that FtsZ and tubulin form protofilaments with
similar longitudinal contacts (4). However, the TubZ structure
reveals key differences, primarily in its C-domain and C-terminal
regions, which suggest that it forms protofilaments distinct from
those formed by tubulin/FtsZ. A notable difference is the struc-
ture of loop 7 (L7). This loop inserts into the adjacent subunit
providing the key catalytic residues required for GTP hydrolysis.
In tubulin/FtsZ proteins, L7 has the consensus GXXNXDXAD.
In TubZ, the loop is very divergent in conformation compared
Fig. 4.
TubZ interacts with TubR-DNA and contains a tubulin/FtsZ fold. (A) FP
assay measuring binding of FL TubZ, TubZ(1-470), TubZ(1-460), TubZ(1-442),
and TubZ(1-407) to TubR-DNA. Below is the control (TubZ titrated into DNA
alone). Millipolarization units and TubZ concentration (nM) are along the y
and x axes, respectively. (B) TubZ(1-428) structure. The N domain or GTP-bind-
ing domain is colored salmon and the C domain purple. The interdomain
helix, H7, is red. TubZ also contains an N-terminal helix, H0 (Yellow), and
a C-terminal helix, H11 (White).
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to that in FtsZ/tubulin and consists of the sequence 256-
DNVTYDPSD-266. In addition to the N-terminal region, the
extreme C-terminal extensions of tubulin, FtsZ, and TubZ are
structurally divergent (Fig. S3B). In FtsZ, the C-terminal region
forms a small, two-stranded β-sheet and continues into an ex-
tended region that is involved in binding adaptor proteins such
as FtsA and ZipA (6, 25). By contrast, the C-terminal regions
of tubulin proteins consist of a two-helix bundle followed by an
extended region. Like FtsZ, however, these regions interact with
numerous target proteins such as the microtubule-associated pro-
teins (MAPs) (24). Consistent with this, the C-terminal regions of
tubulin have been shown to face the outside of the microtubule. A
characteristic feature of the extreme C-terminal extensions of
tubulin proteins is their highly acidic nature (3). This acidic region
has been shown to be critical for binding to several MAPs that
harbor a substantial basic character, such as tau, MAP2, and
MAP4 (24, 27).
The TubZ C-terminal region is also helical, but it contains a
single, long helix. Notably, the TubZ-tubulin overlay shows that
the long C-terminal helix of TubZ would dramatically clash with
the adjacent subunit in a polymer, providing support for the no-
tion that TubZ forms protofilaments different from tubulin/FtsZ
(Figs. S3B and S4). Interestingly, and in contrast to tubulin pro-
teins, the flexible C-terminal region of TubZ that follows H11 is
highly basic, in particular the last 14 residues. We have shown that
these residues play a central role in TubR binding (Fig. 4A). TubR
uses its electropositive face for DNA binding, leaving exposed its
opposite face for TubZ interaction. Notably, this exposed face is
strongly electronegative and hence would complement the basic
C-terminal tail of TubZ (Fig. 3A).
Tubulin/FtsZ protofilaments combine to form higher-order
structures. In tubulin, the protofilaments interact in a parallel
manner to form microtubules. Central to microtubule formation
are lateral contacts between protofilaments from the so-called M
loop, between H10 and S9. In tubulin, this loop is composed of 13
residues (1–3). The corresponding loop is much shorter in FtsZ
proteins, consistent with the fact that FtsZ does not form tubulin
microtubule-like structures (5, 28–30). In TubZ, the M loop is
even shorter than in FtsZ, spanning only four residues. In fact,
the TubZ/tubulin overlay shows that the side of the molecule con-
taining the M loop is the most divergent between these proteins.
These combined findings suggest that TubZ not only forms pro-
tofilaments with distinct longitudinal contacts compared to FtsZ
and tubulin, but it also does not form tubulin-like microtubule
structures.
TubZ Interactions with Guanine Nucleotides. Consistent with TubZ
being a member of the tubulin/FtsZ family, our isothermal titra-
tion calorimetry (ITC) studies showed that TubZ binds guanine
nucleotides with high affinity; Kds for GTP-γ-S and GDP were
∼0.69 and 26 μM, respectively (Fig. 5A). We next determined
the structure of the TubZ-GTP-γ-S complex by soaking GTP-γ-S
into preformed TubZ(1-428) crystals. The TubZ-GTP-γ-S struc-
ture contains TubZ residues 1–79 and 91–404, one GTP-γ-S,
and has Rwork∕Rfree ¼ 21.8%∕25.5% (Fig. 5B and Table S2).
The structure shows that TubZ binds GTP-γ-S in the same GTP
binding pocket as tubulin/FtsZ (1–5). Comparison of the apo
and GTP-γ-S bound TubZ structures indicated that, like FtsZ,
guanine nucleotide binding does not lead to significant conforma-
tional changes (5). The phosphate binding pocket is formed by
two of the most highly conserved regions between TubZ and
tubulin/FtsZ called loops 1 and 4 (L1 and L4) (1–3). L1 contacts
the GTP-γ-S α- and β-phosphate groups via the Gln-32 and Lys-33
amide nitrogens. The L1 region of FtsZ and tubulin contain the
sequences GQ(A/G)G and GQCG, respectively, whereas in TubZ
the motif is 31-GQKG-34. However, the alanine/glycine and
cysteine residues in FtsZ and tubulin do not contact the bound
nucleotide; the TubZ Lys-33 side chain makes stacking interac-
tions with the guanine base (Fig. 5B). L4 represents the so-called
signature motif [GGGTG(T/S)G], which serves as an identifier of
tubulin/FtsZ family members. Like FtsZ and tubulin, the L4
region of TubZ-GTP-γ-S makes phosphate interactions via its
glycine amide nitrogens. Whereas L1 and L4 residues of the N
domain mediate phosphate contacts, the GTP-γ-S guanine moiety
is specified from residues in the core helix, H5, and C domain. In
this regard, an important motif is loop 6 (L6). In FtsZ and tubulin,
L6 has the consensus (F/Y)XXX(N/D) and the conserved (F/Y)
residue functions in guanine base stacking. This region in TubZ,
236-WKXXXN-241, is in an altered conformation compared to
FtsZ and tubulin structures. Despite the presence of the trypto-
phan, which might be expected to interact with the guanine,
the side chain of Lys-237 instead stacks with the guanine ring.
Hence, in the TubZ-GTP-γ-S structure, the guanine base does
not interact with aromatic residues as in tubulin/FtsZ but is sand-
wiched between the aliphatic portions of two lysine side chains,
Lys-33 and Lys-237. Finally, two asparagine residues, Asn-213
and Asn-241, from L6 effectively read the guanine N2/N3 and
N1/O6 atoms, respectively, providing high specificity in TubZ’s in-
teraction with guanine nucleotides.
pBtoxis DNA Segregation: TubR Plasmid Partition Model. Our data
show that TubR binds to the flexible, C-terminal, basic region
of TubZ. The flexibility and location of the TubZ C-terminal
extension suggest that it is not required for polymerization and
thus may be exposed on the surface of TubZ filaments. Indeed,
negative stain EM images show that TubZ(1-407) forms polymers
in a GTP-dependent manner similar to the FL protein (Fig. S5).
Recent data suggesting that TubZ filaments are stabilized by
Fig. 5.
TubZ-guanine nucleotide interactions. (A) ITC binding isotherms
showing TubZ-GDP (Left) and TubZ-GTP-γ-S interaction (Right). (B) Left:
Overall structure of the TubZ-GTP-γ-S complex. β-strands are colored magen-
ta and helices cyan, and the GTP-γ-S molecule is shown as cpk. Right: Close-up
view of the GTP binding pocket with the initial Fo-Fc electron density map
(Blue Mesh), contoured at 4.5σ, and calculated before the GTP-γ-S was
included in refinement.
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the presence of a GTP cap and undergo treadmilling are consis-
tent with the notion that TubZ displays tubulin-like polymer dy-
namics (12). Thus, on the basis of the combined data, we suggest
a model for TubR/TubZ mediated pBtoxis plasmid segregation
shown in Fig. 6. In this model, multiple TubR dimers first bind
to the iteronic DNA on the pBtoxis plasmid leading to the crea-
tion of a high local concentration of TubR, which can recruit a
TubZ polymer, likely by interactions between the acidic TubR
dimer face and the basic C-terminal TubZ region. Importantly,
this interaction serves to attach the pBtoxis plasmid to the TubZ
polymer, which undergoes treadmilling, adding subunits at the þ
end and losing subunits at the −end. The bound TubR-pBtoxis
can be handed off from the −end to the molecules in the growing
þ end, leading to the transport of the pBtoxis plasmid to the cell
pole. Interestingly, it has been shown that once TubZ polymers
reach and interact with the cell pole, they bend around the curved
pole and continue growing in the other direction (7). The force of
the interaction with the membrane likely causes the release of
TubR-pBtoxis, the net result being transport of pBtoxis to the cell
pole. Of course, this model is simplified and many questions re-
main. For example, how directionality is achieved and how the
replicated plasmids are driven to opposite cell poles is not clear.
However, given the large size of the pBtoxis plasmid (8), it may be
that only one TubR-pBtoxis “tram” can be bound at once by the
rapidly treadmilling TubZ polymer and that, once one such a tram
is unloaded after reaching the cell pole, another engages when the
now reversed polymer treadmills toward the opposite cell pole.
Materials and Methods Summary
Detailed methods are provided in SI Materials and Methods.
Briefly, the tubR and tubZ genes were codon optimized (for
E. coli expression), subcloned into pET15b, expressed, and pur-
ified. WT TubR crystals were grown with NaCl and phosphate.
TubR S63W was crystallized with PEG and ethylene glycol and
TubZ with sodium formate. Detailed assay conditions for FP,
ITC, electron microscopy, and gel filtration are provided in
SI Materials and Methods.
ACKNOWLEDGMENTS. This work was supported by the Burroughs Wellcome
Career Development Award 992863 and National Institutes of Health Grant
GM074815 (to M.A.S.).
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(2009)
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Fig. 6.
pBtoxis DNA partition model. In the first step, TubR, which is bound
to its centromere on one of the replicated pBtoxis plasmids, contacts the TubZ
C-terminal region (indicated by lines pointing from the TubZ “circles”) in a
treadmilling TubZ polymer. TubZ subunits are lost from the −end and are
added to the þ end. TubR is pulled along the growing polymer by its
TubR-TubZ interaction until it reaches the cell pole and is knocked off when
it comes into contact with the membrane at the cell pole. TubZ reverses
direction and may pick up the other TubR-pBtoxis complex and deliver it
similarly to the opposite cell pole.
11768
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Ni et al.
|
3M8B
|
Crystal structure of spin-labeled BtuB V10R1 in the apo state
|
Conformational Exchange in a Membrane Transport Protein Is Altered
in Protein Crystals
Daniel M. Freed,† Peter S. Horanyi,‡ Michael C. Wiener,‡ and David S. Cafiso†‡*
†Departments of Chemistry and ‡Molecular Physiology and Biological Physics, and the Biophysics Program, University of Virginia,
Charlottesville, Virginia
ABSTRACT
Successful macromolecular crystallography requires solution conditions that may alter the conformational
sampling of a macromolecule. Here, site-directed spin labeling is used to examine a conformational equilibrium within BtuB,
the Escherichia coli outer membrane transporter for vitamin B12. Electron paramagnetic resonance (EPR) spectra from
a spin label placed within the N-terminal energy coupling motif (Ton box) of BtuB indicate that this segment is in equilibrium
between folded and unfolded forms. In bilayers, substrate binding shifts this equilibrium toward the unfolded form; however,
EPR spectra from this same spin-labeled mutant indicate that this unfolding transition is blocked in protein crystals. Moreover,
crystal structures of this spin-labeled mutant are consistent with the EPR result. When the free energy difference between
substates is estimated from the EPR spectra, the crystal environment is found to alter this energy by 3 kcal/mol when compared
to the bilayer state. Approximately half of this energy change is due to solutes or osmolytes in the crystallization buffer, and the
remainder is contributed by the crystal lattice. These data provide a quantitative measure of how a conformational equilibrium in
BtuB is modified in the crystal environment, and suggest that more-compact, less-hydrated substates will be favored in protein
crystals.
INTRODUCTION
Proteins are dynamic and structurally heterogeneous. They
exhibit collective and uncoupled motions over a wide range
of timescales (1,2) and they may assume numerous discrete
structural substates that are in equilibrium. This motion and
sampling of structural states is important and appears to be
critical to enzyme activity and allosteric regulation (3–5). In
protein crystals, dynamics and structural heterogeneity are
present, and at sufficiently high resolution, may be well
represented by the use of multiple conformations of the
side-chain (6) and backbone atoms (7).
There are indications that protein crystallization and the
conditions used for crystallization may alter protein dy-
namics and conformational sampling. Molecular dynamics
simulations of proteins in a crystal lattice have been
performed for both soluble (8–10), and membrane (11)
proteins. These computational efforts suggest that confor-
mational sampling may be altered by protein crystallization.
In addition, experiments employing precipitants or osmo-
lytes similar to those used in protein crystallization demon-
strate that these solutes may have a significant effect on
exchange between long-lived conformational substates; for
example, osmolytes have been found to alter conformational
substates involved in enzymatic activity (12,13) and ion
conduction (14).
In BtuB, the outer membrane Escherichia coli vitamin
B12 (CNCbl) transporter, an electron paramagnetic reso-
nance
(EPR)-based
method
termed
site-directed
spin
labeling (SDSL), has been used to investigate the dynamics
and structural transitions in an N-terminal energy coupling
segment termed the Ton box (15). The Ton box couples
BtuB to the inner membrane protein TonB, which provides
energy for transport (16–18). SDSL provides strong
evidence that the Ton box undergoes a vitamin B12-depen-
dent unfolding (19,20), as depicted in Fig. 1. This event
moves the Ton box 20–30 A˚ into the periplasmic space,
where it may act as a trigger to initiate BtuB-TonB interac-
tions (21). In contrast, the Ton box remains folded within the
transporter in crystal structures of BtuB either in the pres-
ence or absence of substrate. While there are small shifts
in the conformation of the Ton box upon substrate binding,
no evidence is seen for the substrate-dependent unfolding
observed spectroscopically (22).
The discrepancy between the spectroscopic and crystallo-
graphic result might have several origins. EPR spectroscopy
of membrane-associated BtuB demonstrates that there is an
equilibrium between folded and unfolded substates of the
Ton box, and that this equilibrium is shifted toward the
more folded state by the osmolytes used in the BtuB crystal-
lization (23–25). Osmolytes, such as polyethylene glycols
(PEGs), are believed to be excluded from hydrated protein
surfaces (26,27), thereby raising the energy of the protein
and reducing its solubility. As a result, the presence of os-
molytes will favor conformational substates that are less
hydrated (12,28,29). The packing of the protein within the
crystal lattice might also account for the difference between
the spectroscopic and crystallographic result. Although
protein-protein contacts within the unit cell should not
interfere sterically with the unfolding of the Ton box, the
contributions that the lattice might make to the Ton box
equilibrium are not known.
Submitted April 26, 2010, and accepted for publication June 14, 2010.
*Correspondence: cafiso@virginia.edu
Editor: David D. Thomas.
2010 by the Biophysical Society
0006-3495/10/09/1604/7
$2.00
doi: 10.1016/j.bpj.2010.06.026
1604
Biophysical Journal
Volume 99
September 2010
1604–1610
To determine how the Ton box equilibrium is modified
within the protein crystal compared to the bilayer state,
we generated a spin-labeled mutant of BtuB where the nitro-
xide side chain R1 (Fig. 1 c) is incorporated into the Ton box
at position 10. EPR spectroscopy was then carried out in
parallel with x-ray diffraction and structure determination
of this labeled BtuB mutant (V10R1) using the same protein
crystals. The EPR spectra obtained from the protein crystal
indicate that the substrate-dependent Ton box transition is
blocked. This spectroscopic result is consistent with crystal
structures of this BtuB mutant, which indicate that the Ton
box remains folded and the R1 side chain buried with or
without substrate. By comparing EPR spectra of BtuB-
V10R1 in bilayers with spectra from the protein crystal,
we estimate the free energy change induced by the crystal
environment on this conformational equilibrium, and dissect
the energetic contributions made by solute and the crystal
lattice.
MATERIALS AND METHODS
Mutagenesis, expression, and purification
The V10C mutation was introduced into btuB using a QuikChange site-
directed mutagenesis kit (Stratagene, La Jolla, CA), and was subsequently
verified by nucleotide sequencing. Expression and purification of BtuB for
the formation of protein crystals was performed as described previously
(22), and BtuB was reconstituted into vesicle bilayers by following a
procedure described elsewhere (30).
Spin labeling
For spin labeling, the first round of purification was paused before initiation
of the salt gradient. The Q-Sepharose slurry (Amersham, Piscataway, NJ)
bound with BtuB was transferred to a conical tube and reacted with
1 mL of 45 mM S-(1-oxy-2,2,5,6-tetramethylpyrroline-3-methyl)methane-
thiosulfonate (MTSL; Toronto Research Chemicals, Ontario, Canada) for
4 h at room temperature.
Crystallization and crystallographic data
collection
Purified BtuB (11 mg/mL in 30 mM Tris pH 8.0, 20 mM C8E4) was crys-
tallized by mixing 1 mL of BtuB and 1 mL of reservoir buffer in an EasyXtal
hanging-drop tray (Qiagen, Germantown, MD), containing 200 mL of total
reservoir buffer for each crystallization condition, and followed by incuba-
tion at 290 K. The reservoir buffer consisted of 200–550 mM magnesium
acetate, 5.0–7.5% PEG3350, and 20 mM Bis Tris at pH 6.6. Crystals
were visible after 1–2 days, and grew to ~200 mm in the longest dimension
after 1–2 weeks. For crystals to be incubated with substrate, 1 mL of soaking
buffer (150 mM calcium chloride, 2.5% PEG3350, 20 mM Bis Tris at
pH 6.6, and 10 mM C8E4) was added to each well, followed by incubation
overnight. The crystals were subsequently transferred into soaking buffer
containing 1 mM cyanocobalamin and 20% glycerol, and allowed to incu-
bate for at least 4 h. For x-ray diffraction, apo and Ca2þB12-soaked crystals
were transferred to cryo-buffer (150 mM magnesium acetate or calcium
chloride, 2.5% PEG3350, 20 mM Bis Tris at pH 6.6, 10 mM C8E4, and
20% glycerol) for 1–2 min before loop mounting and cryocooling by inser-
tion into liquid nitrogen. Diffraction data were taken at 90 K at the 22ID
beamline at the Advanced Photon Source (Argonne National Laboratory,
Argonne, IL). See Table 1 for more details.
Structure determination
Indexing, integration, and scaling of the diffraction data was performed
using HKL2000 (31). The structures were solved with PHASER (32)
maximum likelihood molecular replacement method, using PDB deposi-
tions 1NQE and 1NQH (22) as search models for the apo- and Ca2þB12-
bound data, respectively. To reduce model bias, V10 was deleted from
the apo-search model, and the entire Ton box was deleted from the Ca2þ-
CNCbl-bound search model. Model building was done in COOT (33),
and unrestrained TLS (34) refinement was performed using REFMAC
(35) and PHENIX was used to refine the occupancy of the spin label
(36). The spin-labeled residue V10R1 was manually built in COOT. Anom-
alous difference Fourier maps were calculated to accurately position bound
cobalt and calcium using Sfall and fast-Fourier transform (37). Completed
structures were evaluated and validated with MolProbity (38).
Electron paramagnetic resonance
Apo- and Ca2þ B12-soaked crystals were incubated for at least 4 h in
cryobuffer and soaking buffer (with 1 mM cyanocobalamin and 20% glyc-
erol), respectively. Crystals were then transferred to a 0.60 ID 0.84 OD
round capillary with a syringe (Hamilton Syringe, Bonaduz, Switzerland)
for EPR spectroscopy, which was performed on an X-band EMX spectrom-
eter (Bruker Biospin, Billerica, MA) equipped with a dielectric resonator.
All EPR spectra were recorded with a 100 G magnetic field sweep at
2.0 mW incident power at a temperature of 298 K. The phasing, normaliza-
tion, and subtraction of EPR spectra was performed using LabVIEW soft-
ware provided by Dr. Christian Altenbach (University of California,
Los Angeles, California).
FIGURE 1
BtuB in the (a) apo form where the Ton box position is
highlighted (PDB ID: 1NQE). (b) Vitamin B12 bound form of BtuB
showing the state of the Ton box as determined by EPR spectra and pulse
EPR distance measurements (based upon PDB ID 1NQH and spectroscopic
restraints obtained for the Ton box in bilayers (21)). This unfolding event
places the Ton box as much as 30 A˚ into the periplasmic space. (c) The
structure of the spin-labeled R1 side chain and dihedral angles that define
the rotamers of R1.
Biophysical Journal 99(5) 1604–1610
Membrane Protein Conformational Exchange
1605
To determine the free energies and free energy changes between Ton box
states, the population of each Ton box conformation was determined by
spectral subtraction and quantitation of the spectral components as
described previously (24). For BtuB V10R1, the EPR spectra are linear
combinations of the spectra resulting from the folded and unfolded Ton
box conformations. As a result, the fraction of spins in each population
may be estimated by determining the contribution that each conformation
makes to the total spectrum. The label at position 10 was chosen for these
measurements, because EPR spectra for V10R1 yield dramatically different
lineshapes for the folded and unfolded forms of the Ton box. As a result, it
is easy to simulate both the folded and unfolded lineshape. In this case, the
mobile lineshape was simulated (using Redfield theory (39)) and subtracted
from the composite spectrum until a spectrum corresponding to the purely
folded Ton box conformation was obtained. Double integration of the first
derivative EPR spectra yields numbers that are proportional to spin number
and was used to estimate the populations of folded and unfolded Ton box.
RESULTS
The Ton box exhibits a substrate-dependent
unfolding in bilayers but not in protein crystals
The label at position 10 was chosen for these experiments
for two reasons. First, the incorporation of R1 at some sites
may perturb the Ton box fold; however, the incorporation of
R1 at position 10 does not appear to be highly perturbing
(20). Second, the spectra from BtuB-V10R1 are particularly
good at revealing different conformational substates of the
Ton box, and these states are easily quantitated from the
EPR spectra of V10R1.
Shown in Fig. 2 a are EPR spectra for BtuB-V10R1 with
and without substrate in lipid bilayers composed of POPC.
Spectra for BtuB-V10R1 in bilayers have been reported
previously (20), and in the absence of substrate the spectrum
is dominated by a broad component resulting from an
immobile spin-labeled side chain that is near the rigid-limit
of nitroxide motion at X-band (tc 30–50 ns). This broad
component results from a label that is in strong tertiary
contact with other side chains in BtuB. In the presence of
substrate, the spectrum changes dramatically and is domi-
nated by a narrow high-amplitude component arising from a
motionally averaged nitroxide attached to a disordered
backbone segment. A careful examination of the EPR line-
shapes in Fig. 2 a indicates that in each case (with or without
substrate), both immobile and mobile components can be
distinguished. These components represent folded and
unfolded substates of the Ton box in equilibrium (40), and
the populations of these substates may be estimated from
the EPR spectra using spectral subtraction (see Methods).
This estimate shows that in the presence of substrate
TABLE 1
Data collection and refinement for BtuB-V10R1
Structure
BtuB-V10R1 apo
BtuB-V10R1 þCa2þB12
Data collection
Beamline
APS-22ID
APS-22ID
Wavelength (A˚ )
1.000
1.000
Temperature (K)
90
90
Reflections observed
311,539
294,094
Unique reflections
32,472
32,358
Resolution range (A˚ )* 50–2.40 (2.49–2.40)
50–2.45 (2.54–2.45)
Space group
P3121
P3121
Cell dimensions
a ¼ b ¼ 81.3 A˚ ,
c ¼ 226.6 A˚
a ¼ b ¼ 82.1 A˚ ,
c ¼ 224.5 A˚
a ¼ b ¼ 90,
g ¼ 120
a ¼ b ¼ 90,
g ¼ 120
Rsym (%)
9.1 (38.3)
12.1 (45.8)
Redundancy
9.6
9.1
Refinement
Resolution range (A˚ )
44.1–2.44 (2.50–2.44) 44.0–2.44 (2.51–2.44)
Reflections used
30,769
30,642
Completeness (%)
97.6 (79.3)
96.6 (67.3)
Rcryst (%)y
22.1
22.9
Rfree (%)z
24.8
27.5
Root-mean-square deviations
Bond lengths (A˚ )
0.021
0.019
Bond angles ()
1.839
2.037
Number of atoms
Protein
4605
4865
Water
113
76
Other
C8E4 (7), Mg (4)
CNCbl (1), Ca2þ (3),
C8E4 (6)
PDB accession code
3M8B
3M8D
*Highest resolution shell data shown in parentheses.
yRcryst ¼ SkFobsj-jFcalck / SjFobsj, where Fobs and Fcalc are the observed and
calculated structure factor amplitudes, respectively.
zRfree is Rcryst calculated using 5% of the data which is randomly chosen and
omitted from the refinement.
FIGURE 2
EPR spectra for V10R1 with (red traces) and without (blue
traces) substrate when BtuB is incorporated into (a) POPC bilayers, or
(b) in the protein crystal. The inset below is a 10 vertical expansion
showing a small signal from unfolded Ton box. The dashed vertical lines
indicate the positions of signals resulting from immobilized (i) and mobile
(m) nitroxide side chain, corresponding to folded and unfolded Ton box,
respectively.
Biophysical Journal 99(5) 1604–1610
1606
Freed et al.
~50% of the Ton box is unfolded, and the free energy differ-
ence (DG) between these two states is approximately zero.
Fig. 2 b shows an analogous pair of spectra obtained for
V10R1 in protein crystals in cryo buffer (see Methods)
with and without substrate. In the protein crystal, each
spectrum reflects a nitroxide near the rigid limit of motion
at X-band. The substrate-induced transition, which is clearly
seen in bilayers (Fig. 2 a), is absent. A careful examination
of the EPR spectrum for crystallized BtuB in the presence
of vitamin B12 (Fig. 2 b) reveals a very minor mobile
component (arrow in Fig. 2). This component matches the
lineshape obtained for V10R1 in the unfolded state and
appears to represent a small fraction of unfolded Ton box
in the presence of substrate. Quantitation of this minor
component by spectral subtraction indicates that it repre-
sents <0.5% of the total spin signal from V10R1, and that
the folded form of the Ton box is stabilized by at least
3 kcal/mol for BtuB bound to substrate in the protein crystal.
Because the energy difference between the folded and
unfolded states of the Ton box is close to zero in bilayers,
the free energy difference between these two protein sub-
states is altered (a DDG) by ~3 kcal/mol for BtuB-V10R1
in the protein crystal.
Structures from crystals of BtuB-V10R1 show
no evidence for a substrate-dependent unfolding
Protein crystals of BtuB-V10R1 in the absence and presence
of substrate diffracted to 2.4 A˚ and the refinement details are
given in Table 1. In both cases, the Ton box is resolved and
folded within the protein interior, and several extracellular
loops become resolved in presence of ligand, as seen previ-
ously for wild-type (wt) BtuB (22). Fig. 3, a and b, displays
periplasmic views of BtuB-V10R1, where the position of
V10R1 in the protein interior as well as the configuration
of the Ton box is shown. The label is sitting at the bottom
of a pocket facing the periplasmic surface; and as expected,
it is interacting with a number of side chains, including
R219 and R255. As a result, conversion between label ro-
tamers should be highly restricted, consistent with the rigid
limit spectra seen by EPR (Fig. 2 b).
The angles for c1 and c2 (Fig. 1 c) for R1 typically
assume a limited set of rotameric states on protein surface
sites, where the rotamers allow for an interaction between
Sd and HCa (41). Here V10R1 is found to have c1 and c2
angles of 56 and 69 in the apo form and 49 and 60 in
the CNCbl-bound form, which are both in a {p, p} configu-
ration using the conventions of Lovell et al. (42). The entire
set of spin-label dihedral angles for V10R1 is given in
Table 2. The Sd-HCa distance for the R1 side chain is
~4.5 A˚ , which is longer than that typically seen for R1 on
helix surface sites. Although this rotamer is energetically
allowed, it has not previously been observed in crystal
structures (41), presumably due to the sterically restricted
environment surrounding V10R1.
Fig. 3 c compares the Ton box for the V10R1 mutant
with and without CNCbl. The R1 side chain and the Ton
box to which it is attached remain folded into the protein
interior upon the addition of substrate, consistent with a
lack of change in the EPR spectra shown in Fig. 2 b for
FIGURE 3
(a) Periplasmic view of the structure and electron density
(1s) showing the placement of the spin-labeled side chain V10R1 and
residues that closely interact with the label in the apo form (PDB ID:
3M8B) of BtuB. (Magenta) Backbone of the Ton box. (Beige) N-terminal
fold. (b) Periplasmic view of BtuB-V10R1 similar to that shown in panel
a, except with van der Waals surfaces rendered for the atoms. The label,
V10R1, is at the base of a periplasmic pocket in close tertiary contact
with a number of atoms. (c) A comparison of the Ton box of BtuB-
V10R1 with and without substrate. A side view of the crystal structure of
the Ca2þ-B12 bound form of V10R1 (PDB ID: 3M8D) is shown with B12
bound, and the Ton box (magenta). This structure was aligned with the
apo form of BtuB-V10R1 where only the Ton box is rendered (blue).
Biophysical Journal 99(5) 1604–1610
Membrane Protein Conformational Exchange
1607
the protein crystal. Substrate addition to BtuB-V10R1
produces a change in the position of residue 7, as seen previ-
ously for wt protein (22). However, residue 6, which is
resolved in the wt structure, is not resolved for BtuB-
V10R1 once substrate is bound. A B-factor analysis of the
Ton box backbone and side-chain atoms indicates that
when compared to wt BtuB structures, the BtuB-V10R1
has higher B-factors for the Ton box N-terminal to position
10, and a larger difference between apo- and ligand-bound
forms. Nonetheless, the fold of the Ton box in BtuB-
V10R1 is virtually identical to that seen in the wt structure
(the root-mean-square deviation is 1.2 A˚ and 1.5 A˚ for the
apo and CNCbl-bound forms of the Ton box, respectively,
when V10R1 and wt are compared). Hence, minimal struc-
tural changes in the Ton box are induced by this particular
label.
Both the crystal lattice and solutes shift
the equilibrium between Ton box substates
To determine whether the crystal lattice makes a contribu-
tion to the free energy change when bilayer and crystal
forms of BtuB are compared, EPR spectra from V10R1
were compared for the protein crystal and the protein solu-
bilized into the cryo buffer. The two spectra for BtuB-
V10R1 (in the CNCbl bound form) are compared in
Fig. 4, a and b, and are clearly different. In particular,
the spectrum from solubilized protein (Fig. 4 b) yields a
mobile component with much higher amplitude than that
for the protein crystal (Fig. 4 a). This mobile signal has
a lineshape identical to that seen for the unfolded state in
the bilayer (Fig. 2 a). Quantitation of the two components
in this spectrum indicates that the mobile population makes
up ~8 5 2% of the total spins. This fraction of unfolded
Ton box corresponds to a change in free energy (a DDG
for this transition relative to the bilayer reconstituted
BtuB) of ~1.5 5 0.2 kcal/mol, indicating that solutes and
the crystal lattice make roughly equal contributions to the
change in conformational energy that is seen in the protein
crystal.
The lineshapes for the immobilized component in the
absence of substrate for the bilayer reconstituted and crys-
tallized BtuB-V10R1 are shown in Fig. 4, c and d, respec-
tively. In this case the mobile component was subtracted
from the bilayer BtuB-V10R1 (Fig. 2 a) to yield the immo-
bile component in Fig. 4 c. Both these lineshapes result from
immobile spin labels near the rigid limit of nitroxide
motion. However, the hyperfine extrema in Fig. 4 c are
not as distinct as in Fig. 4 d, and components representing
the g-tensor anisotropy in the central (mI ¼ 0) resonance
of BtuB-V10R1 are better resolved in the protein crystal
(Fig. 4 d). This difference provides an indication that addi-
tional motional modes are available for V10R1 in the
bilayer environment.
DISCUSSION
In this work, SDSL was used to examine a conformational
equilibrium in the Escherichia coli outer membrane trans-
porter, BtuB, both in lipid bilayers and in protein crystals.
The results indicate that the equilibrium between folded and
unfolded forms of the Ton box is shifted by ~3 kcal/mol
when the protein is taken from the bilayer phase to the protein
crystal phase. This has the effect of stabilizing the folded form
of the Ton box in the protein crystal, and it provides an expla-
nation for the observation that the Ton box is resolved both in
the absence and presenceofsubstrate in crystal structures(22),
but is seen to unfold in bilayers. It should be emphasized that
protein crystallography does not provide an incorrect structure
for BtuB. The conditions of the protein crystal alter the equi-
librium distribution of substates, compared to the distribution
found by EPR, to favor the more-compact-ordered conformer.
Osmolytes, such as PEGs, are well known to modify
protein behavior (43), and previous work has demonstrated
that osmolytes stabilize a folded form of the Ton box
(15,24,25)
and
stabilize
more-compact,
less-hydrated
conformations of the extracellular ligand-binding loops
(44). The data obtained here indicate that solutes and the
crystal lattice contribute almost equally to the energy
change seen in the Ton box equilibrium. While the action
of PEGs and other osmolytes is reasonably well understood,
FIGURE 4
EPR spectra from BtuB-V10R1 with bound ligand in (a) the
protein crystal, (b) in the crystallization buffer at a protein concentration too
dilute to form crystals, and in the apo state in (c) lipid bilayers and (d) the
protein crystal. The symbols i and m indicate immobilized and mobile
components in the spectra for panel b. The spectrum in panel c is identical
to the spectrum in Fig. 2 a, except that the small mobile component seen in
Fig. 2 a has been subtracted. All spectra are 100 Gauss scans.
TABLE 2
Summary of R1 side-chain dihedral angles and
rotamer designation
Mutant
Rotamer
c1
c2
c3
c4
c5
V10R1 apo
{p,p}
56
69
83
67
67
V10R1 Ca2þ and B12
{p,p}
49
60
70
93
46
Biophysical Journal 99(5) 1604–1610
1608
Freed et al.
it is not presently known how the protein lattice in the
crystal couples to the Ton box equilibrium and stabilizes
its folded form in BtuB. Previous work has shown that there
is an interaction between charged residues near the Ton box
and the BtuB b-barrel (40), and that eliminating this interac-
tion unfolds the Ton box. Conceivably, a change in the
dynamics or structure of the BtuB b-barrel when the protein
is in the crystal lattice might alter the energy of this interac-
tion and account for the effect of the lattice upon the Ton
box.
The structural biology of membrane proteins is far from
mature, and it is not known whether the effects seen here
on protein conformational sampling apply to a wider range
of membrane proteins. Bacteriorhodopsin is perhaps the
best-studied membrane protein, and there are indications
that the kinetics of the photocycle are modified in the
three-dimensional crystalline lattice when compared to the
native membrane (45). Solution NMR can provide high-
resolution structural data on membrane proteins in micelles,
allowing comparisons to be made with crystal structures.
NMR spectroscopy is often found to resolve portions of
proteins that are not resolved by crystallography (as seen
for DsbB (46)), presumably because NMR is better at exam-
ining structures that are inherently dynamic. In outer
membrane porins, such as OmpA, the strands of the b-barrel
are shorter in the NMR-derived structures than in the crystal
structures (47); however, it is not clear whether this differ-
ence is a result of crystallization conditions or the micellar
environment used for NMR. Even in a reconstituted bilayer
environment (which is a much better approximation to the
native environment than the protein crystal), protein confor-
mational sampling may be modulated relative to the native
environment (48–51); however, many of these effects appear
to be due to the fraction of acidic lipids selected for the
reconstitution, which in turn control local ion concentrations
and pH. As for BtuB, the Ton box equilibrium does not
appear to be modulated by lipid composition (Q. Xu and
D. S. Cafiso, unpublished); this equilibrium is maintained
within range of reconstituted bilayers as well as intact outer
membrane preparations (19).
Changes in the equilibrium distribution of protein confor-
mational substates are thought to underlie protein signaling
events (52) and allostery (53). Because of its fast timescale,
EPR spectroscopy is particularly well suited to detect these
conformational substates and to measure conformational
equilibria in proteins. In BtuB, SDSL demonstrates that
both the folded and unfolded states of the Ton box are
sampled and that substrate binding shifts the equilibrium
to the more disordered state. Furthermore, colicin E3, which
is also a ligand for BtuB, shifts the Ton box equilibrium to
favor the folded, ordered state of the Ton box (54). These are
precisely the types of changes that are proposed to underlie
protein signaling, and in the present case, they may function
to regulate coupling between BtuB and inner membrane
protein TonB.
We thank Dr. Robert Nakamoto for careful reading of this manuscript.
This work was supported by National Institutes of Health grants Nos.
NIGMS 035215 to D.S.C. and NIGMS 079800 to M.C.W. Use of the
Advanced Photon Source was supported by the U. S. Department of Energy,
Office of Science, Office of Basic Energy Sciences, under contract No. DE-
AC02-06CH11357. Data were collected at Southeast Regional Collabora-
tive Access Team (SER-CAT) 22-ID beamline at the Advanced Photon
Source, Argon National Laboratory, Argonne, IL. Supporting institutions
may be found at http://www.ser-cat.org/members.html.
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|
3M8D
|
Crystal structure of spin-labeled BtuB V10R1 with bound calcium and cyanocobalamin
|
Conformational Exchange in a Membrane Transport Protein Is Altered
in Protein Crystals
Daniel M. Freed,† Peter S. Horanyi,‡ Michael C. Wiener,‡ and David S. Cafiso†‡*
†Departments of Chemistry and ‡Molecular Physiology and Biological Physics, and the Biophysics Program, University of Virginia,
Charlottesville, Virginia
ABSTRACT
Successful macromolecular crystallography requires solution conditions that may alter the conformational
sampling of a macromolecule. Here, site-directed spin labeling is used to examine a conformational equilibrium within BtuB,
the Escherichia coli outer membrane transporter for vitamin B12. Electron paramagnetic resonance (EPR) spectra from
a spin label placed within the N-terminal energy coupling motif (Ton box) of BtuB indicate that this segment is in equilibrium
between folded and unfolded forms. In bilayers, substrate binding shifts this equilibrium toward the unfolded form; however,
EPR spectra from this same spin-labeled mutant indicate that this unfolding transition is blocked in protein crystals. Moreover,
crystal structures of this spin-labeled mutant are consistent with the EPR result. When the free energy difference between
substates is estimated from the EPR spectra, the crystal environment is found to alter this energy by 3 kcal/mol when compared
to the bilayer state. Approximately half of this energy change is due to solutes or osmolytes in the crystallization buffer, and the
remainder is contributed by the crystal lattice. These data provide a quantitative measure of how a conformational equilibrium in
BtuB is modified in the crystal environment, and suggest that more-compact, less-hydrated substates will be favored in protein
crystals.
INTRODUCTION
Proteins are dynamic and structurally heterogeneous. They
exhibit collective and uncoupled motions over a wide range
of timescales (1,2) and they may assume numerous discrete
structural substates that are in equilibrium. This motion and
sampling of structural states is important and appears to be
critical to enzyme activity and allosteric regulation (3–5). In
protein crystals, dynamics and structural heterogeneity are
present, and at sufficiently high resolution, may be well
represented by the use of multiple conformations of the
side-chain (6) and backbone atoms (7).
There are indications that protein crystallization and the
conditions used for crystallization may alter protein dy-
namics and conformational sampling. Molecular dynamics
simulations of proteins in a crystal lattice have been
performed for both soluble (8–10), and membrane (11)
proteins. These computational efforts suggest that confor-
mational sampling may be altered by protein crystallization.
In addition, experiments employing precipitants or osmo-
lytes similar to those used in protein crystallization demon-
strate that these solutes may have a significant effect on
exchange between long-lived conformational substates; for
example, osmolytes have been found to alter conformational
substates involved in enzymatic activity (12,13) and ion
conduction (14).
In BtuB, the outer membrane Escherichia coli vitamin
B12 (CNCbl) transporter, an electron paramagnetic reso-
nance
(EPR)-based
method
termed
site-directed
spin
labeling (SDSL), has been used to investigate the dynamics
and structural transitions in an N-terminal energy coupling
segment termed the Ton box (15). The Ton box couples
BtuB to the inner membrane protein TonB, which provides
energy for transport (16–18). SDSL provides strong
evidence that the Ton box undergoes a vitamin B12-depen-
dent unfolding (19,20), as depicted in Fig. 1. This event
moves the Ton box 20–30 A˚ into the periplasmic space,
where it may act as a trigger to initiate BtuB-TonB interac-
tions (21). In contrast, the Ton box remains folded within the
transporter in crystal structures of BtuB either in the pres-
ence or absence of substrate. While there are small shifts
in the conformation of the Ton box upon substrate binding,
no evidence is seen for the substrate-dependent unfolding
observed spectroscopically (22).
The discrepancy between the spectroscopic and crystallo-
graphic result might have several origins. EPR spectroscopy
of membrane-associated BtuB demonstrates that there is an
equilibrium between folded and unfolded substates of the
Ton box, and that this equilibrium is shifted toward the
more folded state by the osmolytes used in the BtuB crystal-
lization (23–25). Osmolytes, such as polyethylene glycols
(PEGs), are believed to be excluded from hydrated protein
surfaces (26,27), thereby raising the energy of the protein
and reducing its solubility. As a result, the presence of os-
molytes will favor conformational substates that are less
hydrated (12,28,29). The packing of the protein within the
crystal lattice might also account for the difference between
the spectroscopic and crystallographic result. Although
protein-protein contacts within the unit cell should not
interfere sterically with the unfolding of the Ton box, the
contributions that the lattice might make to the Ton box
equilibrium are not known.
Submitted April 26, 2010, and accepted for publication June 14, 2010.
*Correspondence: cafiso@virginia.edu
Editor: David D. Thomas.
2010 by the Biophysical Society
0006-3495/10/09/1604/7
$2.00
doi: 10.1016/j.bpj.2010.06.026
1604
Biophysical Journal
Volume 99
September 2010
1604–1610
To determine how the Ton box equilibrium is modified
within the protein crystal compared to the bilayer state,
we generated a spin-labeled mutant of BtuB where the nitro-
xide side chain R1 (Fig. 1 c) is incorporated into the Ton box
at position 10. EPR spectroscopy was then carried out in
parallel with x-ray diffraction and structure determination
of this labeled BtuB mutant (V10R1) using the same protein
crystals. The EPR spectra obtained from the protein crystal
indicate that the substrate-dependent Ton box transition is
blocked. This spectroscopic result is consistent with crystal
structures of this BtuB mutant, which indicate that the Ton
box remains folded and the R1 side chain buried with or
without substrate. By comparing EPR spectra of BtuB-
V10R1 in bilayers with spectra from the protein crystal,
we estimate the free energy change induced by the crystal
environment on this conformational equilibrium, and dissect
the energetic contributions made by solute and the crystal
lattice.
MATERIALS AND METHODS
Mutagenesis, expression, and purification
The V10C mutation was introduced into btuB using a QuikChange site-
directed mutagenesis kit (Stratagene, La Jolla, CA), and was subsequently
verified by nucleotide sequencing. Expression and purification of BtuB for
the formation of protein crystals was performed as described previously
(22), and BtuB was reconstituted into vesicle bilayers by following a
procedure described elsewhere (30).
Spin labeling
For spin labeling, the first round of purification was paused before initiation
of the salt gradient. The Q-Sepharose slurry (Amersham, Piscataway, NJ)
bound with BtuB was transferred to a conical tube and reacted with
1 mL of 45 mM S-(1-oxy-2,2,5,6-tetramethylpyrroline-3-methyl)methane-
thiosulfonate (MTSL; Toronto Research Chemicals, Ontario, Canada) for
4 h at room temperature.
Crystallization and crystallographic data
collection
Purified BtuB (11 mg/mL in 30 mM Tris pH 8.0, 20 mM C8E4) was crys-
tallized by mixing 1 mL of BtuB and 1 mL of reservoir buffer in an EasyXtal
hanging-drop tray (Qiagen, Germantown, MD), containing 200 mL of total
reservoir buffer for each crystallization condition, and followed by incuba-
tion at 290 K. The reservoir buffer consisted of 200–550 mM magnesium
acetate, 5.0–7.5% PEG3350, and 20 mM Bis Tris at pH 6.6. Crystals
were visible after 1–2 days, and grew to ~200 mm in the longest dimension
after 1–2 weeks. For crystals to be incubated with substrate, 1 mL of soaking
buffer (150 mM calcium chloride, 2.5% PEG3350, 20 mM Bis Tris at
pH 6.6, and 10 mM C8E4) was added to each well, followed by incubation
overnight. The crystals were subsequently transferred into soaking buffer
containing 1 mM cyanocobalamin and 20% glycerol, and allowed to incu-
bate for at least 4 h. For x-ray diffraction, apo and Ca2þB12-soaked crystals
were transferred to cryo-buffer (150 mM magnesium acetate or calcium
chloride, 2.5% PEG3350, 20 mM Bis Tris at pH 6.6, 10 mM C8E4, and
20% glycerol) for 1–2 min before loop mounting and cryocooling by inser-
tion into liquid nitrogen. Diffraction data were taken at 90 K at the 22ID
beamline at the Advanced Photon Source (Argonne National Laboratory,
Argonne, IL). See Table 1 for more details.
Structure determination
Indexing, integration, and scaling of the diffraction data was performed
using HKL2000 (31). The structures were solved with PHASER (32)
maximum likelihood molecular replacement method, using PDB deposi-
tions 1NQE and 1NQH (22) as search models for the apo- and Ca2þB12-
bound data, respectively. To reduce model bias, V10 was deleted from
the apo-search model, and the entire Ton box was deleted from the Ca2þ-
CNCbl-bound search model. Model building was done in COOT (33),
and unrestrained TLS (34) refinement was performed using REFMAC
(35) and PHENIX was used to refine the occupancy of the spin label
(36). The spin-labeled residue V10R1 was manually built in COOT. Anom-
alous difference Fourier maps were calculated to accurately position bound
cobalt and calcium using Sfall and fast-Fourier transform (37). Completed
structures were evaluated and validated with MolProbity (38).
Electron paramagnetic resonance
Apo- and Ca2þ B12-soaked crystals were incubated for at least 4 h in
cryobuffer and soaking buffer (with 1 mM cyanocobalamin and 20% glyc-
erol), respectively. Crystals were then transferred to a 0.60 ID 0.84 OD
round capillary with a syringe (Hamilton Syringe, Bonaduz, Switzerland)
for EPR spectroscopy, which was performed on an X-band EMX spectrom-
eter (Bruker Biospin, Billerica, MA) equipped with a dielectric resonator.
All EPR spectra were recorded with a 100 G magnetic field sweep at
2.0 mW incident power at a temperature of 298 K. The phasing, normaliza-
tion, and subtraction of EPR spectra was performed using LabVIEW soft-
ware provided by Dr. Christian Altenbach (University of California,
Los Angeles, California).
FIGURE 1
BtuB in the (a) apo form where the Ton box position is
highlighted (PDB ID: 1NQE). (b) Vitamin B12 bound form of BtuB
showing the state of the Ton box as determined by EPR spectra and pulse
EPR distance measurements (based upon PDB ID 1NQH and spectroscopic
restraints obtained for the Ton box in bilayers (21)). This unfolding event
places the Ton box as much as 30 A˚ into the periplasmic space. (c) The
structure of the spin-labeled R1 side chain and dihedral angles that define
the rotamers of R1.
Biophysical Journal 99(5) 1604–1610
Membrane Protein Conformational Exchange
1605
To determine the free energies and free energy changes between Ton box
states, the population of each Ton box conformation was determined by
spectral subtraction and quantitation of the spectral components as
described previously (24). For BtuB V10R1, the EPR spectra are linear
combinations of the spectra resulting from the folded and unfolded Ton
box conformations. As a result, the fraction of spins in each population
may be estimated by determining the contribution that each conformation
makes to the total spectrum. The label at position 10 was chosen for these
measurements, because EPR spectra for V10R1 yield dramatically different
lineshapes for the folded and unfolded forms of the Ton box. As a result, it
is easy to simulate both the folded and unfolded lineshape. In this case, the
mobile lineshape was simulated (using Redfield theory (39)) and subtracted
from the composite spectrum until a spectrum corresponding to the purely
folded Ton box conformation was obtained. Double integration of the first
derivative EPR spectra yields numbers that are proportional to spin number
and was used to estimate the populations of folded and unfolded Ton box.
RESULTS
The Ton box exhibits a substrate-dependent
unfolding in bilayers but not in protein crystals
The label at position 10 was chosen for these experiments
for two reasons. First, the incorporation of R1 at some sites
may perturb the Ton box fold; however, the incorporation of
R1 at position 10 does not appear to be highly perturbing
(20). Second, the spectra from BtuB-V10R1 are particularly
good at revealing different conformational substates of the
Ton box, and these states are easily quantitated from the
EPR spectra of V10R1.
Shown in Fig. 2 a are EPR spectra for BtuB-V10R1 with
and without substrate in lipid bilayers composed of POPC.
Spectra for BtuB-V10R1 in bilayers have been reported
previously (20), and in the absence of substrate the spectrum
is dominated by a broad component resulting from an
immobile spin-labeled side chain that is near the rigid-limit
of nitroxide motion at X-band (tc 30–50 ns). This broad
component results from a label that is in strong tertiary
contact with other side chains in BtuB. In the presence of
substrate, the spectrum changes dramatically and is domi-
nated by a narrow high-amplitude component arising from a
motionally averaged nitroxide attached to a disordered
backbone segment. A careful examination of the EPR line-
shapes in Fig. 2 a indicates that in each case (with or without
substrate), both immobile and mobile components can be
distinguished. These components represent folded and
unfolded substates of the Ton box in equilibrium (40), and
the populations of these substates may be estimated from
the EPR spectra using spectral subtraction (see Methods).
This estimate shows that in the presence of substrate
TABLE 1
Data collection and refinement for BtuB-V10R1
Structure
BtuB-V10R1 apo
BtuB-V10R1 þCa2þB12
Data collection
Beamline
APS-22ID
APS-22ID
Wavelength (A˚ )
1.000
1.000
Temperature (K)
90
90
Reflections observed
311,539
294,094
Unique reflections
32,472
32,358
Resolution range (A˚ )* 50–2.40 (2.49–2.40)
50–2.45 (2.54–2.45)
Space group
P3121
P3121
Cell dimensions
a ¼ b ¼ 81.3 A˚ ,
c ¼ 226.6 A˚
a ¼ b ¼ 82.1 A˚ ,
c ¼ 224.5 A˚
a ¼ b ¼ 90,
g ¼ 120
a ¼ b ¼ 90,
g ¼ 120
Rsym (%)
9.1 (38.3)
12.1 (45.8)
Redundancy
9.6
9.1
Refinement
Resolution range (A˚ )
44.1–2.44 (2.50–2.44) 44.0–2.44 (2.51–2.44)
Reflections used
30,769
30,642
Completeness (%)
97.6 (79.3)
96.6 (67.3)
Rcryst (%)y
22.1
22.9
Rfree (%)z
24.8
27.5
Root-mean-square deviations
Bond lengths (A˚ )
0.021
0.019
Bond angles ()
1.839
2.037
Number of atoms
Protein
4605
4865
Water
113
76
Other
C8E4 (7), Mg (4)
CNCbl (1), Ca2þ (3),
C8E4 (6)
PDB accession code
3M8B
3M8D
*Highest resolution shell data shown in parentheses.
yRcryst ¼ SkFobsj-jFcalck / SjFobsj, where Fobs and Fcalc are the observed and
calculated structure factor amplitudes, respectively.
zRfree is Rcryst calculated using 5% of the data which is randomly chosen and
omitted from the refinement.
FIGURE 2
EPR spectra for V10R1 with (red traces) and without (blue
traces) substrate when BtuB is incorporated into (a) POPC bilayers, or
(b) in the protein crystal. The inset below is a 10 vertical expansion
showing a small signal from unfolded Ton box. The dashed vertical lines
indicate the positions of signals resulting from immobilized (i) and mobile
(m) nitroxide side chain, corresponding to folded and unfolded Ton box,
respectively.
Biophysical Journal 99(5) 1604–1610
1606
Freed et al.
~50% of the Ton box is unfolded, and the free energy differ-
ence (DG) between these two states is approximately zero.
Fig. 2 b shows an analogous pair of spectra obtained for
V10R1 in protein crystals in cryo buffer (see Methods)
with and without substrate. In the protein crystal, each
spectrum reflects a nitroxide near the rigid limit of motion
at X-band. The substrate-induced transition, which is clearly
seen in bilayers (Fig. 2 a), is absent. A careful examination
of the EPR spectrum for crystallized BtuB in the presence
of vitamin B12 (Fig. 2 b) reveals a very minor mobile
component (arrow in Fig. 2). This component matches the
lineshape obtained for V10R1 in the unfolded state and
appears to represent a small fraction of unfolded Ton box
in the presence of substrate. Quantitation of this minor
component by spectral subtraction indicates that it repre-
sents <0.5% of the total spin signal from V10R1, and that
the folded form of the Ton box is stabilized by at least
3 kcal/mol for BtuB bound to substrate in the protein crystal.
Because the energy difference between the folded and
unfolded states of the Ton box is close to zero in bilayers,
the free energy difference between these two protein sub-
states is altered (a DDG) by ~3 kcal/mol for BtuB-V10R1
in the protein crystal.
Structures from crystals of BtuB-V10R1 show
no evidence for a substrate-dependent unfolding
Protein crystals of BtuB-V10R1 in the absence and presence
of substrate diffracted to 2.4 A˚ and the refinement details are
given in Table 1. In both cases, the Ton box is resolved and
folded within the protein interior, and several extracellular
loops become resolved in presence of ligand, as seen previ-
ously for wild-type (wt) BtuB (22). Fig. 3, a and b, displays
periplasmic views of BtuB-V10R1, where the position of
V10R1 in the protein interior as well as the configuration
of the Ton box is shown. The label is sitting at the bottom
of a pocket facing the periplasmic surface; and as expected,
it is interacting with a number of side chains, including
R219 and R255. As a result, conversion between label ro-
tamers should be highly restricted, consistent with the rigid
limit spectra seen by EPR (Fig. 2 b).
The angles for c1 and c2 (Fig. 1 c) for R1 typically
assume a limited set of rotameric states on protein surface
sites, where the rotamers allow for an interaction between
Sd and HCa (41). Here V10R1 is found to have c1 and c2
angles of 56 and 69 in the apo form and 49 and 60 in
the CNCbl-bound form, which are both in a {p, p} configu-
ration using the conventions of Lovell et al. (42). The entire
set of spin-label dihedral angles for V10R1 is given in
Table 2. The Sd-HCa distance for the R1 side chain is
~4.5 A˚ , which is longer than that typically seen for R1 on
helix surface sites. Although this rotamer is energetically
allowed, it has not previously been observed in crystal
structures (41), presumably due to the sterically restricted
environment surrounding V10R1.
Fig. 3 c compares the Ton box for the V10R1 mutant
with and without CNCbl. The R1 side chain and the Ton
box to which it is attached remain folded into the protein
interior upon the addition of substrate, consistent with a
lack of change in the EPR spectra shown in Fig. 2 b for
FIGURE 3
(a) Periplasmic view of the structure and electron density
(1s) showing the placement of the spin-labeled side chain V10R1 and
residues that closely interact with the label in the apo form (PDB ID:
3M8B) of BtuB. (Magenta) Backbone of the Ton box. (Beige) N-terminal
fold. (b) Periplasmic view of BtuB-V10R1 similar to that shown in panel
a, except with van der Waals surfaces rendered for the atoms. The label,
V10R1, is at the base of a periplasmic pocket in close tertiary contact
with a number of atoms. (c) A comparison of the Ton box of BtuB-
V10R1 with and without substrate. A side view of the crystal structure of
the Ca2þ-B12 bound form of V10R1 (PDB ID: 3M8D) is shown with B12
bound, and the Ton box (magenta). This structure was aligned with the
apo form of BtuB-V10R1 where only the Ton box is rendered (blue).
Biophysical Journal 99(5) 1604–1610
Membrane Protein Conformational Exchange
1607
the protein crystal. Substrate addition to BtuB-V10R1
produces a change in the position of residue 7, as seen previ-
ously for wt protein (22). However, residue 6, which is
resolved in the wt structure, is not resolved for BtuB-
V10R1 once substrate is bound. A B-factor analysis of the
Ton box backbone and side-chain atoms indicates that
when compared to wt BtuB structures, the BtuB-V10R1
has higher B-factors for the Ton box N-terminal to position
10, and a larger difference between apo- and ligand-bound
forms. Nonetheless, the fold of the Ton box in BtuB-
V10R1 is virtually identical to that seen in the wt structure
(the root-mean-square deviation is 1.2 A˚ and 1.5 A˚ for the
apo and CNCbl-bound forms of the Ton box, respectively,
when V10R1 and wt are compared). Hence, minimal struc-
tural changes in the Ton box are induced by this particular
label.
Both the crystal lattice and solutes shift
the equilibrium between Ton box substates
To determine whether the crystal lattice makes a contribu-
tion to the free energy change when bilayer and crystal
forms of BtuB are compared, EPR spectra from V10R1
were compared for the protein crystal and the protein solu-
bilized into the cryo buffer. The two spectra for BtuB-
V10R1 (in the CNCbl bound form) are compared in
Fig. 4, a and b, and are clearly different. In particular,
the spectrum from solubilized protein (Fig. 4 b) yields a
mobile component with much higher amplitude than that
for the protein crystal (Fig. 4 a). This mobile signal has
a lineshape identical to that seen for the unfolded state in
the bilayer (Fig. 2 a). Quantitation of the two components
in this spectrum indicates that the mobile population makes
up ~8 5 2% of the total spins. This fraction of unfolded
Ton box corresponds to a change in free energy (a DDG
for this transition relative to the bilayer reconstituted
BtuB) of ~1.5 5 0.2 kcal/mol, indicating that solutes and
the crystal lattice make roughly equal contributions to the
change in conformational energy that is seen in the protein
crystal.
The lineshapes for the immobilized component in the
absence of substrate for the bilayer reconstituted and crys-
tallized BtuB-V10R1 are shown in Fig. 4, c and d, respec-
tively. In this case the mobile component was subtracted
from the bilayer BtuB-V10R1 (Fig. 2 a) to yield the immo-
bile component in Fig. 4 c. Both these lineshapes result from
immobile spin labels near the rigid limit of nitroxide
motion. However, the hyperfine extrema in Fig. 4 c are
not as distinct as in Fig. 4 d, and components representing
the g-tensor anisotropy in the central (mI ¼ 0) resonance
of BtuB-V10R1 are better resolved in the protein crystal
(Fig. 4 d). This difference provides an indication that addi-
tional motional modes are available for V10R1 in the
bilayer environment.
DISCUSSION
In this work, SDSL was used to examine a conformational
equilibrium in the Escherichia coli outer membrane trans-
porter, BtuB, both in lipid bilayers and in protein crystals.
The results indicate that the equilibrium between folded and
unfolded forms of the Ton box is shifted by ~3 kcal/mol
when the protein is taken from the bilayer phase to the protein
crystal phase. This has the effect of stabilizing the folded form
of the Ton box in the protein crystal, and it provides an expla-
nation for the observation that the Ton box is resolved both in
the absence and presenceofsubstrate in crystal structures(22),
but is seen to unfold in bilayers. It should be emphasized that
protein crystallography does not provide an incorrect structure
for BtuB. The conditions of the protein crystal alter the equi-
librium distribution of substates, compared to the distribution
found by EPR, to favor the more-compact-ordered conformer.
Osmolytes, such as PEGs, are well known to modify
protein behavior (43), and previous work has demonstrated
that osmolytes stabilize a folded form of the Ton box
(15,24,25)
and
stabilize
more-compact,
less-hydrated
conformations of the extracellular ligand-binding loops
(44). The data obtained here indicate that solutes and the
crystal lattice contribute almost equally to the energy
change seen in the Ton box equilibrium. While the action
of PEGs and other osmolytes is reasonably well understood,
FIGURE 4
EPR spectra from BtuB-V10R1 with bound ligand in (a) the
protein crystal, (b) in the crystallization buffer at a protein concentration too
dilute to form crystals, and in the apo state in (c) lipid bilayers and (d) the
protein crystal. The symbols i and m indicate immobilized and mobile
components in the spectra for panel b. The spectrum in panel c is identical
to the spectrum in Fig. 2 a, except that the small mobile component seen in
Fig. 2 a has been subtracted. All spectra are 100 Gauss scans.
TABLE 2
Summary of R1 side-chain dihedral angles and
rotamer designation
Mutant
Rotamer
c1
c2
c3
c4
c5
V10R1 apo
{p,p}
56
69
83
67
67
V10R1 Ca2þ and B12
{p,p}
49
60
70
93
46
Biophysical Journal 99(5) 1604–1610
1608
Freed et al.
it is not presently known how the protein lattice in the
crystal couples to the Ton box equilibrium and stabilizes
its folded form in BtuB. Previous work has shown that there
is an interaction between charged residues near the Ton box
and the BtuB b-barrel (40), and that eliminating this interac-
tion unfolds the Ton box. Conceivably, a change in the
dynamics or structure of the BtuB b-barrel when the protein
is in the crystal lattice might alter the energy of this interac-
tion and account for the effect of the lattice upon the Ton
box.
The structural biology of membrane proteins is far from
mature, and it is not known whether the effects seen here
on protein conformational sampling apply to a wider range
of membrane proteins. Bacteriorhodopsin is perhaps the
best-studied membrane protein, and there are indications
that the kinetics of the photocycle are modified in the
three-dimensional crystalline lattice when compared to the
native membrane (45). Solution NMR can provide high-
resolution structural data on membrane proteins in micelles,
allowing comparisons to be made with crystal structures.
NMR spectroscopy is often found to resolve portions of
proteins that are not resolved by crystallography (as seen
for DsbB (46)), presumably because NMR is better at exam-
ining structures that are inherently dynamic. In outer
membrane porins, such as OmpA, the strands of the b-barrel
are shorter in the NMR-derived structures than in the crystal
structures (47); however, it is not clear whether this differ-
ence is a result of crystallization conditions or the micellar
environment used for NMR. Even in a reconstituted bilayer
environment (which is a much better approximation to the
native environment than the protein crystal), protein confor-
mational sampling may be modulated relative to the native
environment (48–51); however, many of these effects appear
to be due to the fraction of acidic lipids selected for the
reconstitution, which in turn control local ion concentrations
and pH. As for BtuB, the Ton box equilibrium does not
appear to be modulated by lipid composition (Q. Xu and
D. S. Cafiso, unpublished); this equilibrium is maintained
within range of reconstituted bilayers as well as intact outer
membrane preparations (19).
Changes in the equilibrium distribution of protein confor-
mational substates are thought to underlie protein signaling
events (52) and allostery (53). Because of its fast timescale,
EPR spectroscopy is particularly well suited to detect these
conformational substates and to measure conformational
equilibria in proteins. In BtuB, SDSL demonstrates that
both the folded and unfolded states of the Ton box are
sampled and that substrate binding shifts the equilibrium
to the more disordered state. Furthermore, colicin E3, which
is also a ligand for BtuB, shifts the Ton box equilibrium to
favor the folded, ordered state of the Ton box (54). These are
precisely the types of changes that are proposed to underlie
protein signaling, and in the present case, they may function
to regulate coupling between BtuB and inner membrane
protein TonB.
We thank Dr. Robert Nakamoto for careful reading of this manuscript.
This work was supported by National Institutes of Health grants Nos.
NIGMS 035215 to D.S.C. and NIGMS 079800 to M.C.W. Use of the
Advanced Photon Source was supported by the U. S. Department of Energy,
Office of Science, Office of Basic Energy Sciences, under contract No. DE-
AC02-06CH11357. Data were collected at Southeast Regional Collabora-
tive Access Team (SER-CAT) 22-ID beamline at the Advanced Photon
Source, Argon National Laboratory, Argonne, IL. Supporting institutions
may be found at http://www.ser-cat.org/members.html.
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|
3M8E
|
Protein structure of Type III plasmid segregation TubR
|
Plasmid protein TubR uses a distinct mode of HTH-
DNA binding and recruits the prokaryotic tubulin
homolog TubZ to effect DNA partition
Lisheng Ni, Weijun Xu, Muthiah Kumaraswami, and Maria A. Schumacher1
Department of Biochemistry and Molecular Biology, University of Texas, M. D. Anderson Cancer Center, Unit 1000, Houston, TX 77030
Edited by Robert T. Sauer, Massachusetts Institute of Technology, Cambridge, MA, and approved May 11, 2010 (received for review March 22, 2010)
The segregation of plasmid DNA typically requires three elements:
a DNA centromere site, an NTPase, and a centromere-binding
protein. Because of their simplicity, plasmid partition systems
represent tractable models to study the molecular basis of DNA
segregation. Unlike eukaryotes, which utilize the GTPase tubulin
to segregate DNA, the most common plasmid-encoded NTPases
contain Walker-box and actin-like folds. Recently, a plasmid stabi-
lity cassette on Bacillus thuringiensis pBtoxis encoding a putative
FtsZ/tubulin-like NTPase called TubZ and DNA-binding protein
called TubR has been described. How these proteins collaborate
to impart plasmid stability, however, is unknown. Here we show
that the TubR structure consists of an intertwined dimer with a
winged helix-turn-helix (HTH) motif. Strikingly, however, the TubR
recognition helices mediate dimerization, making canonical HTH–
DNA interactions impossible. Mutagenesis data indicate that a
basic patch, encompassing the two wing regions and the N termini
of the recognition helices, mediates DNA binding, which indicates
an unusual HTH–DNA interaction mode in which the N termini of
the recognition helices insert into a single DNA groove and the
wings into adjacent DNA grooves. The TubZ structure shows that
it is as similar structurally to eukaryotic tubulin as it is to bacterial
FtsZ. TubZ forms polymers with guanine nucleotide-binding
characteristics and polymer dynamics similar to tubulin. Finally,
we show that the exposed TubZ C-terminal region interacts with
TubR-DNA, linking the TubR-bound pBtoxis to TubZ polymerization.
The combined data suggest a mechanism for TubZ-polymer pow-
ered plasmid movement.
T
he cytoskeletons of eukaryotic cells are constructed of three
primary elements: actin, tubulin, and intermediate filaments.
Although it had long been presumed that the proteins forming
these elements were absent in prokaryotes, it is now known that
prokaryotes contain structural homologs to all three components.
These prokaryotic proteins appear to carry out distinct functions
compared to their eukaryotic counterparts; however, their roles
are similar enough to indicate a likely common ancestor. The best
known prokaryotic homolog of tubulin is FtsZ. Both FtsZ and
tubulin form long filamentous structures by head to tail associa-
tion in a manner regulated by GTP, which binds between adjacent
subunits (1–4). However, unlike tubulin, FtsZ does not function
in DNA segregation but rather cell division. Specifically, it forms
a cytokinetic ring called the Z ring at midcell, which mediates
septation (5, 6). Recently, however, prokaryotic proteins encoded
on large plasmids harbored in bacilli showing 15–20% sequence
similarity to both FtsZ and tubulin have been identified and
dubbed TubZ (7–12). Studies showed that the Bacillus thuringien-
sis TubZ protein from the pBtoxis plasmid is essential for plasmid
DNA segregation.
DNA segregation of most low copy number plasmids is carried
out by specific partition (par) systems. These systems require only
three elements: a centromere DNA site, a centromere-binding
protein, and a partition NTPase (13, 14). Partition systems have
been classified into two main types on the basis of the kind
of NTPase present (15). Type I systems contain NTPases with
deviant Walker A-type ATPase folds, whereas type II systems uti-
lize actin-like NTPases. Interestingly, both types of NTPases form
polymers in NTP-dependent manners that are implicated to play
a role in plasmid DNA separation (16–19). The recent discovery
of TubZ NTPases has led to the designation of “type III” par sys-
tems (13, 14). The best studied of these systems is that found on
the pBtoxis plasmid in B. thuringiensis. This plasmid stability sys-
tem is represented by an operon encoding two proteins: ORF156
(TubZ) and ORF157 (TubR) (7–9, 11). TubR is a 11.6 kDa DNA-
binding protein that shows no sequence homology to any known
protein. Studies showed that TubR binds a 48-bp centromere con-
taining four repeat sites in the pBtoxis plasmid and also autore-
gulates tubRZ transcription (8, 9). TubZ is a 54.4 kDa protein that
can assemble into filaments in a GTP-dependent manner (12).
Both proteins were found to be required for plasmid stability
(9). However, how the TubR and TubZ proteins work together
to effect pBtoxis plasmid segregation is not known. To gain insight
into the molecular mechanism utilized by these proteins in DNA
segregation, we carried out structural and biochemical studies on
the pBtoxis TubR and TubZ proteins. The TubR structure reveals
that it employs a helix-turn-helix (HTH) motif in a previously un-
described manner to bind DNA. TubZ contains a tubulin/FtsZ
fold but has structural distinctions from these proteins indicating
that it forms distinct protofilaments. TubR binds the flexible
C-terminal region of TubZ, thus attaching the TubZ filament
to the pBtoxis plasmid, providing a mechanism for plasmid move-
ment and, ultimately, segregation.
Results and Discussion
Overall Structure of pBtoxis TubR. The crystal structure of the 107-
residue pBtoxis TubR protein was solved to 2.0-Å resolution by
selenomethionine multiple wavelength anomalous diffraction
(MAD) methods (Table S1). The structure contains two TubR
molecules in the crystallographic asymmetric unit and consists
of residues 6–102 of one subunit and 4–100 of the second subunit,
and has Rwork∕Rfree ¼ 23.8%∕27.0%. The TubR structure forms a
highly
intertwined
dimer
with
dimensions
30 × 30 × 60 Å3
(Fig. 1A). Each TubR subunit has the topology β1-α1-α2-α3-α4-β2-
β3-α5, which is similar to winged HTH motifs found in a number
of DNA-binding proteins in both prokaryotes and eukaryotes
(20). In TubR, α3-α4 forms the HTH motif and the loop between
Author contributions: M.A.S. designed research; L.N., W.X., M.K., and M.A.S. performed
research; M.A.S. analyzed data; and M.A.S. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Freely available online through the PNAS open access option.
Data deposition: The atomic coordinates and structure factor amplitudes for the WT TubR
(C2), WT TubR (I222), TubR(S63W), TubZ, and TubZ-GTP-γ-S structures have been
deposited with the Protein Data Bank, www.pdb.org (PDB ID codes 3M8E, 3M9A,
3M8F, 3M8K, and 3M89).
1To whom correspondence may be addressed. E-mail: maschuma@mdanderson.org.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1003817107/-/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.1003817107
PNAS ∣June 29, 2010 ∣vol. 107 ∣no. 26 ∣11763–11768
BIOCHEMISTRY
β2 and β3, the wing. Indeed, each TubR subunit shows the stron-
gest structural similarity to members of the ArsR family of
prokaryotic winged helix transcription regulators, in particular
the Staphylococcus aureus CzrA protein (21, 22). Superposition
of one subunit of TubR onto that of CzrA results in a rmsd of
2.7 Å. This similarity includes the core regions of the winged
HTH, because the loops and N-terminal regions of the proteins
are structurally distinct. For example, TubR contains a β-strand in
its N-terminal region compared to a long helix in the CzrA struc-
ture (Fig. 1B). This structural similarity initially suggested that
TubR may be a member of the ArsR family of proteins. However,
the arrangement of the TubR dimer was found to be strikingly
different from the dimer organization exhibited by the ArsR
proteins (Fig. 1C).
ArsR family members are involved in metal-regulated tran-
scription processes whereby they act as repressors in their apo
forms and are induced off their DNA sites upon metal binding
(22). The specific dimer structures of the ArsR proteins are cri-
tical for creation of their metal-binding motifs. Not only does
TubR form a very different dimer from the ArsR proteins, it also
does not harbor any of their metal-binding signatures nor does it
contain any other characterized metal-binding motif. Consistent
with this, we find that the addition of metals has no effect on TubR
DNA binding. Dimerization of the ArsR proteins is imparted by
residues from the N-terminal regions, α1 and α5, which, impor-
tantly, leaves its recognition helices exposed for DNA interaction
(22). By contrast, the TubR dimer is formed primarily by contacts
between its twofold related “recognition helices” α4 and α4·. This
interaction results in a near complete burial of these helices, leav-
ing only the N-terminal residues exposed to solvent. Whereas the
α4 and α4· interaction creates the dimer core, the dimer is further
stabilized by interactions between the twofold related β1 strands,
which swap to form an antiparallel β-sheet. Residues from α1 and
α5 interact with β1 to further seal the top of the dimer. The dimer
interface formed by these interactions is predominantly hydropho-
bic and buries a large 1;200 Å2 of subunit surface from solvent.
TubR Forms a “Recognition Helix Dimer”: Implications for DNA Bind-
ing. Gel filtration studies on TubR confirmed that it is a dimer in
solution. However, the finding from the structure that the TubR
α4 recognition helices are buried in the dimer core has important
implications in terms of its DNA-binding mechanism. Indeed, it
suggests that, although TubR contains a structurally canonical
HTH, it is not utilized for DNA binding in a manner typical
of HTH proteins. A second crystal form (I222) of TubR, which
was solved to 2.5-Å resolution, revealed the same TubR dimer.
The presence of the identical dimer in two different crystal forms
and its large buried surface area supports that the dimer observed
in the crystal structures is physiologically relevant. However, to
test this, we mutated residues within the recognition helices that
the structure indicates are critical for dimerization and assayed
the ability of the mutant proteins to dimerize via gel filtration.
Specifically, we mutated Ser-63 and Ala-67 individually to tryp-
tophan and arginine.
The structure shows that residues occupying positions 63 and
67 must be small and largely hydrophobic to permit the proper
packing of the α4 helices in the dimer (Fig. S1A). Hence, the in-
troduction of the bulky side chain of tryptophan and, in particu-
lar, the large as well as charged side chain of arginine would be
predicted to be highly disruptive to dimerization. Gel filtration
analyses on purified mutant proteins clearly showed that the ar-
ginine mutants exist primarily as monomers in solution (>80%),
whereas the tryptophan mutants were able to maintain the di-
meric state (Fig. 2A and Fig. S1B). However, all mutant proteins
showed reduced or loss of DNA-binding activity as ascertained by
fluorescence polarization (FP) studies, which examined TubR
protein binding to its centromere site (Fig. 2B and Fig. S1 C
and D) (9). The fact that the monomeric mutants were severely
impaired in DNA binding was not surprising. However, the
finding that the tryptophan mutants, which were largely dimeric,
displayed reduced DNA-binding activity suggested that their oli-
gomer structures might be altered. To address this issue, the struc-
ture of S63W TubR was solved to 2.8-Å resolution, resulting in
Rwork∕Rfree values of 20.4%∕26.6% (Table S1). The subunit struc-
ture of S63W TubR is essentially identical to that of WT TubR as
revealed by their superimposition (rmsd of 1.2 Å) (Fig. 2 C and
D). However, this single subunit overlay shows that the S63W
TubR dimer, although the same as the WT in general arrange-
ment, is forced into a more open oligomer conformation in which
one subunit is rotated 20° away from its dimer mate compared to
WT. This rotation is required to accommodate the bulky S63W
side chains (Fig. 2D). The N-terminal β1–β1′-sheet interaction
appears to play a key role in holding the TubR subunits together.
In addition, the tight stacking of the twofold related Trp63 indole
groups (3.5 Å) provides a compensatory interaction that, com-
bined with the β1–β1′ interaction, apparently permits retention
of the dimer state, indicating why the TubR tryptophan mutants
were able to maintain the dimer state, albeit an altered dimer
state relative to WT (Fig. 2 C and D). By contrast, the TubR
arginine mutations, which introduced both bulk and charge with-
in the predominantly hydrophobic dimer interface, were highly
destabilizing for dimerization. In addition, the finding that the
S63W TubR mutant forms an altered dimer explains the severe
effect on DNA binding because a correct dimer orientation is
likely essential for binding to its palindromic DNA sites (9).
TubR-DNA Model. Because all but the N-terminal residues of the
TubR recognition helices are buried in the dimer interface, TubR
must use a different mode of DNA binding than the ArsR or other
HTH containing proteins (23). Examination of surface electro-
statics of TubR reveals that one face of the protein is electro-
negative, whereas the other is strongly electropositive (Fig. 3A).
Notably, the positive region is composed of one large and contig-
uous basic patch. Basic residues in this region correspond to Arg-
74, Arg-77, and Lys-79, in the wing and Lys-43, which is on α3, the
helix preceding α4 in the HTH motif. These residues were mutated
singly to alanine to examine their roles in DNA binding (Fig. 3B).
FP experiments showed that mutation of the basic wing residues
resulted in either a complete (R74A and R77A) or nearly com-
plete (K79A) abrogation of DNA binding, indicating that the
wings play a major role in TubR DNA binding. Residue Lys-43
is surface exposed and located at the center of the basic region
on the TubR dimer (Fig. 3A). The K43A mutant also showed
no binding to TubR, supporting the notion that the continuous
basic patch of TubR represents its DNA-binding surface.
To gain insight into the structural mechanism of DNA binding,
a DNA duplex was docked onto the basic patch of the TubR di-
Fig. 1.
B. thuringiensis pBtoxis TubR structure. (A) One TubR subunit is red
and the other cyan. Secondary structural elements and N and C termini are
labeled. (B) Superimposition of one subunit of TubR (Red) onto a S. aureus
CzrA subunit (Yellow). Regions with different structures are labeled. (C) Same
superimposition as B showing the location of the other subunit in the TubR
and CzrA dimers after one subunit is overlaid. A–C are in the same orienta-
tion to highlight differences. Figs. 1 A–C, 2 C and D, 3 A, C, and E, 4B, and 5B
were made by using PyMOL (31).
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mer by using the location of the mutations that affected DNA
binding as a guide (Fig. 3C). This model revealed that the wings
are positioned to interact with successive minor grooves, with
either the bases or the phosphate backbone depending on the
ability of the DNA to deform. In the model TubR interacts with
a minimum of 14 bp of DNA. However, the centromere bound by
Fig. 2.
The TubR “recognition helix” dimer. (A) Gel fil-
tration studies on TubR mutants S63W and S63R show-
ing that the S63W mutant remains dimeric whereas
S63R is >80% monomer. (B) Fluorescence polarization
studies examining the ability of WT TubR, S63R, and
S63W TubR mutants to bind iteronic DNA. Fluorescence
polarization units (millipolarization) and TubR concen-
tration (nM) are along the y and x axes, respectively.
The Kd of WT TubR for the centromere DNA is
8 2 nm. (C) Superimposition of WT TubR (Green) onto
the TubR S63W mutant structure (Tan). (D) Close-up of
the site of the S63W mutation in the expanded TubR
S63W dimer showing stacking interactions between
the twofold related tryptophans.
Fig. 3.
TubR-DNA binding. (A) Electrostatic surface potential of the TubR dimer. Blue and red represent electropositive and electronegative regions, respec-
tively. (Left) The electronegative side of the TubR dimer, and the side on the right is the electropositive side. Labeled on the left side are the locations of the
mutated residues. (B) FP binding isotherms showing the DNA binding of WT TubR and the K43A, R74A, R77A, and K79A mutants. Fluorescence polarization
units (millipolarization) and TubR concentration (nM) are along the y and x axes, respectively. (C) TubR-DNA model showing TubR electrostatic potential.
(D) Stoichiometry of TubR (subunit) binding showing titration curve of TubR into the 48-mer iteron resulting in a molar ratio of TubR subunit to DNA of
eight (or four) dimers. (E) Left: Ribbon diagram of the TubR-DNA model with the recognition helices colored yellow. Right: Ribbon diagram showing a canonical
HTH–DNA interaction (the λ repressor-DNA complex) with the recognition helices colored yellow (32).
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TubR consists of four 12-bp sites with the consensus T(T/A)(T/A)
(C/A)(G/A)GTTTA(A/C)(A/C) (9). Thus, we used FP to ascer-
tain the binding stoichiometry of TubR for its 48-bp centromere.
As shown in Fig. 3D, eight TubR subunits or four TubR dimers
bind the 48-mer centromere, consistent with a dimer of TubR
binding each palindrome. Thus, either TubR distorts its DNA
or the TubR dimers bind with some degree of overlap on their
DNA sites perhaps imparting cooperativity, as observed in other
centromere-binding protein–DNA interactions (13, 14). In addi-
tion to the insertion of the wings, a striking outcome of the mod-
eling was the finding that the N termini of the recognition helices,
which interact with each other in a parallel, coiled-coil-like man-
ner, are in position to insert into a single major groove. Structures
of HTH proteins bound to DNA have thus far shown that the
recognition helices insert singly into successive major grooves
by using residues in the first few turns or the central portion
of the recognition helix to contact the DNA bases (Fig. 3E). Thus,
in the TubR-DNA model, the HTH–DNA interaction is drama-
tically different from any displayed previously by a HTH protein.
TubZ Binds TubR-DNA. A characteristic feature exhibited by
partition centromere-binding proteins is the ability to bind their
partner NTPase (13, 14). To determine if TubR binds TubZ, we
utilized a FP assay and found that full length (FL) TubZ bound
avidly to the TubR-centromere complex (Fig. 4A). However,
unlike other partition systems in which the NTPase must be com-
plexed with nucleotide to bind its centromere-binding protein,
the interaction of TubR with TubZ did not require the presence
of GTP-Mg2þ. Previous studies have shown that the C-terminal
regions of tubulin and FtsZ mediate key binding events with their
target proteins (24–26). We noted that the terminal region of
TubZ, consisting of residues 407–484, is the most divergent region
between TubZ proteins and between TubZ and tubulin/FtsZ pro-
teins, suggesting that it may be similarly utilized and bind TubR.
To test this hypothesis, we constructed various TubZ truncations,
TubZ(1-407), TubZ(1-442), TubZ(1-460), and TubZ(1-470), and
examined the ability of each protein to bind TubR-DNA. TubZ
(1-407) showed no binding to TubR, whereas the remaining trun-
cation mutants bound weakly to TubR-DNA (Fig. 4A). Thus, the
data indicate that the last 14 amino acids of TubZ are critical for
the ability of TubZ to form a tight interaction with TubR but that
residues 408–470 also play an important role in this interaction.
These data demonstrate that TubR acts as a partition partner for
TubZ, linking it to pBtoxis plasmid DNA. Although TubZ has
been shown to form polymers in a GTP-dependent manner,
the TubZ protein displays limited sequence similarity to tubu-
lin/FtsZ, suggesting potential differences in TubZ and tubulin/
FtsZ structures (7–9). To gain insight into TubZ function, we next
determined structures of B. thuringiensis pBtoxis TubZ.
Structure of TubZ. Crystallization of FLTubZ was not successful, in
either its apo form or bound to guanine nucleotides. We noted that
FL TubZ degraded over time whereby C-terminal residues were
proteolyzed. Therefore, truncated TubZ proteins were utilized
in crystallization trials, and crystals were obtained of apo TubZ
(1-428) and the structure solved by MAD (Table S2). The model
consists of residues 1–79 and 91–404 and has an Rwork∕Rfree of
21.4∕24.9% to 2.3-Å resolution (Fig. 4B). No discernible oligomer-
ization of apo TubZ(1-428) was observed in the crystal packing,
and gel filtration analyses confirmed that it is monomeric
(Fig. S2). The overall TubZ structure can be divided into two main
domains: an N domain (residues 25–235) and a C domain (resi-
dues 258–377). These domains are connected by a long, core helix,
H7. The TubZ N domain has a Rossman fold and consists of six
parallel β-strands with topology 3-2-1-4-5-6. The resulting β-sheet
is sandwiched by five α-helices, with two helices on one side and
three on the other. The C domain consists of four β-strands with
the topology 1-4-2-3. The C-domain β-strands are arranged nearly
perpendicular to those in the N domain. In addition to these main
protein domains, there are two helices: one at the N terminus, H0,
and a long helix at the C terminus, H11 (Fig. 4B). Database
searches showed that TubZ indeed belongs to the tubulin/FtsZ fa-
mily of proteins and is similar to both eukaryotic and prokaryotic
members of the family; TubZ can be optimally superimposed with
rmsds of 3.4 Å onto both bovine α tubulin and Pseudomonas
aeruginosa FtsZ (1, 5). Whereas the two-domain architectures
of tubulin, FtsZ, and TubZ are similar in overall structure, the
extreme N- and C-terminal regions of these proteins are very di-
vergent (Fig. S3 A–D).
N-Terminal and C-Terminal Differences in TubZ, FtsZ, and Tubulin: Im-
plications for Polymer Formation and Target Protein Binding. Tubulin
proteins do not contain significant N-terminal extensions,
whereas FtsZ proteins from different organisms show structural
variability within their N-terminal regions. For instance, in the
Escherichia coli FtsZ structure the N-terminal residues are disor-
dered, whereas Methanococcus jannaschii FtsZ has an extra
N-terminal helix, H0, which is flexibly attached to the body of
the protein and has been captured in multiple orientations (5).
Although H0 is not conserved in FtsZ proteins, one M. jannaschii
FtsZ structure revealed a semicontinuous polymer in the crystal,
thought to closely represent in vivo protofilaments, which utilizes
H0 in subunit-subunit contacts (4). This finding suggests that
the flexibly attached H0 is stabilized in a specific orientation
by protofilament formation, at least in the M. jannaschii protein.
The TubZ H0 helix extends in the opposite direction compared to
that of the protofilament stabilized FtsZ H0 helix. Moreover, in
TubZ, H0 is not flexibly attached to the N domain but is tightly
anchored to the C domain through numerous interactions with
the core helix and C-domain residues. The large number of inter-
actions involving H0, and the fact that it covers what would other-
wise be a surface exposed hydrophobic patch, indicate that the
TubZ H0 does not undergo conformational changes during
protofilament formation and is important for the general fold
of TubZ (Fig. S3 A and C).
Data suggest that FtsZ and tubulin form protofilaments with
similar longitudinal contacts (4). However, the TubZ structure
reveals key differences, primarily in its C-domain and C-terminal
regions, which suggest that it forms protofilaments distinct from
those formed by tubulin/FtsZ. A notable difference is the struc-
ture of loop 7 (L7). This loop inserts into the adjacent subunit
providing the key catalytic residues required for GTP hydrolysis.
In tubulin/FtsZ proteins, L7 has the consensus GXXNXDXAD.
In TubZ, the loop is very divergent in conformation compared
Fig. 4.
TubZ interacts with TubR-DNA and contains a tubulin/FtsZ fold. (A) FP
assay measuring binding of FL TubZ, TubZ(1-470), TubZ(1-460), TubZ(1-442),
and TubZ(1-407) to TubR-DNA. Below is the control (TubZ titrated into DNA
alone). Millipolarization units and TubZ concentration (nM) are along the y
and x axes, respectively. (B) TubZ(1-428) structure. The N domain or GTP-bind-
ing domain is colored salmon and the C domain purple. The interdomain
helix, H7, is red. TubZ also contains an N-terminal helix, H0 (Yellow), and
a C-terminal helix, H11 (White).
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to that in FtsZ/tubulin and consists of the sequence 256-
DNVTYDPSD-266. In addition to the N-terminal region, the
extreme C-terminal extensions of tubulin, FtsZ, and TubZ are
structurally divergent (Fig. S3B). In FtsZ, the C-terminal region
forms a small, two-stranded β-sheet and continues into an ex-
tended region that is involved in binding adaptor proteins such
as FtsA and ZipA (6, 25). By contrast, the C-terminal regions
of tubulin proteins consist of a two-helix bundle followed by an
extended region. Like FtsZ, however, these regions interact with
numerous target proteins such as the microtubule-associated pro-
teins (MAPs) (24). Consistent with this, the C-terminal regions of
tubulin have been shown to face the outside of the microtubule. A
characteristic feature of the extreme C-terminal extensions of
tubulin proteins is their highly acidic nature (3). This acidic region
has been shown to be critical for binding to several MAPs that
harbor a substantial basic character, such as tau, MAP2, and
MAP4 (24, 27).
The TubZ C-terminal region is also helical, but it contains a
single, long helix. Notably, the TubZ-tubulin overlay shows that
the long C-terminal helix of TubZ would dramatically clash with
the adjacent subunit in a polymer, providing support for the no-
tion that TubZ forms protofilaments different from tubulin/FtsZ
(Figs. S3B and S4). Interestingly, and in contrast to tubulin pro-
teins, the flexible C-terminal region of TubZ that follows H11 is
highly basic, in particular the last 14 residues. We have shown that
these residues play a central role in TubR binding (Fig. 4A). TubR
uses its electropositive face for DNA binding, leaving exposed its
opposite face for TubZ interaction. Notably, this exposed face is
strongly electronegative and hence would complement the basic
C-terminal tail of TubZ (Fig. 3A).
Tubulin/FtsZ protofilaments combine to form higher-order
structures. In tubulin, the protofilaments interact in a parallel
manner to form microtubules. Central to microtubule formation
are lateral contacts between protofilaments from the so-called M
loop, between H10 and S9. In tubulin, this loop is composed of 13
residues (1–3). The corresponding loop is much shorter in FtsZ
proteins, consistent with the fact that FtsZ does not form tubulin
microtubule-like structures (5, 28–30). In TubZ, the M loop is
even shorter than in FtsZ, spanning only four residues. In fact,
the TubZ/tubulin overlay shows that the side of the molecule con-
taining the M loop is the most divergent between these proteins.
These combined findings suggest that TubZ not only forms pro-
tofilaments with distinct longitudinal contacts compared to FtsZ
and tubulin, but it also does not form tubulin-like microtubule
structures.
TubZ Interactions with Guanine Nucleotides. Consistent with TubZ
being a member of the tubulin/FtsZ family, our isothermal titra-
tion calorimetry (ITC) studies showed that TubZ binds guanine
nucleotides with high affinity; Kds for GTP-γ-S and GDP were
∼0.69 and 26 μM, respectively (Fig. 5A). We next determined
the structure of the TubZ-GTP-γ-S complex by soaking GTP-γ-S
into preformed TubZ(1-428) crystals. The TubZ-GTP-γ-S struc-
ture contains TubZ residues 1–79 and 91–404, one GTP-γ-S,
and has Rwork∕Rfree ¼ 21.8%∕25.5% (Fig. 5B and Table S2).
The structure shows that TubZ binds GTP-γ-S in the same GTP
binding pocket as tubulin/FtsZ (1–5). Comparison of the apo
and GTP-γ-S bound TubZ structures indicated that, like FtsZ,
guanine nucleotide binding does not lead to significant conforma-
tional changes (5). The phosphate binding pocket is formed by
two of the most highly conserved regions between TubZ and
tubulin/FtsZ called loops 1 and 4 (L1 and L4) (1–3). L1 contacts
the GTP-γ-S α- and β-phosphate groups via the Gln-32 and Lys-33
amide nitrogens. The L1 region of FtsZ and tubulin contain the
sequences GQ(A/G)G and GQCG, respectively, whereas in TubZ
the motif is 31-GQKG-34. However, the alanine/glycine and
cysteine residues in FtsZ and tubulin do not contact the bound
nucleotide; the TubZ Lys-33 side chain makes stacking interac-
tions with the guanine base (Fig. 5B). L4 represents the so-called
signature motif [GGGTG(T/S)G], which serves as an identifier of
tubulin/FtsZ family members. Like FtsZ and tubulin, the L4
region of TubZ-GTP-γ-S makes phosphate interactions via its
glycine amide nitrogens. Whereas L1 and L4 residues of the N
domain mediate phosphate contacts, the GTP-γ-S guanine moiety
is specified from residues in the core helix, H5, and C domain. In
this regard, an important motif is loop 6 (L6). In FtsZ and tubulin,
L6 has the consensus (F/Y)XXX(N/D) and the conserved (F/Y)
residue functions in guanine base stacking. This region in TubZ,
236-WKXXXN-241, is in an altered conformation compared to
FtsZ and tubulin structures. Despite the presence of the trypto-
phan, which might be expected to interact with the guanine,
the side chain of Lys-237 instead stacks with the guanine ring.
Hence, in the TubZ-GTP-γ-S structure, the guanine base does
not interact with aromatic residues as in tubulin/FtsZ but is sand-
wiched between the aliphatic portions of two lysine side chains,
Lys-33 and Lys-237. Finally, two asparagine residues, Asn-213
and Asn-241, from L6 effectively read the guanine N2/N3 and
N1/O6 atoms, respectively, providing high specificity in TubZ’s in-
teraction with guanine nucleotides.
pBtoxis DNA Segregation: TubR Plasmid Partition Model. Our data
show that TubR binds to the flexible, C-terminal, basic region
of TubZ. The flexibility and location of the TubZ C-terminal
extension suggest that it is not required for polymerization and
thus may be exposed on the surface of TubZ filaments. Indeed,
negative stain EM images show that TubZ(1-407) forms polymers
in a GTP-dependent manner similar to the FL protein (Fig. S5).
Recent data suggesting that TubZ filaments are stabilized by
Fig. 5.
TubZ-guanine nucleotide interactions. (A) ITC binding isotherms
showing TubZ-GDP (Left) and TubZ-GTP-γ-S interaction (Right). (B) Left:
Overall structure of the TubZ-GTP-γ-S complex. β-strands are colored magen-
ta and helices cyan, and the GTP-γ-S molecule is shown as cpk. Right: Close-up
view of the GTP binding pocket with the initial Fo-Fc electron density map
(Blue Mesh), contoured at 4.5σ, and calculated before the GTP-γ-S was
included in refinement.
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the presence of a GTP cap and undergo treadmilling are consis-
tent with the notion that TubZ displays tubulin-like polymer dy-
namics (12). Thus, on the basis of the combined data, we suggest
a model for TubR/TubZ mediated pBtoxis plasmid segregation
shown in Fig. 6. In this model, multiple TubR dimers first bind
to the iteronic DNA on the pBtoxis plasmid leading to the crea-
tion of a high local concentration of TubR, which can recruit a
TubZ polymer, likely by interactions between the acidic TubR
dimer face and the basic C-terminal TubZ region. Importantly,
this interaction serves to attach the pBtoxis plasmid to the TubZ
polymer, which undergoes treadmilling, adding subunits at the þ
end and losing subunits at the −end. The bound TubR-pBtoxis
can be handed off from the −end to the molecules in the growing
þ end, leading to the transport of the pBtoxis plasmid to the cell
pole. Interestingly, it has been shown that once TubZ polymers
reach and interact with the cell pole, they bend around the curved
pole and continue growing in the other direction (7). The force of
the interaction with the membrane likely causes the release of
TubR-pBtoxis, the net result being transport of pBtoxis to the cell
pole. Of course, this model is simplified and many questions re-
main. For example, how directionality is achieved and how the
replicated plasmids are driven to opposite cell poles is not clear.
However, given the large size of the pBtoxis plasmid (8), it may be
that only one TubR-pBtoxis “tram” can be bound at once by the
rapidly treadmilling TubZ polymer and that, once one such a tram
is unloaded after reaching the cell pole, another engages when the
now reversed polymer treadmills toward the opposite cell pole.
Materials and Methods Summary
Detailed methods are provided in SI Materials and Methods.
Briefly, the tubR and tubZ genes were codon optimized (for
E. coli expression), subcloned into pET15b, expressed, and pur-
ified. WT TubR crystals were grown with NaCl and phosphate.
TubR S63W was crystallized with PEG and ethylene glycol and
TubZ with sodium formate. Detailed assay conditions for FP,
ITC, electron microscopy, and gel filtration are provided in
SI Materials and Methods.
ACKNOWLEDGMENTS. This work was supported by the Burroughs Wellcome
Career Development Award 992863 and National Institutes of Health Grant
GM074815 (to M.A.S.).
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Fig. 6.
pBtoxis DNA partition model. In the first step, TubR, which is bound
to its centromere on one of the replicated pBtoxis plasmids, contacts the TubZ
C-terminal region (indicated by lines pointing from the TubZ “circles”) in a
treadmilling TubZ polymer. TubZ subunits are lost from the −end and are
added to the þ end. TubR is pulled along the growing polymer by its
TubR-TubZ interaction until it reaches the cell pole and is knocked off when
it comes into contact with the membrane at the cell pole. TubZ reverses
direction and may pick up the other TubR-pBtoxis complex and deliver it
similarly to the opposite cell pole.
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|
3M8F
|
Protein structure of type III plasmid segregation TubR mutant
|
Plasmid protein TubR uses a distinct mode of HTH-
DNA binding and recruits the prokaryotic tubulin
homolog TubZ to effect DNA partition
Lisheng Ni, Weijun Xu, Muthiah Kumaraswami, and Maria A. Schumacher1
Department of Biochemistry and Molecular Biology, University of Texas, M. D. Anderson Cancer Center, Unit 1000, Houston, TX 77030
Edited by Robert T. Sauer, Massachusetts Institute of Technology, Cambridge, MA, and approved May 11, 2010 (received for review March 22, 2010)
The segregation of plasmid DNA typically requires three elements:
a DNA centromere site, an NTPase, and a centromere-binding
protein. Because of their simplicity, plasmid partition systems
represent tractable models to study the molecular basis of DNA
segregation. Unlike eukaryotes, which utilize the GTPase tubulin
to segregate DNA, the most common plasmid-encoded NTPases
contain Walker-box and actin-like folds. Recently, a plasmid stabi-
lity cassette on Bacillus thuringiensis pBtoxis encoding a putative
FtsZ/tubulin-like NTPase called TubZ and DNA-binding protein
called TubR has been described. How these proteins collaborate
to impart plasmid stability, however, is unknown. Here we show
that the TubR structure consists of an intertwined dimer with a
winged helix-turn-helix (HTH) motif. Strikingly, however, the TubR
recognition helices mediate dimerization, making canonical HTH–
DNA interactions impossible. Mutagenesis data indicate that a
basic patch, encompassing the two wing regions and the N termini
of the recognition helices, mediates DNA binding, which indicates
an unusual HTH–DNA interaction mode in which the N termini of
the recognition helices insert into a single DNA groove and the
wings into adjacent DNA grooves. The TubZ structure shows that
it is as similar structurally to eukaryotic tubulin as it is to bacterial
FtsZ. TubZ forms polymers with guanine nucleotide-binding
characteristics and polymer dynamics similar to tubulin. Finally,
we show that the exposed TubZ C-terminal region interacts with
TubR-DNA, linking the TubR-bound pBtoxis to TubZ polymerization.
The combined data suggest a mechanism for TubZ-polymer pow-
ered plasmid movement.
T
he cytoskeletons of eukaryotic cells are constructed of three
primary elements: actin, tubulin, and intermediate filaments.
Although it had long been presumed that the proteins forming
these elements were absent in prokaryotes, it is now known that
prokaryotes contain structural homologs to all three components.
These prokaryotic proteins appear to carry out distinct functions
compared to their eukaryotic counterparts; however, their roles
are similar enough to indicate a likely common ancestor. The best
known prokaryotic homolog of tubulin is FtsZ. Both FtsZ and
tubulin form long filamentous structures by head to tail associa-
tion in a manner regulated by GTP, which binds between adjacent
subunits (1–4). However, unlike tubulin, FtsZ does not function
in DNA segregation but rather cell division. Specifically, it forms
a cytokinetic ring called the Z ring at midcell, which mediates
septation (5, 6). Recently, however, prokaryotic proteins encoded
on large plasmids harbored in bacilli showing 15–20% sequence
similarity to both FtsZ and tubulin have been identified and
dubbed TubZ (7–12). Studies showed that the Bacillus thuringien-
sis TubZ protein from the pBtoxis plasmid is essential for plasmid
DNA segregation.
DNA segregation of most low copy number plasmids is carried
out by specific partition (par) systems. These systems require only
three elements: a centromere DNA site, a centromere-binding
protein, and a partition NTPase (13, 14). Partition systems have
been classified into two main types on the basis of the kind
of NTPase present (15). Type I systems contain NTPases with
deviant Walker A-type ATPase folds, whereas type II systems uti-
lize actin-like NTPases. Interestingly, both types of NTPases form
polymers in NTP-dependent manners that are implicated to play
a role in plasmid DNA separation (16–19). The recent discovery
of TubZ NTPases has led to the designation of “type III” par sys-
tems (13, 14). The best studied of these systems is that found on
the pBtoxis plasmid in B. thuringiensis. This plasmid stability sys-
tem is represented by an operon encoding two proteins: ORF156
(TubZ) and ORF157 (TubR) (7–9, 11). TubR is a 11.6 kDa DNA-
binding protein that shows no sequence homology to any known
protein. Studies showed that TubR binds a 48-bp centromere con-
taining four repeat sites in the pBtoxis plasmid and also autore-
gulates tubRZ transcription (8, 9). TubZ is a 54.4 kDa protein that
can assemble into filaments in a GTP-dependent manner (12).
Both proteins were found to be required for plasmid stability
(9). However, how the TubR and TubZ proteins work together
to effect pBtoxis plasmid segregation is not known. To gain insight
into the molecular mechanism utilized by these proteins in DNA
segregation, we carried out structural and biochemical studies on
the pBtoxis TubR and TubZ proteins. The TubR structure reveals
that it employs a helix-turn-helix (HTH) motif in a previously un-
described manner to bind DNA. TubZ contains a tubulin/FtsZ
fold but has structural distinctions from these proteins indicating
that it forms distinct protofilaments. TubR binds the flexible
C-terminal region of TubZ, thus attaching the TubZ filament
to the pBtoxis plasmid, providing a mechanism for plasmid move-
ment and, ultimately, segregation.
Results and Discussion
Overall Structure of pBtoxis TubR. The crystal structure of the 107-
residue pBtoxis TubR protein was solved to 2.0-Å resolution by
selenomethionine multiple wavelength anomalous diffraction
(MAD) methods (Table S1). The structure contains two TubR
molecules in the crystallographic asymmetric unit and consists
of residues 6–102 of one subunit and 4–100 of the second subunit,
and has Rwork∕Rfree ¼ 23.8%∕27.0%. The TubR structure forms a
highly
intertwined
dimer
with
dimensions
30 × 30 × 60 Å3
(Fig. 1A). Each TubR subunit has the topology β1-α1-α2-α3-α4-β2-
β3-α5, which is similar to winged HTH motifs found in a number
of DNA-binding proteins in both prokaryotes and eukaryotes
(20). In TubR, α3-α4 forms the HTH motif and the loop between
Author contributions: M.A.S. designed research; L.N., W.X., M.K., and M.A.S. performed
research; M.A.S. analyzed data; and M.A.S. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Freely available online through the PNAS open access option.
Data deposition: The atomic coordinates and structure factor amplitudes for the WT TubR
(C2), WT TubR (I222), TubR(S63W), TubZ, and TubZ-GTP-γ-S structures have been
deposited with the Protein Data Bank, www.pdb.org (PDB ID codes 3M8E, 3M9A,
3M8F, 3M8K, and 3M89).
1To whom correspondence may be addressed. E-mail: maschuma@mdanderson.org.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1003817107/-/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.1003817107
PNAS ∣June 29, 2010 ∣vol. 107 ∣no. 26 ∣11763–11768
BIOCHEMISTRY
β2 and β3, the wing. Indeed, each TubR subunit shows the stron-
gest structural similarity to members of the ArsR family of
prokaryotic winged helix transcription regulators, in particular
the Staphylococcus aureus CzrA protein (21, 22). Superposition
of one subunit of TubR onto that of CzrA results in a rmsd of
2.7 Å. This similarity includes the core regions of the winged
HTH, because the loops and N-terminal regions of the proteins
are structurally distinct. For example, TubR contains a β-strand in
its N-terminal region compared to a long helix in the CzrA struc-
ture (Fig. 1B). This structural similarity initially suggested that
TubR may be a member of the ArsR family of proteins. However,
the arrangement of the TubR dimer was found to be strikingly
different from the dimer organization exhibited by the ArsR
proteins (Fig. 1C).
ArsR family members are involved in metal-regulated tran-
scription processes whereby they act as repressors in their apo
forms and are induced off their DNA sites upon metal binding
(22). The specific dimer structures of the ArsR proteins are cri-
tical for creation of their metal-binding motifs. Not only does
TubR form a very different dimer from the ArsR proteins, it also
does not harbor any of their metal-binding signatures nor does it
contain any other characterized metal-binding motif. Consistent
with this, we find that the addition of metals has no effect on TubR
DNA binding. Dimerization of the ArsR proteins is imparted by
residues from the N-terminal regions, α1 and α5, which, impor-
tantly, leaves its recognition helices exposed for DNA interaction
(22). By contrast, the TubR dimer is formed primarily by contacts
between its twofold related “recognition helices” α4 and α4·. This
interaction results in a near complete burial of these helices, leav-
ing only the N-terminal residues exposed to solvent. Whereas the
α4 and α4· interaction creates the dimer core, the dimer is further
stabilized by interactions between the twofold related β1 strands,
which swap to form an antiparallel β-sheet. Residues from α1 and
α5 interact with β1 to further seal the top of the dimer. The dimer
interface formed by these interactions is predominantly hydropho-
bic and buries a large 1;200 Å2 of subunit surface from solvent.
TubR Forms a “Recognition Helix Dimer”: Implications for DNA Bind-
ing. Gel filtration studies on TubR confirmed that it is a dimer in
solution. However, the finding from the structure that the TubR
α4 recognition helices are buried in the dimer core has important
implications in terms of its DNA-binding mechanism. Indeed, it
suggests that, although TubR contains a structurally canonical
HTH, it is not utilized for DNA binding in a manner typical
of HTH proteins. A second crystal form (I222) of TubR, which
was solved to 2.5-Å resolution, revealed the same TubR dimer.
The presence of the identical dimer in two different crystal forms
and its large buried surface area supports that the dimer observed
in the crystal structures is physiologically relevant. However, to
test this, we mutated residues within the recognition helices that
the structure indicates are critical for dimerization and assayed
the ability of the mutant proteins to dimerize via gel filtration.
Specifically, we mutated Ser-63 and Ala-67 individually to tryp-
tophan and arginine.
The structure shows that residues occupying positions 63 and
67 must be small and largely hydrophobic to permit the proper
packing of the α4 helices in the dimer (Fig. S1A). Hence, the in-
troduction of the bulky side chain of tryptophan and, in particu-
lar, the large as well as charged side chain of arginine would be
predicted to be highly disruptive to dimerization. Gel filtration
analyses on purified mutant proteins clearly showed that the ar-
ginine mutants exist primarily as monomers in solution (>80%),
whereas the tryptophan mutants were able to maintain the di-
meric state (Fig. 2A and Fig. S1B). However, all mutant proteins
showed reduced or loss of DNA-binding activity as ascertained by
fluorescence polarization (FP) studies, which examined TubR
protein binding to its centromere site (Fig. 2B and Fig. S1 C
and D) (9). The fact that the monomeric mutants were severely
impaired in DNA binding was not surprising. However, the
finding that the tryptophan mutants, which were largely dimeric,
displayed reduced DNA-binding activity suggested that their oli-
gomer structures might be altered. To address this issue, the struc-
ture of S63W TubR was solved to 2.8-Å resolution, resulting in
Rwork∕Rfree values of 20.4%∕26.6% (Table S1). The subunit struc-
ture of S63W TubR is essentially identical to that of WT TubR as
revealed by their superimposition (rmsd of 1.2 Å) (Fig. 2 C and
D). However, this single subunit overlay shows that the S63W
TubR dimer, although the same as the WT in general arrange-
ment, is forced into a more open oligomer conformation in which
one subunit is rotated 20° away from its dimer mate compared to
WT. This rotation is required to accommodate the bulky S63W
side chains (Fig. 2D). The N-terminal β1–β1′-sheet interaction
appears to play a key role in holding the TubR subunits together.
In addition, the tight stacking of the twofold related Trp63 indole
groups (3.5 Å) provides a compensatory interaction that, com-
bined with the β1–β1′ interaction, apparently permits retention
of the dimer state, indicating why the TubR tryptophan mutants
were able to maintain the dimer state, albeit an altered dimer
state relative to WT (Fig. 2 C and D). By contrast, the TubR
arginine mutations, which introduced both bulk and charge with-
in the predominantly hydrophobic dimer interface, were highly
destabilizing for dimerization. In addition, the finding that the
S63W TubR mutant forms an altered dimer explains the severe
effect on DNA binding because a correct dimer orientation is
likely essential for binding to its palindromic DNA sites (9).
TubR-DNA Model. Because all but the N-terminal residues of the
TubR recognition helices are buried in the dimer interface, TubR
must use a different mode of DNA binding than the ArsR or other
HTH containing proteins (23). Examination of surface electro-
statics of TubR reveals that one face of the protein is electro-
negative, whereas the other is strongly electropositive (Fig. 3A).
Notably, the positive region is composed of one large and contig-
uous basic patch. Basic residues in this region correspond to Arg-
74, Arg-77, and Lys-79, in the wing and Lys-43, which is on α3, the
helix preceding α4 in the HTH motif. These residues were mutated
singly to alanine to examine their roles in DNA binding (Fig. 3B).
FP experiments showed that mutation of the basic wing residues
resulted in either a complete (R74A and R77A) or nearly com-
plete (K79A) abrogation of DNA binding, indicating that the
wings play a major role in TubR DNA binding. Residue Lys-43
is surface exposed and located at the center of the basic region
on the TubR dimer (Fig. 3A). The K43A mutant also showed
no binding to TubR, supporting the notion that the continuous
basic patch of TubR represents its DNA-binding surface.
To gain insight into the structural mechanism of DNA binding,
a DNA duplex was docked onto the basic patch of the TubR di-
Fig. 1.
B. thuringiensis pBtoxis TubR structure. (A) One TubR subunit is red
and the other cyan. Secondary structural elements and N and C termini are
labeled. (B) Superimposition of one subunit of TubR (Red) onto a S. aureus
CzrA subunit (Yellow). Regions with different structures are labeled. (C) Same
superimposition as B showing the location of the other subunit in the TubR
and CzrA dimers after one subunit is overlaid. A–C are in the same orienta-
tion to highlight differences. Figs. 1 A–C, 2 C and D, 3 A, C, and E, 4B, and 5B
were made by using PyMOL (31).
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Ni et al.
mer by using the location of the mutations that affected DNA
binding as a guide (Fig. 3C). This model revealed that the wings
are positioned to interact with successive minor grooves, with
either the bases or the phosphate backbone depending on the
ability of the DNA to deform. In the model TubR interacts with
a minimum of 14 bp of DNA. However, the centromere bound by
Fig. 2.
The TubR “recognition helix” dimer. (A) Gel fil-
tration studies on TubR mutants S63W and S63R show-
ing that the S63W mutant remains dimeric whereas
S63R is >80% monomer. (B) Fluorescence polarization
studies examining the ability of WT TubR, S63R, and
S63W TubR mutants to bind iteronic DNA. Fluorescence
polarization units (millipolarization) and TubR concen-
tration (nM) are along the y and x axes, respectively.
The Kd of WT TubR for the centromere DNA is
8 2 nm. (C) Superimposition of WT TubR (Green) onto
the TubR S63W mutant structure (Tan). (D) Close-up of
the site of the S63W mutation in the expanded TubR
S63W dimer showing stacking interactions between
the twofold related tryptophans.
Fig. 3.
TubR-DNA binding. (A) Electrostatic surface potential of the TubR dimer. Blue and red represent electropositive and electronegative regions, respec-
tively. (Left) The electronegative side of the TubR dimer, and the side on the right is the electropositive side. Labeled on the left side are the locations of the
mutated residues. (B) FP binding isotherms showing the DNA binding of WT TubR and the K43A, R74A, R77A, and K79A mutants. Fluorescence polarization
units (millipolarization) and TubR concentration (nM) are along the y and x axes, respectively. (C) TubR-DNA model showing TubR electrostatic potential.
(D) Stoichiometry of TubR (subunit) binding showing titration curve of TubR into the 48-mer iteron resulting in a molar ratio of TubR subunit to DNA of
eight (or four) dimers. (E) Left: Ribbon diagram of the TubR-DNA model with the recognition helices colored yellow. Right: Ribbon diagram showing a canonical
HTH–DNA interaction (the λ repressor-DNA complex) with the recognition helices colored yellow (32).
Ni et al.
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TubR consists of four 12-bp sites with the consensus T(T/A)(T/A)
(C/A)(G/A)GTTTA(A/C)(A/C) (9). Thus, we used FP to ascer-
tain the binding stoichiometry of TubR for its 48-bp centromere.
As shown in Fig. 3D, eight TubR subunits or four TubR dimers
bind the 48-mer centromere, consistent with a dimer of TubR
binding each palindrome. Thus, either TubR distorts its DNA
or the TubR dimers bind with some degree of overlap on their
DNA sites perhaps imparting cooperativity, as observed in other
centromere-binding protein–DNA interactions (13, 14). In addi-
tion to the insertion of the wings, a striking outcome of the mod-
eling was the finding that the N termini of the recognition helices,
which interact with each other in a parallel, coiled-coil-like man-
ner, are in position to insert into a single major groove. Structures
of HTH proteins bound to DNA have thus far shown that the
recognition helices insert singly into successive major grooves
by using residues in the first few turns or the central portion
of the recognition helix to contact the DNA bases (Fig. 3E). Thus,
in the TubR-DNA model, the HTH–DNA interaction is drama-
tically different from any displayed previously by a HTH protein.
TubZ Binds TubR-DNA. A characteristic feature exhibited by
partition centromere-binding proteins is the ability to bind their
partner NTPase (13, 14). To determine if TubR binds TubZ, we
utilized a FP assay and found that full length (FL) TubZ bound
avidly to the TubR-centromere complex (Fig. 4A). However,
unlike other partition systems in which the NTPase must be com-
plexed with nucleotide to bind its centromere-binding protein,
the interaction of TubR with TubZ did not require the presence
of GTP-Mg2þ. Previous studies have shown that the C-terminal
regions of tubulin and FtsZ mediate key binding events with their
target proteins (24–26). We noted that the terminal region of
TubZ, consisting of residues 407–484, is the most divergent region
between TubZ proteins and between TubZ and tubulin/FtsZ pro-
teins, suggesting that it may be similarly utilized and bind TubR.
To test this hypothesis, we constructed various TubZ truncations,
TubZ(1-407), TubZ(1-442), TubZ(1-460), and TubZ(1-470), and
examined the ability of each protein to bind TubR-DNA. TubZ
(1-407) showed no binding to TubR, whereas the remaining trun-
cation mutants bound weakly to TubR-DNA (Fig. 4A). Thus, the
data indicate that the last 14 amino acids of TubZ are critical for
the ability of TubZ to form a tight interaction with TubR but that
residues 408–470 also play an important role in this interaction.
These data demonstrate that TubR acts as a partition partner for
TubZ, linking it to pBtoxis plasmid DNA. Although TubZ has
been shown to form polymers in a GTP-dependent manner,
the TubZ protein displays limited sequence similarity to tubu-
lin/FtsZ, suggesting potential differences in TubZ and tubulin/
FtsZ structures (7–9). To gain insight into TubZ function, we next
determined structures of B. thuringiensis pBtoxis TubZ.
Structure of TubZ. Crystallization of FLTubZ was not successful, in
either its apo form or bound to guanine nucleotides. We noted that
FL TubZ degraded over time whereby C-terminal residues were
proteolyzed. Therefore, truncated TubZ proteins were utilized
in crystallization trials, and crystals were obtained of apo TubZ
(1-428) and the structure solved by MAD (Table S2). The model
consists of residues 1–79 and 91–404 and has an Rwork∕Rfree of
21.4∕24.9% to 2.3-Å resolution (Fig. 4B). No discernible oligomer-
ization of apo TubZ(1-428) was observed in the crystal packing,
and gel filtration analyses confirmed that it is monomeric
(Fig. S2). The overall TubZ structure can be divided into two main
domains: an N domain (residues 25–235) and a C domain (resi-
dues 258–377). These domains are connected by a long, core helix,
H7. The TubZ N domain has a Rossman fold and consists of six
parallel β-strands with topology 3-2-1-4-5-6. The resulting β-sheet
is sandwiched by five α-helices, with two helices on one side and
three on the other. The C domain consists of four β-strands with
the topology 1-4-2-3. The C-domain β-strands are arranged nearly
perpendicular to those in the N domain. In addition to these main
protein domains, there are two helices: one at the N terminus, H0,
and a long helix at the C terminus, H11 (Fig. 4B). Database
searches showed that TubZ indeed belongs to the tubulin/FtsZ fa-
mily of proteins and is similar to both eukaryotic and prokaryotic
members of the family; TubZ can be optimally superimposed with
rmsds of 3.4 Å onto both bovine α tubulin and Pseudomonas
aeruginosa FtsZ (1, 5). Whereas the two-domain architectures
of tubulin, FtsZ, and TubZ are similar in overall structure, the
extreme N- and C-terminal regions of these proteins are very di-
vergent (Fig. S3 A–D).
N-Terminal and C-Terminal Differences in TubZ, FtsZ, and Tubulin: Im-
plications for Polymer Formation and Target Protein Binding. Tubulin
proteins do not contain significant N-terminal extensions,
whereas FtsZ proteins from different organisms show structural
variability within their N-terminal regions. For instance, in the
Escherichia coli FtsZ structure the N-terminal residues are disor-
dered, whereas Methanococcus jannaschii FtsZ has an extra
N-terminal helix, H0, which is flexibly attached to the body of
the protein and has been captured in multiple orientations (5).
Although H0 is not conserved in FtsZ proteins, one M. jannaschii
FtsZ structure revealed a semicontinuous polymer in the crystal,
thought to closely represent in vivo protofilaments, which utilizes
H0 in subunit-subunit contacts (4). This finding suggests that
the flexibly attached H0 is stabilized in a specific orientation
by protofilament formation, at least in the M. jannaschii protein.
The TubZ H0 helix extends in the opposite direction compared to
that of the protofilament stabilized FtsZ H0 helix. Moreover, in
TubZ, H0 is not flexibly attached to the N domain but is tightly
anchored to the C domain through numerous interactions with
the core helix and C-domain residues. The large number of inter-
actions involving H0, and the fact that it covers what would other-
wise be a surface exposed hydrophobic patch, indicate that the
TubZ H0 does not undergo conformational changes during
protofilament formation and is important for the general fold
of TubZ (Fig. S3 A and C).
Data suggest that FtsZ and tubulin form protofilaments with
similar longitudinal contacts (4). However, the TubZ structure
reveals key differences, primarily in its C-domain and C-terminal
regions, which suggest that it forms protofilaments distinct from
those formed by tubulin/FtsZ. A notable difference is the struc-
ture of loop 7 (L7). This loop inserts into the adjacent subunit
providing the key catalytic residues required for GTP hydrolysis.
In tubulin/FtsZ proteins, L7 has the consensus GXXNXDXAD.
In TubZ, the loop is very divergent in conformation compared
Fig. 4.
TubZ interacts with TubR-DNA and contains a tubulin/FtsZ fold. (A) FP
assay measuring binding of FL TubZ, TubZ(1-470), TubZ(1-460), TubZ(1-442),
and TubZ(1-407) to TubR-DNA. Below is the control (TubZ titrated into DNA
alone). Millipolarization units and TubZ concentration (nM) are along the y
and x axes, respectively. (B) TubZ(1-428) structure. The N domain or GTP-bind-
ing domain is colored salmon and the C domain purple. The interdomain
helix, H7, is red. TubZ also contains an N-terminal helix, H0 (Yellow), and
a C-terminal helix, H11 (White).
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Ni et al.
to that in FtsZ/tubulin and consists of the sequence 256-
DNVTYDPSD-266. In addition to the N-terminal region, the
extreme C-terminal extensions of tubulin, FtsZ, and TubZ are
structurally divergent (Fig. S3B). In FtsZ, the C-terminal region
forms a small, two-stranded β-sheet and continues into an ex-
tended region that is involved in binding adaptor proteins such
as FtsA and ZipA (6, 25). By contrast, the C-terminal regions
of tubulin proteins consist of a two-helix bundle followed by an
extended region. Like FtsZ, however, these regions interact with
numerous target proteins such as the microtubule-associated pro-
teins (MAPs) (24). Consistent with this, the C-terminal regions of
tubulin have been shown to face the outside of the microtubule. A
characteristic feature of the extreme C-terminal extensions of
tubulin proteins is their highly acidic nature (3). This acidic region
has been shown to be critical for binding to several MAPs that
harbor a substantial basic character, such as tau, MAP2, and
MAP4 (24, 27).
The TubZ C-terminal region is also helical, but it contains a
single, long helix. Notably, the TubZ-tubulin overlay shows that
the long C-terminal helix of TubZ would dramatically clash with
the adjacent subunit in a polymer, providing support for the no-
tion that TubZ forms protofilaments different from tubulin/FtsZ
(Figs. S3B and S4). Interestingly, and in contrast to tubulin pro-
teins, the flexible C-terminal region of TubZ that follows H11 is
highly basic, in particular the last 14 residues. We have shown that
these residues play a central role in TubR binding (Fig. 4A). TubR
uses its electropositive face for DNA binding, leaving exposed its
opposite face for TubZ interaction. Notably, this exposed face is
strongly electronegative and hence would complement the basic
C-terminal tail of TubZ (Fig. 3A).
Tubulin/FtsZ protofilaments combine to form higher-order
structures. In tubulin, the protofilaments interact in a parallel
manner to form microtubules. Central to microtubule formation
are lateral contacts between protofilaments from the so-called M
loop, between H10 and S9. In tubulin, this loop is composed of 13
residues (1–3). The corresponding loop is much shorter in FtsZ
proteins, consistent with the fact that FtsZ does not form tubulin
microtubule-like structures (5, 28–30). In TubZ, the M loop is
even shorter than in FtsZ, spanning only four residues. In fact,
the TubZ/tubulin overlay shows that the side of the molecule con-
taining the M loop is the most divergent between these proteins.
These combined findings suggest that TubZ not only forms pro-
tofilaments with distinct longitudinal contacts compared to FtsZ
and tubulin, but it also does not form tubulin-like microtubule
structures.
TubZ Interactions with Guanine Nucleotides. Consistent with TubZ
being a member of the tubulin/FtsZ family, our isothermal titra-
tion calorimetry (ITC) studies showed that TubZ binds guanine
nucleotides with high affinity; Kds for GTP-γ-S and GDP were
∼0.69 and 26 μM, respectively (Fig. 5A). We next determined
the structure of the TubZ-GTP-γ-S complex by soaking GTP-γ-S
into preformed TubZ(1-428) crystals. The TubZ-GTP-γ-S struc-
ture contains TubZ residues 1–79 and 91–404, one GTP-γ-S,
and has Rwork∕Rfree ¼ 21.8%∕25.5% (Fig. 5B and Table S2).
The structure shows that TubZ binds GTP-γ-S in the same GTP
binding pocket as tubulin/FtsZ (1–5). Comparison of the apo
and GTP-γ-S bound TubZ structures indicated that, like FtsZ,
guanine nucleotide binding does not lead to significant conforma-
tional changes (5). The phosphate binding pocket is formed by
two of the most highly conserved regions between TubZ and
tubulin/FtsZ called loops 1 and 4 (L1 and L4) (1–3). L1 contacts
the GTP-γ-S α- and β-phosphate groups via the Gln-32 and Lys-33
amide nitrogens. The L1 region of FtsZ and tubulin contain the
sequences GQ(A/G)G and GQCG, respectively, whereas in TubZ
the motif is 31-GQKG-34. However, the alanine/glycine and
cysteine residues in FtsZ and tubulin do not contact the bound
nucleotide; the TubZ Lys-33 side chain makes stacking interac-
tions with the guanine base (Fig. 5B). L4 represents the so-called
signature motif [GGGTG(T/S)G], which serves as an identifier of
tubulin/FtsZ family members. Like FtsZ and tubulin, the L4
region of TubZ-GTP-γ-S makes phosphate interactions via its
glycine amide nitrogens. Whereas L1 and L4 residues of the N
domain mediate phosphate contacts, the GTP-γ-S guanine moiety
is specified from residues in the core helix, H5, and C domain. In
this regard, an important motif is loop 6 (L6). In FtsZ and tubulin,
L6 has the consensus (F/Y)XXX(N/D) and the conserved (F/Y)
residue functions in guanine base stacking. This region in TubZ,
236-WKXXXN-241, is in an altered conformation compared to
FtsZ and tubulin structures. Despite the presence of the trypto-
phan, which might be expected to interact with the guanine,
the side chain of Lys-237 instead stacks with the guanine ring.
Hence, in the TubZ-GTP-γ-S structure, the guanine base does
not interact with aromatic residues as in tubulin/FtsZ but is sand-
wiched between the aliphatic portions of two lysine side chains,
Lys-33 and Lys-237. Finally, two asparagine residues, Asn-213
and Asn-241, from L6 effectively read the guanine N2/N3 and
N1/O6 atoms, respectively, providing high specificity in TubZ’s in-
teraction with guanine nucleotides.
pBtoxis DNA Segregation: TubR Plasmid Partition Model. Our data
show that TubR binds to the flexible, C-terminal, basic region
of TubZ. The flexibility and location of the TubZ C-terminal
extension suggest that it is not required for polymerization and
thus may be exposed on the surface of TubZ filaments. Indeed,
negative stain EM images show that TubZ(1-407) forms polymers
in a GTP-dependent manner similar to the FL protein (Fig. S5).
Recent data suggesting that TubZ filaments are stabilized by
Fig. 5.
TubZ-guanine nucleotide interactions. (A) ITC binding isotherms
showing TubZ-GDP (Left) and TubZ-GTP-γ-S interaction (Right). (B) Left:
Overall structure of the TubZ-GTP-γ-S complex. β-strands are colored magen-
ta and helices cyan, and the GTP-γ-S molecule is shown as cpk. Right: Close-up
view of the GTP binding pocket with the initial Fo-Fc electron density map
(Blue Mesh), contoured at 4.5σ, and calculated before the GTP-γ-S was
included in refinement.
Ni et al.
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the presence of a GTP cap and undergo treadmilling are consis-
tent with the notion that TubZ displays tubulin-like polymer dy-
namics (12). Thus, on the basis of the combined data, we suggest
a model for TubR/TubZ mediated pBtoxis plasmid segregation
shown in Fig. 6. In this model, multiple TubR dimers first bind
to the iteronic DNA on the pBtoxis plasmid leading to the crea-
tion of a high local concentration of TubR, which can recruit a
TubZ polymer, likely by interactions between the acidic TubR
dimer face and the basic C-terminal TubZ region. Importantly,
this interaction serves to attach the pBtoxis plasmid to the TubZ
polymer, which undergoes treadmilling, adding subunits at the þ
end and losing subunits at the −end. The bound TubR-pBtoxis
can be handed off from the −end to the molecules in the growing
þ end, leading to the transport of the pBtoxis plasmid to the cell
pole. Interestingly, it has been shown that once TubZ polymers
reach and interact with the cell pole, they bend around the curved
pole and continue growing in the other direction (7). The force of
the interaction with the membrane likely causes the release of
TubR-pBtoxis, the net result being transport of pBtoxis to the cell
pole. Of course, this model is simplified and many questions re-
main. For example, how directionality is achieved and how the
replicated plasmids are driven to opposite cell poles is not clear.
However, given the large size of the pBtoxis plasmid (8), it may be
that only one TubR-pBtoxis “tram” can be bound at once by the
rapidly treadmilling TubZ polymer and that, once one such a tram
is unloaded after reaching the cell pole, another engages when the
now reversed polymer treadmills toward the opposite cell pole.
Materials and Methods Summary
Detailed methods are provided in SI Materials and Methods.
Briefly, the tubR and tubZ genes were codon optimized (for
E. coli expression), subcloned into pET15b, expressed, and pur-
ified. WT TubR crystals were grown with NaCl and phosphate.
TubR S63W was crystallized with PEG and ethylene glycol and
TubZ with sodium formate. Detailed assay conditions for FP,
ITC, electron microscopy, and gel filtration are provided in
SI Materials and Methods.
ACKNOWLEDGMENTS. This work was supported by the Burroughs Wellcome
Career Development Award 992863 and National Institutes of Health Grant
GM074815 (to M.A.S.).
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Fig. 6.
pBtoxis DNA partition model. In the first step, TubR, which is bound
to its centromere on one of the replicated pBtoxis plasmids, contacts the TubZ
C-terminal region (indicated by lines pointing from the TubZ “circles”) in a
treadmilling TubZ polymer. TubZ subunits are lost from the −end and are
added to the þ end. TubR is pulled along the growing polymer by its
TubR-TubZ interaction until it reaches the cell pole and is knocked off when
it comes into contact with the membrane at the cell pole. TubZ reverses
direction and may pick up the other TubR-pBtoxis complex and deliver it
similarly to the opposite cell pole.
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|
3M8K
|
Protein structure of type III plasmid segregation TubZ
|
Plasmid protein TubR uses a distinct mode of HTH-
DNA binding and recruits the prokaryotic tubulin
homolog TubZ to effect DNA partition
Lisheng Ni, Weijun Xu, Muthiah Kumaraswami, and Maria A. Schumacher1
Department of Biochemistry and Molecular Biology, University of Texas, M. D. Anderson Cancer Center, Unit 1000, Houston, TX 77030
Edited by Robert T. Sauer, Massachusetts Institute of Technology, Cambridge, MA, and approved May 11, 2010 (received for review March 22, 2010)
The segregation of plasmid DNA typically requires three elements:
a DNA centromere site, an NTPase, and a centromere-binding
protein. Because of their simplicity, plasmid partition systems
represent tractable models to study the molecular basis of DNA
segregation. Unlike eukaryotes, which utilize the GTPase tubulin
to segregate DNA, the most common plasmid-encoded NTPases
contain Walker-box and actin-like folds. Recently, a plasmid stabi-
lity cassette on Bacillus thuringiensis pBtoxis encoding a putative
FtsZ/tubulin-like NTPase called TubZ and DNA-binding protein
called TubR has been described. How these proteins collaborate
to impart plasmid stability, however, is unknown. Here we show
that the TubR structure consists of an intertwined dimer with a
winged helix-turn-helix (HTH) motif. Strikingly, however, the TubR
recognition helices mediate dimerization, making canonical HTH–
DNA interactions impossible. Mutagenesis data indicate that a
basic patch, encompassing the two wing regions and the N termini
of the recognition helices, mediates DNA binding, which indicates
an unusual HTH–DNA interaction mode in which the N termini of
the recognition helices insert into a single DNA groove and the
wings into adjacent DNA grooves. The TubZ structure shows that
it is as similar structurally to eukaryotic tubulin as it is to bacterial
FtsZ. TubZ forms polymers with guanine nucleotide-binding
characteristics and polymer dynamics similar to tubulin. Finally,
we show that the exposed TubZ C-terminal region interacts with
TubR-DNA, linking the TubR-bound pBtoxis to TubZ polymerization.
The combined data suggest a mechanism for TubZ-polymer pow-
ered plasmid movement.
T
he cytoskeletons of eukaryotic cells are constructed of three
primary elements: actin, tubulin, and intermediate filaments.
Although it had long been presumed that the proteins forming
these elements were absent in prokaryotes, it is now known that
prokaryotes contain structural homologs to all three components.
These prokaryotic proteins appear to carry out distinct functions
compared to their eukaryotic counterparts; however, their roles
are similar enough to indicate a likely common ancestor. The best
known prokaryotic homolog of tubulin is FtsZ. Both FtsZ and
tubulin form long filamentous structures by head to tail associa-
tion in a manner regulated by GTP, which binds between adjacent
subunits (1–4). However, unlike tubulin, FtsZ does not function
in DNA segregation but rather cell division. Specifically, it forms
a cytokinetic ring called the Z ring at midcell, which mediates
septation (5, 6). Recently, however, prokaryotic proteins encoded
on large plasmids harbored in bacilli showing 15–20% sequence
similarity to both FtsZ and tubulin have been identified and
dubbed TubZ (7–12). Studies showed that the Bacillus thuringien-
sis TubZ protein from the pBtoxis plasmid is essential for plasmid
DNA segregation.
DNA segregation of most low copy number plasmids is carried
out by specific partition (par) systems. These systems require only
three elements: a centromere DNA site, a centromere-binding
protein, and a partition NTPase (13, 14). Partition systems have
been classified into two main types on the basis of the kind
of NTPase present (15). Type I systems contain NTPases with
deviant Walker A-type ATPase folds, whereas type II systems uti-
lize actin-like NTPases. Interestingly, both types of NTPases form
polymers in NTP-dependent manners that are implicated to play
a role in plasmid DNA separation (16–19). The recent discovery
of TubZ NTPases has led to the designation of “type III” par sys-
tems (13, 14). The best studied of these systems is that found on
the pBtoxis plasmid in B. thuringiensis. This plasmid stability sys-
tem is represented by an operon encoding two proteins: ORF156
(TubZ) and ORF157 (TubR) (7–9, 11). TubR is a 11.6 kDa DNA-
binding protein that shows no sequence homology to any known
protein. Studies showed that TubR binds a 48-bp centromere con-
taining four repeat sites in the pBtoxis plasmid and also autore-
gulates tubRZ transcription (8, 9). TubZ is a 54.4 kDa protein that
can assemble into filaments in a GTP-dependent manner (12).
Both proteins were found to be required for plasmid stability
(9). However, how the TubR and TubZ proteins work together
to effect pBtoxis plasmid segregation is not known. To gain insight
into the molecular mechanism utilized by these proteins in DNA
segregation, we carried out structural and biochemical studies on
the pBtoxis TubR and TubZ proteins. The TubR structure reveals
that it employs a helix-turn-helix (HTH) motif in a previously un-
described manner to bind DNA. TubZ contains a tubulin/FtsZ
fold but has structural distinctions from these proteins indicating
that it forms distinct protofilaments. TubR binds the flexible
C-terminal region of TubZ, thus attaching the TubZ filament
to the pBtoxis plasmid, providing a mechanism for plasmid move-
ment and, ultimately, segregation.
Results and Discussion
Overall Structure of pBtoxis TubR. The crystal structure of the 107-
residue pBtoxis TubR protein was solved to 2.0-Å resolution by
selenomethionine multiple wavelength anomalous diffraction
(MAD) methods (Table S1). The structure contains two TubR
molecules in the crystallographic asymmetric unit and consists
of residues 6–102 of one subunit and 4–100 of the second subunit,
and has Rwork∕Rfree ¼ 23.8%∕27.0%. The TubR structure forms a
highly
intertwined
dimer
with
dimensions
30 × 30 × 60 Å3
(Fig. 1A). Each TubR subunit has the topology β1-α1-α2-α3-α4-β2-
β3-α5, which is similar to winged HTH motifs found in a number
of DNA-binding proteins in both prokaryotes and eukaryotes
(20). In TubR, α3-α4 forms the HTH motif and the loop between
Author contributions: M.A.S. designed research; L.N., W.X., M.K., and M.A.S. performed
research; M.A.S. analyzed data; and M.A.S. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Freely available online through the PNAS open access option.
Data deposition: The atomic coordinates and structure factor amplitudes for the WT TubR
(C2), WT TubR (I222), TubR(S63W), TubZ, and TubZ-GTP-γ-S structures have been
deposited with the Protein Data Bank, www.pdb.org (PDB ID codes 3M8E, 3M9A,
3M8F, 3M8K, and 3M89).
1To whom correspondence may be addressed. E-mail: maschuma@mdanderson.org.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1003817107/-/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.1003817107
PNAS ∣June 29, 2010 ∣vol. 107 ∣no. 26 ∣11763–11768
BIOCHEMISTRY
β2 and β3, the wing. Indeed, each TubR subunit shows the stron-
gest structural similarity to members of the ArsR family of
prokaryotic winged helix transcription regulators, in particular
the Staphylococcus aureus CzrA protein (21, 22). Superposition
of one subunit of TubR onto that of CzrA results in a rmsd of
2.7 Å. This similarity includes the core regions of the winged
HTH, because the loops and N-terminal regions of the proteins
are structurally distinct. For example, TubR contains a β-strand in
its N-terminal region compared to a long helix in the CzrA struc-
ture (Fig. 1B). This structural similarity initially suggested that
TubR may be a member of the ArsR family of proteins. However,
the arrangement of the TubR dimer was found to be strikingly
different from the dimer organization exhibited by the ArsR
proteins (Fig. 1C).
ArsR family members are involved in metal-regulated tran-
scription processes whereby they act as repressors in their apo
forms and are induced off their DNA sites upon metal binding
(22). The specific dimer structures of the ArsR proteins are cri-
tical for creation of their metal-binding motifs. Not only does
TubR form a very different dimer from the ArsR proteins, it also
does not harbor any of their metal-binding signatures nor does it
contain any other characterized metal-binding motif. Consistent
with this, we find that the addition of metals has no effect on TubR
DNA binding. Dimerization of the ArsR proteins is imparted by
residues from the N-terminal regions, α1 and α5, which, impor-
tantly, leaves its recognition helices exposed for DNA interaction
(22). By contrast, the TubR dimer is formed primarily by contacts
between its twofold related “recognition helices” α4 and α4·. This
interaction results in a near complete burial of these helices, leav-
ing only the N-terminal residues exposed to solvent. Whereas the
α4 and α4· interaction creates the dimer core, the dimer is further
stabilized by interactions between the twofold related β1 strands,
which swap to form an antiparallel β-sheet. Residues from α1 and
α5 interact with β1 to further seal the top of the dimer. The dimer
interface formed by these interactions is predominantly hydropho-
bic and buries a large 1;200 Å2 of subunit surface from solvent.
TubR Forms a “Recognition Helix Dimer”: Implications for DNA Bind-
ing. Gel filtration studies on TubR confirmed that it is a dimer in
solution. However, the finding from the structure that the TubR
α4 recognition helices are buried in the dimer core has important
implications in terms of its DNA-binding mechanism. Indeed, it
suggests that, although TubR contains a structurally canonical
HTH, it is not utilized for DNA binding in a manner typical
of HTH proteins. A second crystal form (I222) of TubR, which
was solved to 2.5-Å resolution, revealed the same TubR dimer.
The presence of the identical dimer in two different crystal forms
and its large buried surface area supports that the dimer observed
in the crystal structures is physiologically relevant. However, to
test this, we mutated residues within the recognition helices that
the structure indicates are critical for dimerization and assayed
the ability of the mutant proteins to dimerize via gel filtration.
Specifically, we mutated Ser-63 and Ala-67 individually to tryp-
tophan and arginine.
The structure shows that residues occupying positions 63 and
67 must be small and largely hydrophobic to permit the proper
packing of the α4 helices in the dimer (Fig. S1A). Hence, the in-
troduction of the bulky side chain of tryptophan and, in particu-
lar, the large as well as charged side chain of arginine would be
predicted to be highly disruptive to dimerization. Gel filtration
analyses on purified mutant proteins clearly showed that the ar-
ginine mutants exist primarily as monomers in solution (>80%),
whereas the tryptophan mutants were able to maintain the di-
meric state (Fig. 2A and Fig. S1B). However, all mutant proteins
showed reduced or loss of DNA-binding activity as ascertained by
fluorescence polarization (FP) studies, which examined TubR
protein binding to its centromere site (Fig. 2B and Fig. S1 C
and D) (9). The fact that the monomeric mutants were severely
impaired in DNA binding was not surprising. However, the
finding that the tryptophan mutants, which were largely dimeric,
displayed reduced DNA-binding activity suggested that their oli-
gomer structures might be altered. To address this issue, the struc-
ture of S63W TubR was solved to 2.8-Å resolution, resulting in
Rwork∕Rfree values of 20.4%∕26.6% (Table S1). The subunit struc-
ture of S63W TubR is essentially identical to that of WT TubR as
revealed by their superimposition (rmsd of 1.2 Å) (Fig. 2 C and
D). However, this single subunit overlay shows that the S63W
TubR dimer, although the same as the WT in general arrange-
ment, is forced into a more open oligomer conformation in which
one subunit is rotated 20° away from its dimer mate compared to
WT. This rotation is required to accommodate the bulky S63W
side chains (Fig. 2D). The N-terminal β1–β1′-sheet interaction
appears to play a key role in holding the TubR subunits together.
In addition, the tight stacking of the twofold related Trp63 indole
groups (3.5 Å) provides a compensatory interaction that, com-
bined with the β1–β1′ interaction, apparently permits retention
of the dimer state, indicating why the TubR tryptophan mutants
were able to maintain the dimer state, albeit an altered dimer
state relative to WT (Fig. 2 C and D). By contrast, the TubR
arginine mutations, which introduced both bulk and charge with-
in the predominantly hydrophobic dimer interface, were highly
destabilizing for dimerization. In addition, the finding that the
S63W TubR mutant forms an altered dimer explains the severe
effect on DNA binding because a correct dimer orientation is
likely essential for binding to its palindromic DNA sites (9).
TubR-DNA Model. Because all but the N-terminal residues of the
TubR recognition helices are buried in the dimer interface, TubR
must use a different mode of DNA binding than the ArsR or other
HTH containing proteins (23). Examination of surface electro-
statics of TubR reveals that one face of the protein is electro-
negative, whereas the other is strongly electropositive (Fig. 3A).
Notably, the positive region is composed of one large and contig-
uous basic patch. Basic residues in this region correspond to Arg-
74, Arg-77, and Lys-79, in the wing and Lys-43, which is on α3, the
helix preceding α4 in the HTH motif. These residues were mutated
singly to alanine to examine their roles in DNA binding (Fig. 3B).
FP experiments showed that mutation of the basic wing residues
resulted in either a complete (R74A and R77A) or nearly com-
plete (K79A) abrogation of DNA binding, indicating that the
wings play a major role in TubR DNA binding. Residue Lys-43
is surface exposed and located at the center of the basic region
on the TubR dimer (Fig. 3A). The K43A mutant also showed
no binding to TubR, supporting the notion that the continuous
basic patch of TubR represents its DNA-binding surface.
To gain insight into the structural mechanism of DNA binding,
a DNA duplex was docked onto the basic patch of the TubR di-
Fig. 1.
B. thuringiensis pBtoxis TubR structure. (A) One TubR subunit is red
and the other cyan. Secondary structural elements and N and C termini are
labeled. (B) Superimposition of one subunit of TubR (Red) onto a S. aureus
CzrA subunit (Yellow). Regions with different structures are labeled. (C) Same
superimposition as B showing the location of the other subunit in the TubR
and CzrA dimers after one subunit is overlaid. A–C are in the same orienta-
tion to highlight differences. Figs. 1 A–C, 2 C and D, 3 A, C, and E, 4B, and 5B
were made by using PyMOL (31).
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mer by using the location of the mutations that affected DNA
binding as a guide (Fig. 3C). This model revealed that the wings
are positioned to interact with successive minor grooves, with
either the bases or the phosphate backbone depending on the
ability of the DNA to deform. In the model TubR interacts with
a minimum of 14 bp of DNA. However, the centromere bound by
Fig. 2.
The TubR “recognition helix” dimer. (A) Gel fil-
tration studies on TubR mutants S63W and S63R show-
ing that the S63W mutant remains dimeric whereas
S63R is >80% monomer. (B) Fluorescence polarization
studies examining the ability of WT TubR, S63R, and
S63W TubR mutants to bind iteronic DNA. Fluorescence
polarization units (millipolarization) and TubR concen-
tration (nM) are along the y and x axes, respectively.
The Kd of WT TubR for the centromere DNA is
8 2 nm. (C) Superimposition of WT TubR (Green) onto
the TubR S63W mutant structure (Tan). (D) Close-up of
the site of the S63W mutation in the expanded TubR
S63W dimer showing stacking interactions between
the twofold related tryptophans.
Fig. 3.
TubR-DNA binding. (A) Electrostatic surface potential of the TubR dimer. Blue and red represent electropositive and electronegative regions, respec-
tively. (Left) The electronegative side of the TubR dimer, and the side on the right is the electropositive side. Labeled on the left side are the locations of the
mutated residues. (B) FP binding isotherms showing the DNA binding of WT TubR and the K43A, R74A, R77A, and K79A mutants. Fluorescence polarization
units (millipolarization) and TubR concentration (nM) are along the y and x axes, respectively. (C) TubR-DNA model showing TubR electrostatic potential.
(D) Stoichiometry of TubR (subunit) binding showing titration curve of TubR into the 48-mer iteron resulting in a molar ratio of TubR subunit to DNA of
eight (or four) dimers. (E) Left: Ribbon diagram of the TubR-DNA model with the recognition helices colored yellow. Right: Ribbon diagram showing a canonical
HTH–DNA interaction (the λ repressor-DNA complex) with the recognition helices colored yellow (32).
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TubR consists of four 12-bp sites with the consensus T(T/A)(T/A)
(C/A)(G/A)GTTTA(A/C)(A/C) (9). Thus, we used FP to ascer-
tain the binding stoichiometry of TubR for its 48-bp centromere.
As shown in Fig. 3D, eight TubR subunits or four TubR dimers
bind the 48-mer centromere, consistent with a dimer of TubR
binding each palindrome. Thus, either TubR distorts its DNA
or the TubR dimers bind with some degree of overlap on their
DNA sites perhaps imparting cooperativity, as observed in other
centromere-binding protein–DNA interactions (13, 14). In addi-
tion to the insertion of the wings, a striking outcome of the mod-
eling was the finding that the N termini of the recognition helices,
which interact with each other in a parallel, coiled-coil-like man-
ner, are in position to insert into a single major groove. Structures
of HTH proteins bound to DNA have thus far shown that the
recognition helices insert singly into successive major grooves
by using residues in the first few turns or the central portion
of the recognition helix to contact the DNA bases (Fig. 3E). Thus,
in the TubR-DNA model, the HTH–DNA interaction is drama-
tically different from any displayed previously by a HTH protein.
TubZ Binds TubR-DNA. A characteristic feature exhibited by
partition centromere-binding proteins is the ability to bind their
partner NTPase (13, 14). To determine if TubR binds TubZ, we
utilized a FP assay and found that full length (FL) TubZ bound
avidly to the TubR-centromere complex (Fig. 4A). However,
unlike other partition systems in which the NTPase must be com-
plexed with nucleotide to bind its centromere-binding protein,
the interaction of TubR with TubZ did not require the presence
of GTP-Mg2þ. Previous studies have shown that the C-terminal
regions of tubulin and FtsZ mediate key binding events with their
target proteins (24–26). We noted that the terminal region of
TubZ, consisting of residues 407–484, is the most divergent region
between TubZ proteins and between TubZ and tubulin/FtsZ pro-
teins, suggesting that it may be similarly utilized and bind TubR.
To test this hypothesis, we constructed various TubZ truncations,
TubZ(1-407), TubZ(1-442), TubZ(1-460), and TubZ(1-470), and
examined the ability of each protein to bind TubR-DNA. TubZ
(1-407) showed no binding to TubR, whereas the remaining trun-
cation mutants bound weakly to TubR-DNA (Fig. 4A). Thus, the
data indicate that the last 14 amino acids of TubZ are critical for
the ability of TubZ to form a tight interaction with TubR but that
residues 408–470 also play an important role in this interaction.
These data demonstrate that TubR acts as a partition partner for
TubZ, linking it to pBtoxis plasmid DNA. Although TubZ has
been shown to form polymers in a GTP-dependent manner,
the TubZ protein displays limited sequence similarity to tubu-
lin/FtsZ, suggesting potential differences in TubZ and tubulin/
FtsZ structures (7–9). To gain insight into TubZ function, we next
determined structures of B. thuringiensis pBtoxis TubZ.
Structure of TubZ. Crystallization of FLTubZ was not successful, in
either its apo form or bound to guanine nucleotides. We noted that
FL TubZ degraded over time whereby C-terminal residues were
proteolyzed. Therefore, truncated TubZ proteins were utilized
in crystallization trials, and crystals were obtained of apo TubZ
(1-428) and the structure solved by MAD (Table S2). The model
consists of residues 1–79 and 91–404 and has an Rwork∕Rfree of
21.4∕24.9% to 2.3-Å resolution (Fig. 4B). No discernible oligomer-
ization of apo TubZ(1-428) was observed in the crystal packing,
and gel filtration analyses confirmed that it is monomeric
(Fig. S2). The overall TubZ structure can be divided into two main
domains: an N domain (residues 25–235) and a C domain (resi-
dues 258–377). These domains are connected by a long, core helix,
H7. The TubZ N domain has a Rossman fold and consists of six
parallel β-strands with topology 3-2-1-4-5-6. The resulting β-sheet
is sandwiched by five α-helices, with two helices on one side and
three on the other. The C domain consists of four β-strands with
the topology 1-4-2-3. The C-domain β-strands are arranged nearly
perpendicular to those in the N domain. In addition to these main
protein domains, there are two helices: one at the N terminus, H0,
and a long helix at the C terminus, H11 (Fig. 4B). Database
searches showed that TubZ indeed belongs to the tubulin/FtsZ fa-
mily of proteins and is similar to both eukaryotic and prokaryotic
members of the family; TubZ can be optimally superimposed with
rmsds of 3.4 Å onto both bovine α tubulin and Pseudomonas
aeruginosa FtsZ (1, 5). Whereas the two-domain architectures
of tubulin, FtsZ, and TubZ are similar in overall structure, the
extreme N- and C-terminal regions of these proteins are very di-
vergent (Fig. S3 A–D).
N-Terminal and C-Terminal Differences in TubZ, FtsZ, and Tubulin: Im-
plications for Polymer Formation and Target Protein Binding. Tubulin
proteins do not contain significant N-terminal extensions,
whereas FtsZ proteins from different organisms show structural
variability within their N-terminal regions. For instance, in the
Escherichia coli FtsZ structure the N-terminal residues are disor-
dered, whereas Methanococcus jannaschii FtsZ has an extra
N-terminal helix, H0, which is flexibly attached to the body of
the protein and has been captured in multiple orientations (5).
Although H0 is not conserved in FtsZ proteins, one M. jannaschii
FtsZ structure revealed a semicontinuous polymer in the crystal,
thought to closely represent in vivo protofilaments, which utilizes
H0 in subunit-subunit contacts (4). This finding suggests that
the flexibly attached H0 is stabilized in a specific orientation
by protofilament formation, at least in the M. jannaschii protein.
The TubZ H0 helix extends in the opposite direction compared to
that of the protofilament stabilized FtsZ H0 helix. Moreover, in
TubZ, H0 is not flexibly attached to the N domain but is tightly
anchored to the C domain through numerous interactions with
the core helix and C-domain residues. The large number of inter-
actions involving H0, and the fact that it covers what would other-
wise be a surface exposed hydrophobic patch, indicate that the
TubZ H0 does not undergo conformational changes during
protofilament formation and is important for the general fold
of TubZ (Fig. S3 A and C).
Data suggest that FtsZ and tubulin form protofilaments with
similar longitudinal contacts (4). However, the TubZ structure
reveals key differences, primarily in its C-domain and C-terminal
regions, which suggest that it forms protofilaments distinct from
those formed by tubulin/FtsZ. A notable difference is the struc-
ture of loop 7 (L7). This loop inserts into the adjacent subunit
providing the key catalytic residues required for GTP hydrolysis.
In tubulin/FtsZ proteins, L7 has the consensus GXXNXDXAD.
In TubZ, the loop is very divergent in conformation compared
Fig. 4.
TubZ interacts with TubR-DNA and contains a tubulin/FtsZ fold. (A) FP
assay measuring binding of FL TubZ, TubZ(1-470), TubZ(1-460), TubZ(1-442),
and TubZ(1-407) to TubR-DNA. Below is the control (TubZ titrated into DNA
alone). Millipolarization units and TubZ concentration (nM) are along the y
and x axes, respectively. (B) TubZ(1-428) structure. The N domain or GTP-bind-
ing domain is colored salmon and the C domain purple. The interdomain
helix, H7, is red. TubZ also contains an N-terminal helix, H0 (Yellow), and
a C-terminal helix, H11 (White).
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to that in FtsZ/tubulin and consists of the sequence 256-
DNVTYDPSD-266. In addition to the N-terminal region, the
extreme C-terminal extensions of tubulin, FtsZ, and TubZ are
structurally divergent (Fig. S3B). In FtsZ, the C-terminal region
forms a small, two-stranded β-sheet and continues into an ex-
tended region that is involved in binding adaptor proteins such
as FtsA and ZipA (6, 25). By contrast, the C-terminal regions
of tubulin proteins consist of a two-helix bundle followed by an
extended region. Like FtsZ, however, these regions interact with
numerous target proteins such as the microtubule-associated pro-
teins (MAPs) (24). Consistent with this, the C-terminal regions of
tubulin have been shown to face the outside of the microtubule. A
characteristic feature of the extreme C-terminal extensions of
tubulin proteins is their highly acidic nature (3). This acidic region
has been shown to be critical for binding to several MAPs that
harbor a substantial basic character, such as tau, MAP2, and
MAP4 (24, 27).
The TubZ C-terminal region is also helical, but it contains a
single, long helix. Notably, the TubZ-tubulin overlay shows that
the long C-terminal helix of TubZ would dramatically clash with
the adjacent subunit in a polymer, providing support for the no-
tion that TubZ forms protofilaments different from tubulin/FtsZ
(Figs. S3B and S4). Interestingly, and in contrast to tubulin pro-
teins, the flexible C-terminal region of TubZ that follows H11 is
highly basic, in particular the last 14 residues. We have shown that
these residues play a central role in TubR binding (Fig. 4A). TubR
uses its electropositive face for DNA binding, leaving exposed its
opposite face for TubZ interaction. Notably, this exposed face is
strongly electronegative and hence would complement the basic
C-terminal tail of TubZ (Fig. 3A).
Tubulin/FtsZ protofilaments combine to form higher-order
structures. In tubulin, the protofilaments interact in a parallel
manner to form microtubules. Central to microtubule formation
are lateral contacts between protofilaments from the so-called M
loop, between H10 and S9. In tubulin, this loop is composed of 13
residues (1–3). The corresponding loop is much shorter in FtsZ
proteins, consistent with the fact that FtsZ does not form tubulin
microtubule-like structures (5, 28–30). In TubZ, the M loop is
even shorter than in FtsZ, spanning only four residues. In fact,
the TubZ/tubulin overlay shows that the side of the molecule con-
taining the M loop is the most divergent between these proteins.
These combined findings suggest that TubZ not only forms pro-
tofilaments with distinct longitudinal contacts compared to FtsZ
and tubulin, but it also does not form tubulin-like microtubule
structures.
TubZ Interactions with Guanine Nucleotides. Consistent with TubZ
being a member of the tubulin/FtsZ family, our isothermal titra-
tion calorimetry (ITC) studies showed that TubZ binds guanine
nucleotides with high affinity; Kds for GTP-γ-S and GDP were
∼0.69 and 26 μM, respectively (Fig. 5A). We next determined
the structure of the TubZ-GTP-γ-S complex by soaking GTP-γ-S
into preformed TubZ(1-428) crystals. The TubZ-GTP-γ-S struc-
ture contains TubZ residues 1–79 and 91–404, one GTP-γ-S,
and has Rwork∕Rfree ¼ 21.8%∕25.5% (Fig. 5B and Table S2).
The structure shows that TubZ binds GTP-γ-S in the same GTP
binding pocket as tubulin/FtsZ (1–5). Comparison of the apo
and GTP-γ-S bound TubZ structures indicated that, like FtsZ,
guanine nucleotide binding does not lead to significant conforma-
tional changes (5). The phosphate binding pocket is formed by
two of the most highly conserved regions between TubZ and
tubulin/FtsZ called loops 1 and 4 (L1 and L4) (1–3). L1 contacts
the GTP-γ-S α- and β-phosphate groups via the Gln-32 and Lys-33
amide nitrogens. The L1 region of FtsZ and tubulin contain the
sequences GQ(A/G)G and GQCG, respectively, whereas in TubZ
the motif is 31-GQKG-34. However, the alanine/glycine and
cysteine residues in FtsZ and tubulin do not contact the bound
nucleotide; the TubZ Lys-33 side chain makes stacking interac-
tions with the guanine base (Fig. 5B). L4 represents the so-called
signature motif [GGGTG(T/S)G], which serves as an identifier of
tubulin/FtsZ family members. Like FtsZ and tubulin, the L4
region of TubZ-GTP-γ-S makes phosphate interactions via its
glycine amide nitrogens. Whereas L1 and L4 residues of the N
domain mediate phosphate contacts, the GTP-γ-S guanine moiety
is specified from residues in the core helix, H5, and C domain. In
this regard, an important motif is loop 6 (L6). In FtsZ and tubulin,
L6 has the consensus (F/Y)XXX(N/D) and the conserved (F/Y)
residue functions in guanine base stacking. This region in TubZ,
236-WKXXXN-241, is in an altered conformation compared to
FtsZ and tubulin structures. Despite the presence of the trypto-
phan, which might be expected to interact with the guanine,
the side chain of Lys-237 instead stacks with the guanine ring.
Hence, in the TubZ-GTP-γ-S structure, the guanine base does
not interact with aromatic residues as in tubulin/FtsZ but is sand-
wiched between the aliphatic portions of two lysine side chains,
Lys-33 and Lys-237. Finally, two asparagine residues, Asn-213
and Asn-241, from L6 effectively read the guanine N2/N3 and
N1/O6 atoms, respectively, providing high specificity in TubZ’s in-
teraction with guanine nucleotides.
pBtoxis DNA Segregation: TubR Plasmid Partition Model. Our data
show that TubR binds to the flexible, C-terminal, basic region
of TubZ. The flexibility and location of the TubZ C-terminal
extension suggest that it is not required for polymerization and
thus may be exposed on the surface of TubZ filaments. Indeed,
negative stain EM images show that TubZ(1-407) forms polymers
in a GTP-dependent manner similar to the FL protein (Fig. S5).
Recent data suggesting that TubZ filaments are stabilized by
Fig. 5.
TubZ-guanine nucleotide interactions. (A) ITC binding isotherms
showing TubZ-GDP (Left) and TubZ-GTP-γ-S interaction (Right). (B) Left:
Overall structure of the TubZ-GTP-γ-S complex. β-strands are colored magen-
ta and helices cyan, and the GTP-γ-S molecule is shown as cpk. Right: Close-up
view of the GTP binding pocket with the initial Fo-Fc electron density map
(Blue Mesh), contoured at 4.5σ, and calculated before the GTP-γ-S was
included in refinement.
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the presence of a GTP cap and undergo treadmilling are consis-
tent with the notion that TubZ displays tubulin-like polymer dy-
namics (12). Thus, on the basis of the combined data, we suggest
a model for TubR/TubZ mediated pBtoxis plasmid segregation
shown in Fig. 6. In this model, multiple TubR dimers first bind
to the iteronic DNA on the pBtoxis plasmid leading to the crea-
tion of a high local concentration of TubR, which can recruit a
TubZ polymer, likely by interactions between the acidic TubR
dimer face and the basic C-terminal TubZ region. Importantly,
this interaction serves to attach the pBtoxis plasmid to the TubZ
polymer, which undergoes treadmilling, adding subunits at the þ
end and losing subunits at the −end. The bound TubR-pBtoxis
can be handed off from the −end to the molecules in the growing
þ end, leading to the transport of the pBtoxis plasmid to the cell
pole. Interestingly, it has been shown that once TubZ polymers
reach and interact with the cell pole, they bend around the curved
pole and continue growing in the other direction (7). The force of
the interaction with the membrane likely causes the release of
TubR-pBtoxis, the net result being transport of pBtoxis to the cell
pole. Of course, this model is simplified and many questions re-
main. For example, how directionality is achieved and how the
replicated plasmids are driven to opposite cell poles is not clear.
However, given the large size of the pBtoxis plasmid (8), it may be
that only one TubR-pBtoxis “tram” can be bound at once by the
rapidly treadmilling TubZ polymer and that, once one such a tram
is unloaded after reaching the cell pole, another engages when the
now reversed polymer treadmills toward the opposite cell pole.
Materials and Methods Summary
Detailed methods are provided in SI Materials and Methods.
Briefly, the tubR and tubZ genes were codon optimized (for
E. coli expression), subcloned into pET15b, expressed, and pur-
ified. WT TubR crystals were grown with NaCl and phosphate.
TubR S63W was crystallized with PEG and ethylene glycol and
TubZ with sodium formate. Detailed assay conditions for FP,
ITC, electron microscopy, and gel filtration are provided in
SI Materials and Methods.
ACKNOWLEDGMENTS. This work was supported by the Burroughs Wellcome
Career Development Award 992863 and National Institutes of Health Grant
GM074815 (to M.A.S.).
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11768
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Ni et al.
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3M8L
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Crystal Structure Analysis of the Feline Calicivirus Capsid Protein
|
JOURNAL OF VIROLOGY, June 2010, p. 5550–5564
Vol. 84, No. 11
0022-538X/10/$12.00
doi:10.1128/JVI.02371-09
Copyright © 2010, American Society for Microbiology. All Rights Reserved.
Conformational Changes in the Capsid of a Calicivirus upon
Interaction with Its Functional Receptor
Robert J. Ossiboff,1† Yi Zhou,2† Patrick J. Lightfoot,1 B. V. Venkataram Prasad,2 and John S. L. Parker1*
Baker Institute for Animal Health, College of Veterinary Medicine, Cornell University, Ithaca, New York 14853,1 and Verna and
Marrs McClean Department of Biochemistry and Molecular Biology, Baylor College of Medicine, Houston, Texas 77030 2
Received 10 November 2009/Accepted 19 March 2010
Nonenveloped viral capsids are metastable structures that undergo conformational changes during virus
entry that lead to interactions of the capsid or capsid fragments with the cell membrane. For members of the
Caliciviridae, neither the nature of these structural changes in the capsid nor the factor(s) responsible for
inducing these changes is known. Feline junctional adhesion molecule A (fJAM-A) mediates the attachment
and infectious viral entry of feline calicivirus (FCV). Here, we show that the infectivity of some FCV isolates
is neutralized following incubation with the soluble receptor at 37°C. We used this property to select mutants
resistant to preincubation with the soluble receptor. We isolated and sequenced 24 soluble receptor-resistant
(srr) mutants and characterized the growth properties and receptor-binding activities of eight mutants. The
location of the mutations within the capsid structure of FCV was mapped using a new 3.6-Å structure of native
FCV. The srr mutations mapped to the surface of the P2 domain were buried at the protruding domain dimer
interface or were present in inaccessible regions of the capsid protein. Coupled with data showing that both the
parental FCV and the srr mutants underwent increases in hydrophobicity upon incubation with the soluble
receptor at 37°C, these findings indicate that FCV likely undergoes conformational change upon interaction
with its receptor. Changes in FCV capsid conformation following its interaction with fJAM-A may be important
for subsequent interactions of the capsid with cellular membranes, membrane penetration, and genome
delivery.
The interactions between viruses and receptors on the sur-
face of host cells strongly influence viral pathogenesis and
regulate morbidity and mortality in the host. Virus-receptor
interactions determine the types of cells that can be infected,
the pathway of entry into the cell, and the efficiency of pro-
ductive infection. Interactions between nonenveloped virus
capsids and their receptor(s) trigger one or more steps re-
quired for infectious entry. These steps can include interaction
with other receptors, exposure to low pH or endosomal pro-
teases, or other factors. Ultimately, one or more of these in-
teractions induce changes in capsid conformation that result in
the exposure of hydrophobic regions or release of a lipid-
seeking factor that can interact with and disrupt the limiting
cellular membrane to allow the capsid and/or the genome to be
delivered to the interior of the cell (reviewed in reference 60).
The Caliciviridae are small nonenveloped viruses containing
a positive-sense RNA genome (7 to 8 kb). Several important
disease-causing members of the Caliciviridae, including human
noroviruses and rabbit hemorrhagic disease virus, cannot be
propagated in tissue culture systems (19, 56). This has slowed
progress on studies of the mechanisms of cellular entry of these
viruses. In contrast, feline caliciviruses (FCVs) propagate
readily in tissue culture, and two cell surface receptor mole-
cules, feline junctional adhesion molecule A (fJAM-A) and
2,6 sialic acid, are known (29, 55).
The FCV receptor, fJAM-A, is a type I transmembrane
glycoprotein (molecular size of 36 to 41 kDa) member of the
immunoglobulin superfamily (IgSF); it consists of an amino-
terminal signal peptide, an extracellular domain (composed of
two Ig-like domains—a membrane-distal D1 and a membrane-
proximal D2), a transmembrane domain, and a short cytoplas-
mic domain that contains a type II PDZ domain-binding motif
(11, 30). We have previously shown that the D1 domain of the
fJAM-A ectodomain is necessary and sufficient for FCV bind-
ing and that preincubation of FCV with soluble fJAM-A
(sfJAM-A) results in virus neutralization (35). Additional roles
that fJAM-A might play in FCV entry, however, have not been
investigated.
Caliciviruses are composed of 180 copies of a single capsid
protein. Atomic resolution structures of recombinant virus-like
particles of Norwalk virus (genus Norovirus) and native San
Miguel sea lion virus (SMSV) virions (genus Vesivirus) indicate
that the virion consists of 90 dimers of the capsid protein
arranged in T3 icosahedral symmetry (5, 41). Each capsid
monomer contains three structural domains—an N-terminal
arm (NTA), the shell (S), and a protruding domain (P) that is
further subdivided into P1 and P2 subdomains. The distal sub-
domain, P2, is structurally conserved between Norwalk virus
and SMSV, but there is little sequence conservation. In the
primary sequence of the FCV capsid, there are two hypervari-
able regions that contain neutralizing epitopes (18, 34, 58). The
corresponding hypervariable regions (HVRs) of the SMSV
capsid structure map to surface-exposed loops. Surface resi-
dues at the dimeric interface between two capsid monomers
* Corresponding author. Mailing address: Baker Institute for Ani-
mal Health, College of Veterinary Medicine, Cornell University, Hun-
gerford Hill Road, Ithaca, NY 14853. Phone: (607) 256-5626. Fax:
(607) 256-5608. E-mail: jsp7@cornell.edu.
† These authors contributed equally to this publication.
Published ahead of print on 31 March 2010.
5550
are conserved within individual calicivirus genera, and it has
been suggested that this interface is involved in receptor bind-
ing (5). A cryo-electron microscopy (cryo-EM) reconstruction
of the FCV vaccine strain F9 complexed with the ectodomain
of fJAM-A (modeled on the crystal structures of SMSV and
human JAM-A, respectively) shows that fJAM-A engages the
top of the P2 domain and that binding causes a rotation in
the P dimer (1). However, the relatively low resolution and the
lack of atomic resolution structures of FCV and fJAM-A pre-
vented precise identification of residues on the viral capsid that
contact fJAM-A.
A classical approach for identifying virus residues that di-
rectly bind or modulate the binding of a receptor is to select for
mutant viruses resistant to neutralization with soluble recep-
tors (6, 23, 46). Soluble receptor-resistant (srr) mutants of
poliovirus revealed that both surface-exposed and internal res-
idues
regulate
receptor
attachment
and
conformational
changes in the capsid (6, 42). Here, we report 24 srr mutants
and the location of their capsid mutations on a 3.6-Å structure
of FCV. In addition, we describe the growth kinetics and re-
ceptor-binding properties of a subpanel of eight srr mutants
and examine changes in capsid hydrophobicity concurrent with
the interaction of FCV capsids with sfJAM-A.
MATERIALS AND METHODS
Cells and viruses. Crandell-Reese feline kidney (CRFK; ATCC CCL-94) cells
were grown in Eagle’s minimal essential medium (EMEM; CellGro) supple-
mented with 5% fetal bovine serum (FBS; HyClone), 100 U ml1 penicillin, 100
g ml1 streptomycin, 0.25 g ml1 amphotericin B, 1 mM sodium pyruvate,
and nonessential amino acids (CellGro). Suspension Chinese hamster ovary
(CHO-S) cells (Invitrogen) were grown in CHO serum-free medium (Gibco).
The F9 vaccine strain (VR-782) of FCV was obtained from the ATCC. The
viral isolates FCV-5, Deuce, Kaos, FCV-127, FCV-131, and FCV-796 were
previously characterized (36). FCV-Urbana viral stock was generated from a
full-length infectious clone as previously described (50, 51). Briefly, CRFK cells
were infected with vaccinia virus expressing T7 RNA polymerase (MVA/T7) and
then transfected using FuGene 6 (Roche) 1 h later with the FCV-Urbana infec-
tious clone (pQ14). Samples were lysed at 18 h postinfection (h pi) by freeze-
thawing, and viral plaques were selected. The viral capsid gene was sequenced to
verify that FCV-Urbana was present. Third-passage viral stocks were prepared
from twice-plaque-purified viruses amplified in CRFK cells. Plaque assays were
performed as previously described (36).
Antibodies and reagents. A rabbit antiserum against the purified fJAM-A
ectodomain was previously described (35). Virus was detected with either an
anti-FCV mouse monoclonal antibody (MAb) (S1-9; Custom Monoclonal Anti-
bodies International) or rabbit anti-FCV-5 antiserum. The fJAM-A extracellular
domain (amino acid residues 26 to 232) was purified as a glutathione S-trans-
ferase (GST)-tagged fusion as previously described (35). Recombinant protein
was purified by glutathione affinity chromatography. When GST-free fJAM-A
was desired, N-terminal GST and His tags were cleaved from the recombinant
protein by using a human rhinovirus (HRV) 3C protease (Novagen) according to
the manufacturer’s directions.
Neutralization of FCV by fJAM-A and selection of soluble receptor-resistant
(srr) mutants of FCV-5. To investigate the neutralization of FCV by fJAM-A,
1 105 PFU of virus in Dulbecco’s modified Eagle’s medium (DMEM;
CellGro) plus 0.1% bovine serum albumin (BSA; Calbiochem) were incubated in
either the presence or absence of sfJAM-A (concentrations of receptor, temper-
atures of incubation, and lengths of incubation varied by experiment; please refer
to Results for experimental details). The infectious titer of the samples was
determined by plaque assay. srr mutants of FCV-5 were selected by plaque
titrating virus neutralized as described above on CRFK cells in six-well plates and
overlaid with EMEM containing 5% FBS and 1% Bacto agar. Following incu-
bation at 37°C in humidified 5% CO2 for 48 h, individual plaques were selected
and amplified without selection on CRFK cells in 12-well plates (Corning). Once
wells showed 100% cytopathic effect (CPE), a small aliquot from each well was
again incubated with GST–fJAM-A and used to infect CRFK cells in 12-well
plates. In wells that developed CPE, a sample of the lysate was subjected to an
additional round of GST–fJAM-A neutralization, and samples were amplified on
CRFK cells in a 96-well plate (Corning). Twenty-four wells showing CPE were
chosen at random and twice plaque-purified. To verify that the selected srr
mutants were resistant to neutralization by soluble receptor, a subpanel of eight
srr mutants was incubated with a soluble receptor, and the infectious titer of the
samples was determined by plaque assay.
Sequencing the major (VP1) and minor (VP2) capsid proteins of the srr
mutants. Recovery of viral RNA, reverse transcription-PCR (RT-PCR), and
sequencing of the major capsid protein of the srr mutants were performed as
described previously (36). Briefly, total RNA was extracted from infected cell
lysates (RNeasy minikit; Qiagen), and first-strand cDNA synthesis of the capsid
region of the genome was performed using a degenerate primer and AccuScript
high-fidelity reverse transcriptase (Stratagene). The major capsid open reading
frame (ORF) was then amplified from the first-strand cDNA template (primers
available upon request). The resulting 2.1-kb PCR products, encompassing the
entirety of the capsid ORF, were purified and sequenced directly. First-strand
cDNA synthesis of the region of the genome containing VP2 was performed
using RNA extracted from infected cell lysates as described above and using the
3 RACE system for amplification of cDNA ends (Invitrogen). The minor capsid
ORF was amplified from the first-strand cDNA template, and the resulting PCR
products were purified and sequenced directly. The VP2 sequence of FCV-5 was
submitted to GenBank (accession no. HM001263).
Purification of FCV. Roller bottles (Corning) containing confluent CRFK
monolayers were infected with FCV (multiplicity of infection [MOI] 5) and
incubated for 8 h at 37°C. At 8 h pi, cells were removed from the plastic surface
by physical agitation, and the cells were collected by centrifugation of the me-
dium at 500 g. The cell pellet was resuspended in 250 mM NaCl and 85 mM
Tris base (pH 7.5) and frozen at 80°C. The pellet was thawed and briefly
sonicated to lyse the cells, and trichlorotrifluoroethane was used to extract the
virus from the cell lysate. To clarify the phases, the trichlorotrifluoroethane-
lysate mixture was centrifuged for 10 min at 5,850 g. The aqueous phase was
removed and the trichlorotrifluoroethane extraction step repeated. Following the
second centrifugation, the aqueous phase was placed on a CsCl gradient (1.30 to
1.45 g/ml) in an ultracentrifuge tube (Beckman) and centrifuged at 97,000 g for
16 h. Typically, two bands were visible—a lower band containing infectious
particles and an upper band containing noninfectious particles. Occasionally, a
third minor band would be visible between the upper and lower bands; infectivity
of this band when present was variable. Each band was collected separately and
dialyzed exhaustively against virion buffer (150 mM NaCl, 10 mM Tris base, 15
mM MgCl2 [pH 7.2]) for 48 to 72 h at 4°C. Purified virus was stored at 4°C.
Crystallization. “Full” infectious particles of FCV-5 were used for crystalliza-
tion. Crystallization trials were carried out using the hanging drop vapor diffusion
method at room temperature. Crystal screens 1 and 2 from Hampton Research
were used to explore various conditions. Typically, for each condition, 2 l of the
virus solution, at a concentration of 3 mg/ml, was mixed with 2 l of the well
solution and equilibrated with 0.5 ml of the well solution. Initial screening
produced crystals under a few conditions, including the crystal screen 1 condition
3 (0.4 M ammonium phosphate) and condition 13 (30% polyethylene glycol
[PEG] 400, 0.1 M Tris HCl [pH 8.5], 0.2 M sodium citrate). After optimization
of crystallization conditions and synchrotron on-site screening, crystals of the
best diffraction quality were obtained using a well solution containing 0.45 M
ammonium phosphate with additive screen reagent C10 (1.0 M glycine). Cryo-
protection of crystals was carried out by soaking the crystals in the mother liquor
with increasing concentrations of PEG 400 through six steps, as follows: 5%,
10%, 15%, 20%, 22.5%, and 25%. The equilibration time at each solution was at
least 10 min. The crystals were then flash frozen in liquid nitrogen and shipped
to synchrotron sites for data collection.
Data collection and analysis. The diffraction data from single crystals were
collected under cryo-EM conditions at an APS 19ID station with monochromatic
X-rays (wavelength 0.9795 Å) and a detector to crystal distance of 480 mm on
an ADSC 3-by-3 charge-coupled-device (CCD) detector, using an oscillation
angle of 0.3° and an exposure time of 3 s. Indexing, integration, scaling, postre-
finement, and reduction of the data were carried out using the HKL-2000 (37)
and d*TREK (40) packages. Analysis of the diffraction data clearly indicated that
the FCV-5 crystal belonged to the orthorhombic space group I222 (Table 1). The
higher-resolution reflections were generally weak and appeared to be sensitive to
radiation damage. A similar trend was observed with several other crystals that
were used for data collection. The best crystal, which diffracted to 3.6 Å, was
used for data collection. The data between 30 and 3.6 Å from 300 frames were
scaled with an Rmerge factor of about 22%. The overall completeness of the data
to 3.6-Å resolution is about 90.1%. To determine the precise orientation of the
virus particle in the unit cell, self-rotation functions (45) were calculated using
the program GLRF (59). These calculations showed all the peaks corresponding
VOL. 84, 2010
CALICIVIRUS CONFORMATIONAL CHANGE
5551
to 5-fold ( 72°), 3-fold ( 120°), and 2-fold ( 180°) axes, which are
expected from a particle with icosahedral symmetry. In the I222 space group,
consistent with the unit cell dimensions and the particle radius, which is esti-
mated to be 200 Å, the particle position is uniquely defined by the intersection of
the crystallographic 2-fold axes. In such a setting, an icosahedral particle can assume
one of the two possible orientations. This ambiguity in orientation was clearly
resolved by the self-rotation function calculations by use of the X-ray diffraction
data. Each crystallographic asymmetric unit is composed of one-fourth of the virion
with a 15-fold noncrystallographic redundancy due to one set of icosahedral 5- and
3-fold symmetry.
Crystal structure determination and refinement. A properly positioned and
oriented SMSV-4 capsid was placed in an FCV-5 unit cell. One initial model at a
resolution of 10 Å was calculated from the 15 copies of SMSV-4 (Protein Data Bank
[PDB] ID 2GH8) capsid protein related first by icosahedral 5-fold symmetry and
then by icosahedral 3-fold symmetry. Phase refinement and extension were carried
out by iterative cycles of real-space electron density averaging, solvent flattening, and
back transformation with the RAVE and CCP4 (3, 25) program packages. The
phases were extended to the final 3.6-Å resolution with each step being less than one
reciprocal space point. An initial mask was constructed from the cryo-EM recon-
struction with the program MAMA, and the mask was edited at each step to avoid
truncation of the density. The averaged 3.6-Å density map is of good quality and
readily interpretable (see Fig. 2A). After initial model building using COOT (12)
and the final refinement using CNS (3), the structure has an Rfree (2) of 0.39,
calculated using 10% of the data that were not included in the refinement, and a final
R factor of 0.37. The atomic coordinates and associated structure factors have been
deposited into the PDB (www.pdb.org) (PDB ID 3M8L).
Virus-binding assay by flow cytometry. FCV binding to CHO-S cells transiently
expressing fJAM-A was detected as previously described (35). Briefly, CHO-S cells
(106 cells) were transiently transfected with a plasmid encoding fJAM-A. At 24 h
posttransfection, the cells were washed and incubated with FCV (MOI 5) in
phosphate-buffered saline (PBS) (137 mM NaCl, 3 mM KCl, 8 mM Na2HPO4 [pH
7.5]) plus 1% BSA (PBSA) on ice for 30 min. After being washed to remove
unbound virions, the cells were incubated with fJAM-A rabbit antiserum and anti-
FCV mouse MAb in PBSA on ice for 30 min. The cells were then washed in PBSA,
fixed in 2% paraformaldehyde in PBS, and incubated with Alexa 647-conjugated
goat anti-rabbit and Alexa 488-conjugated goat anti-mouse IgG for 30 min at room
temperature. The cells were analyzed using a FACSCalibur (Becton Dickinson).
Cells expressing fJAM-A on their surface were gated and analyzed for virus binding
(10,000 cells were gated for each sample).
Plaque reduction of srr mutants by fJAM-A antiserum. Confluent CRFK
monolayers in 60-mm dishes (Corning) were incubated in PBSA with or without
fJAM-A rabbit antiserum for 30 min at room temperature. Virus (100 PFU per
dish) was adsorbed for 1 h at room temperature. The monolayers were then
overlaid with EMEM containing 5% FBS and 1% Bacto agar. After incubation
at 37°C for 48 h in humidified 5% CO2, the overlay was removed, and the cells
were fixed with 10% buffered formalin and stained with 1% (wt/vol) crystal violet
in 20% ethanol. The plaques were counted, and percent reduction from controls
was calculated.
Single- and multiple-cycle growth kinetics of srr mutants. CRFK cell mono-
layers were inoculated with srr mutants at MOIs of 5 and 0.01 for single- and
multiple-cycle growth curves, respectively. Virus was adsorbed to cells for 1 h at
room temperature in DMEM plus 0.1% BSA; EMEM plus 5% FBS supple-
mented as described above was then added, and the cells were incubated at 37°C
in 5% CO2. At various times postinfection, samples were collected and stored at
80°C for later titration. Prior to plaque assay, all samples were frozen and
thawed three times. Virus yield at each time point was calculated by subtracting
the log10(PFU ml1) at T 0 from the log10(PFU ml1) measured at the time
point.
ELISA binding of FCV to fJAM-A. Purified fJAM-A ectodomain (1 g ml1)
in carbonate/bicarbonate buffer (Sigma) was bound to 96-well enzyme-linked
immunosorbent assay (ELISA) plates (100 ng per well) at 4°C overnight. The
wells were blocked with PBS containing 0.05% Tween 20 (PBS-T) plus 0.5%
BSA for 1 h at room temperature and washed three times with PBS-T. Twofold
dilutions of purified virus (starting concentration, 200 g ml1) were prepared in
PBS-T plus BSA; 100 l of each dilution of the virus was bound to plates for 3 h
on ice. After the plates were washed, bound virus was detected with rabbit
anti-FCV-5, followed by horseradish peroxidase (HRP)-conjugated goat anti-
rabbit IgG. Antibodies diluted in PBS-T plus BSA were incubated with the plates
for 90 min on ice. Following washing, substrate 2,2-azino-bis(3-ethylbenzthia-
zoline-6-sulfonic acid) (Sigma) in a citric acid-sodium phosphate buffer was
added to each well and incubated for 60 min at room temperature. Absorbance
was measured at 595 nm on a Microplate biokinetics reader (BioTek Instru-
ments). The mean standard error (SE) of three plates (two replicates per
plate) is shown.
ELISA competitive binding of FCV in the presence of sfJAM-A. Purified
fJAM-A ectodomain (1 g ml1) in carbonate/bicarbonate buffer was bound to
96-well ELISA plates (100 ng per well) at 4°C overnight. The wells were blocked
with PBS-T plus BSA for 1 h at room temperature and washed three times with
PBS-T. Twofold dilutions of fJAM-A ectodomain were prepared in PBS-T plus
BSA with 750 ng purified virus (starting fJAM-A concentration, 5.3 M); 100 l
of each dilution of the fJAM-A–virus mixture was bound to plates for 3 h on ice.
Plates were washed, and virus binding was detected as described above for the
assay to determine virus binding to fJAM in the absence of soluble receptor.
Absorbance readings for each virus were normalized, and the mean SE of
three plates (two replicates per plate) is shown.
Detection of bis-ANS binding to FCV by fluorescence spectroscopy. Purified
virus (60 g ml1, a 0.5 M concentration of the FCV VP1 dimer) or 1.5 M
purified receptor (fJAM-A or hJAM-A ectodomain) diluted in virion buffer was
incubated in a 400-l fluorometer cell (Varian) at 4° or 37°C for 10 min to allow
temperature equilibration. To detect bis-(8-anilinonaphthalene-1-sulfonate)
(bis-ANS) binding to native virus and receptor, bis-ANS was added to the sample
to a final concentration of 3 M. Bis-ANS was excited at 395 nm (slit width, 5
nm) and emission collected at 495 nm (slit width, 10 nm) at 2-s intervals for 10
min on a Cary Eclipse spectrofluorometer (Varian). To investigate the binding of
bis-ANS to preincubated virus and receptor, virus and receptor were mixed and
incubated for 10 min at 4°or 37°C. Bis-ANS was added (3 M) and sample
readings were performed as detailed above. All spectrofluorometer readings
were taken with the photomultiplier tube set at 700 V.
Statistical analyses. The Analyze-it (Analyze-it Software) statistical analysis
add-in for Microsoft Excel was used to perform analysis of variance (ANOVA)
where necessary. Graphs were prepared using Kaleidagraph (Synergy Software)
and Adobe Illustrator CS3 (Adobe). Protein images were prepared using
PyMOL (DeLano Scientific) (8).
RESULTS
FCV-5 is neutralized by preincubation with the sfJAM-A
ectodomain at 37°C. Preincubation of nonenveloped viruses,
such as rhinoviruses and polioviruses, with their functional
receptors neutralizes virus infectivity (17, 23). In a previous
study, we showed that preincubation on ice of FCV-5 with a
purified soluble GST-fused form of the ectodomain of fJAM-A
modestly reduced infectivity (35); FCV-5 is a field isolate re-
sponsible for highly virulent systemic disease in cats. During
these experiments, we noted that when the virus-receptor pre-
incubation was carried out at 37°C rather than 4°C, substan-
tially more infectivity was lost. To quantify the extent to which
FCV-5 was neutralized by preincubation with soluble receptor
TABLE 1. Data collection and refinement statistics
Parameter
Data
Data collection
Wavelength (Å) .................................................0.97925
Space group........................................................I222
Cell dimensions
a, b, c (Å) .......................................................427.08, 450.73, 467.59
,
, (°)........................................................90.0, 90.0, 90.0
Resolution (Å)...................................................30–3.6 (3.73–3.60)a
Rsym or Rmerge (%) ............................................22 (55)
Completeness (%) .............................................90.1 (87.0)
Total no. of reflections .....................................1,233,681
No. of unique reflections..................................462,277
Refinement
Resolution (Å)...................................................30–3.6
No. of reflections ...............................................462,277
Rwork/Rfree ...........................................................0.37/0.36
a Values in parentheses are for the highest-resolution shell.
5552
OSSIBOFF ET AL.
J. VIROL.
at 37°C and to assess the influence of receptor concentration
and the form of sfJAM-A ectodomain on neutralization, we
preincubated 105 PFU of FCV-5 with increasing concentra-
tions of purified GST–fJAM-A or with a soluble form of
fJAM-A in which the GST had been removed (sfJAM-A) for
30 min at 37°C. We found a clear relationship between the
concentration of soluble receptor and the extent of virus neu-
tralization with concentrations of GST–fJAM-A of 27.5 M
(Fig. 1A) and of sfJAM-A of 2.8 M (Fig. 1B), causing an
250-fold loss of viral titer. Cleavage of GST from the N
terminus of the fJAM-A ectodomain enhanced the capacity of
the protein to neutralize FCV-5 10-fold. As GST was fused to
the D1 portion of the ectodomain, which we have shown is
necessary for binding to FCV, it seems likely that GST steri-
cally hindered the FCV-ectodomain interaction.
As we had previously found only a modest reduction in
FCV-5 infectivity when virus and GST–fJAM-A were preincu-
bated at 4°C, we assessed the loss of infectivity following pre-
incubation of virus with sfJAM-A (4.3 M) at 4°C for 3 h,
followed by an additional incubation at 4, 16, 20, or 37°C for 30
min. We found a temperature dependence with the loss of
infectivity, with a 4-fold loss of infectivity at 4°C compared to
250-fold loss of infectivity at 37°C (Table 2). We conclude
that soluble forms of the fJAM-A ectodomain can substantially
neutralize FCV infectivity when preincubated at 37°C and that
neutralization is temperature and concentration dependent
and saturable.
Selection of soluble receptor-resistant (srr) mutants of FCV.
Because some residual infectivity always remained after pre-
incubation of FCV-5 with sfJAM-A at 37°C, we hypothesized
that resistant mutant viruses were being selected. As we ex-
pected that analysis of these mutants would shed light on
possible capsid residues important in binding to fJAM-A or
receptor-induced conformational changes in the capsid associ-
ated with neutralization, we selected 24 mutants that were
resistant to preincubation with GST–fJAM-A (55 M) at 37°C
for 30 min. All 24 mutants contained one or two point muta-
tions within the major capsid protein (Table 3), giving a total of
20 unique mutations. Three of these mutations were present in
the leader sequence of the capsid, which is cleaved from the
capsid precursor protein during capsid assembly by a virus-
encoded proteinase (4, 33, 54); however, the mutants that
contained mutations in the leader sequence also had an addi-
tional mutation in another region of the capsid protein. Several
residue changes were found in more than one mutant. These
changes were at positions G329 (three mutants), T438 (two
mutants), K480 (three mutants), V516 (five mutants), and
K572 (two mutants). Only two mutants had mutations within
the minor capsid protein, VP2 (Table 3), and both of these
mutants also contained mutations in VP1.
Overview of the crystal structure. To identify the location of
srr mutations on the capsid, we determined the crystal struc-
ture of FCV-5 at 3.6-Å resolution by molecular replacement
and real-space averaging (Fig. 2). The structure was refined to
a crystallographic R factor of 0.37 (Rfree 0.39) at 3.6-Å
resolution. As in other T3 viruses, the icosahedral asymmet-
rical unit consisted of three independent but chemically iden-
tical subunits (Fig. 2C). These subunits are traditionally des-
ignated A, B, and C. The present model of the FCV-5
icosahedral asymmetrical unit consists of residues 129 to 662 of
the A and C subunits and residues 133 to 662 of the B subunit.
Polypeptide fold. The structure of each subunit comprises an
NTA, an S domain, and a P domain (Fig. 2C), similar to that
FIG. 1. Neutralization of FCV-5 by sfJAM-A. FCV-5 (1 105
PFU) was incubated with various concentrations of soluble GST–
fJAM-A (A) or cleaved fJAM-A at 37°C for 30 min (B). The virus
samples were then plaque titrated on CRFK cell monolayers. Each
data point represents the average of two replicates from a single
representative experiment.
TABLE 2. Temperature dependence of FCV-5 neutralization
Incubation
temp (°C)a
Avg log titer (PFU ml1) after
incubation withb:
Change in
log titer SD
Buffer
4.4 M sfJAM-A
4
4.8
4.2
0.6 0.2
16
5.2
3.8
1.4 0.3
24
4.7
2.6
2.1 0.2
37
4.4
2.0
2.4 0.1
a All samples were incubated at 4°C for 3 h and then for 30 min at the
temperature indicated.
b Average log titer (as determined by plaque assay on CRFK cells) of at least
three replicates.
TABLE 3. Capsid mutations in srr mutants
VP1 mutation(s)a
VP2 mutationa
No. of mutants
A168S
F486L
1
T202A
T447I
1
G329D
1
G329S
1
D15N
G329S
1
E401K
I15V
1
D434N
I661T
1
T438I
N13D
1
T438A
1
N443S
2
I94T
S465N
1
K480R
1
K480Q
1
K480E
N483S
1
V516I
2
V516A
3
P116T
I535V
1
K572E
1
K572Q
1
I94T
Y575H
1
a Mutations are identified by the wild-type amino acid position in ORF 2 (VP1;
including leader sequence) or ORF 3 (VP2) of FCV-5.
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CALICIVIRUS CONFORMATIONAL CHANGE
5553
seen in the recombinant Norwalk virus capsid (rNV) and
SMSV-4 (Fig. 3C) (5, 41). Residues 170 to 329 fold into a
canonical eight-stranded
-barrel structure similar to that
found in rNV, SMSV-4, and many other viral capsid proteins.
These eight strands, conventionally denoted as B to I, form two
four-stranded sheets, BIDG and CHEF. Two helices are
situated between strands C and D and strands E and F,
respectively. Structural comparison of the S domains of
FCV-5 and SMSV-4 revealed that the most significant vari-
ations were located in the loop region connecting strands E
and F (Fig. 3C).
The P domain of FCV-5 is comprised of residues 330 to 662
and is linked to the S domain by a flexible hinge. The P domain
is subdivided into P1 and P2 subdomains. The P1 subdomain,
consisting of two segments of polypeptide (residues 330 to 381
and 551 to 662), folds into a structure similar to that seen in
SMSV-4.
The P2 subdomain, the outermost portion of the capsid
protein, is a large insertion into the P1 subdomain. The P2
subdomains of SMSV-4 and FCV-5 form a compact barrel
structure consisting of six strands, sequentially labeled as A to
F (Fig. 3A and B). However, the six strands in FCV-5 vary in
length and relative orientations, compared to their counter-
parts in SMSV-4. These six strands are connected by loops of
various lengths. Structure superimposition showed that loop
CD in FCV-5 is significantly shorter than that of SMSV-4
(Fig. 3C). The P2 subdomain consists of residues 382 to 550
and contains seven fewer residues than that of SMSV-4. Struc-
tural comparison showed that those “inserted” residues are
mainly distributed in the loop connecting strands C and D.
Structural comparison between different subunits. Pairwise
superposition calculations indicated that the structure of the P
domain of FCV-5 adopts similar structures in all three sub-
units. The S domain, however, shows larger variations between
the different subunits to facilitate the assembly of a T3 cap-
sid. In addition, the A and C subunits have four more disor-
dered N-terminal residues than the B subunit.
Superposition of the whole subunit structure revealed that
the A subunit forms a more compacted structure, as evidenced
by the closer association of the S and P domains. The interac-
tions between the S and P domains are less prominent in the B
subunit, resulting in a more relaxed structure. The C subunit
FIG. 2. X-ray structure of FCV (FCV-5) capsid. (A) Two sample regions in the calculated electron density map with modeled amino acid
residues 197 to 207 (S domain) and 586 to 596 (P domain), respectively. (B) X-ray structure of FCV-5 viewed along the icosahedral 2-fold axis.
Location of a set of A/B and C/C dimers and icosahedral 5-fold and 3-fold axes are shown. (C) Ribbon representation of the B subunit structure.
The NTA (green), S domain (blue), and P1 (red) and P2 (yellow) subdomains are indicated.
5554
OSSIBOFF ET AL.
J. VIROL.
has the most open conformation, with the fewest interactions
between the S and P domains. These differences contribute to
the bent and flat conformations of the A/B and C/C dimers,
respectively (Fig. 3D).
Mapping of neutralizing epitopes and conserved surface
regions on the P2 subdomain of FCV-5. Hypervariable regions
(C and E regions) of the FCV linear capsid sequence were
previously identified, and neutralizing epitopes were mapped
FIG. 3. Ribbon representation of the P2 subdomain in FCV-5 (A) and SMSV-4 (B). The
-strands are labeled from A to F in each case.
(C) The superposition of the P domain of FCV-5 (red) and SMSV-4 (blue) shows that the six
-strands in the P2 subdomains are similarly spatially
arranged into a compact barrel, despite the low sequence similarity. Arrows indicate the loops connecting
-strands C and D, and E and F, of the
FCV-5 P2 subdomain. (D) The FCV-5 A/B dimer (top) as viewed along the line joining the icosahedral 3- and 5-fold axes and the C/C dimer
(bottom) as viewed along the line joining the adjacent icosahedral 3-fold axes demonstrate the bent and flat conformations, respectively, assumed
by the subunits. In both cases, the dimeric 2-fold axis is vertical. (E to G) Positions of antigenic sites and conserved residues on the P2 subdomain
of the C/C dimer of FCV-5. (E) Neutralizing epitopes mapped on P2 (gray). Antigenic sites (Ags1 to -4; colored pink) represent B-cell linear
epitopes identified by Radford et al. (43). MAb escape mutations identified by Tohya et al. are colored red (58). (F) Degree of conservation of
surface-exposed residues on the P2 subdomain as predicted by Consurf 3.0. The solid yellow line indicates dimer interface; the dashed yellow line
indicates the axis of conservation. (G) Locations of N-terminal and C-terminal HVRs, region C and central conserved portion of region E mapped
on surface of P2 subdomain.
VOL. 84, 2010
CALICIVIRUS CONFORMATIONAL CHANGE
5555
predominantly to residues within the E region (Fig. 3G) (43,
48, 57, 58). Using the FCV-5 structure, we mapped the loca-
tions of four linear B-cell epitopes identified in FCV-F9 (43).
All of these sites lay on the margins of the P2 subdomain, with
one site, Ags4, at the dimer interface (Fig. 3E). Similarly,
antibody escape mutations found in mutants of FCV-F4 are
also located on the margins of the P2 subdomain within hy-
pervariable regions (Fig. 3E) (58). To more accurately assess
the conservation of individual surface-exposed residues on the
P2 subdomain, we used Consurf 3.0, which maps the rate of
evolution as determined by an empirical Bayesian estimation
among homologous proteins onto the three-dimensional (3-D)
structure (26). As input, we used an alignment of 47 different
FCV capsid sequences to derive an evolutionary conservation
score that was used to color code individual residues on the
surface of the FCV-5 capsid structure (Fig. 3F). This analysis
revealed a patchwork of conserved surface residues along the
dimeric interface of the P domain that also extended laterally
at a 45° angle to the dimer interface on the surface of each
monomer. Highly variable residues on the margins of the P2
subdomain surrounded the more central conserved residues.
Mapping srr mutations on the FCV-5 structure. Sequencing
of the capsid protein of the 24 srr mutants revealed 17 unique
amino acid changes from the parental FCV-5 mature capsid
protein sequence. We mapped the locations of these changed
residues on the structure of FCV-5 (Fig. 4). Three mutations
were found in the NTA and S domains, and 14 mutations were
present in the P domain (Fig. 4A). Of the 24 mutants, 21
FIG. 4. Positions of mutations found in srr mutants in the structure of FCV-5. (A) The residues mutated in the srr mutants are illustrated on
a schematic of the FCV capsid protein. The locations of the different structural domains in the primary capsid sequence are indicated.
(B) Locations of the mutations in the FCV-5 atomic resolution structure.
5556
OSSIBOFF ET AL.
J. VIROL.
(87.5%) contained mutations that mapped to the P domain of
the capsid, and 18 of those mutations were to residues within
the P2 subdomain.
Of the mutations found in the NTA and S domains, one
change was to a residue (A168S) within the NTA, but this
mutation was accompanied by a second amino acid change in
the P2 domain (F486L). Similarly, a mutant with a change in
the shell domain (T202A) also had a change in the P2 domain
(T447I). Three srr mutants contained mutations at residue
G329 in the shell domain on a loop that connects the shell with
the protrusion domain. Residue G329 is located at the point of
the flexible hinge, and mutations at G329 were present alone
or in addition to a change in the leader sequence of the capsid
(D15N) (Fig. 4B and Table 3). Mutations to residue G329 were
the only changes identified in the S domain that occurred
independently of an additional mutation within the P domain.
Three of the 21 mutated residues identified in the P domain
were present within the P1 subdomain (K572E/Q, Y575H, and
I661T). Residues K572 and Y575 were buried at the dimer
interface of the capsid structure (Fig. 4B). Residue I661 was
located at the base of the P1 subdomain, in close proximity to
the S domain (Fig. 4B); the mutant containing the I661T
change also had a second change within the P2 domain
(D434N). The other 11 P domain mutations were in the P2
subdomain. Eight of these mutated residues were within sur-
face-exposed loops of P2: E401K, D434N, T438A, N443S,
T447I, S465N, K480R/Q/E, V516I/A, and I535V (Fig. 4B).
Three of the eight surface-exposed mutations were present in
more than one srr mutant: N443 (two mutants), K480 (three
mutants), and V516 (five mutants). Two mutants had muta-
tions to residues at the P2 dimer interface; one mutant pos-
sessed a change in the surface-exposed residue N483, while
another mutant contained a change to the buried residue F486.
Lastly, two mutants had mutations to residue T438 that is
buried in the present structure and maps to a loop near the
surface of the P2 subdomain. In summary, the location of
mutations present in srr isolates mapped to three general re-
gions of the structure of FCV-5: (i) surface-exposed residues
on the P2 domain; (ii) residues that are buried or not easily
accessible from the surface of the capsid; and (iii) the hinge
region of the capsid.
srr mutants are resistant to neutralization by sfJAM-A but
retain the capacity to bind fJAM-A and require cellular
fJAM-A for infection. From the original 24 srr mutants, we
selected eight mutant viruses to further characterize. We se-
lected these mutants based on the location of their mutated
residues in an effort to have representative mutations from all
regions of VP1. Additionally, we selected mutants based on the
frequency that particular residues were mutated, with overrep-
resented mutations to residues included. All of the eight mu-
tant viruses in the subpanel were resistant to neutralization
with sfJAM-A under the conditions in which they were se-
lected (Fig. 5). To verify that the eight selected srr mutants still
bound to fJAM-A expressed on the cell surface, we used flow
cytometry to measure the capacity of each of the mutants to
bind to CHO-S cells expressing fJAM-A on their surface (Fig.
6A and B). We used CHO-S cells transiently expressing
fJAM-A rather than a stable fJAM-A-expressing CHO-K1 cell
line that we previously described (35), because of technical
difficulties using adherent CHO-K1 cells in flow cytometry
assays. We found that all eight mutants bound to CHO-S cells
expressing fJAM-A. One mutant, the K572E mutant, however,
bound to 50% fewer cells than did the parental FCV-5. In
addition, all eight mutants still required fJAM-A as a func-
tional receptor, as pretreatment of CRFK cells with a rabbit
antiserum against fJAM-A completely blocked infection (data
not shown). We conclude from these findings that the subpanel
of eight srr mutants still binds and requires cellular fJAM-A for
cell infection.
At low MOI, the growth of some srr mutants differs from
that of parental FCV-5. To evaluate the growth properties of
the srr mutants, we generated single- and multiple-cycle growth
curves (Fig. 7A and B, respectively) for each of the eight
selected srr mutants. The growth kinetics and final yields fol-
lowing a single replicative cycle (MOI 5) for the mutant
viruses were similar to those of the parental FCV-5 (Fig. 7A
and data not shown). However, we found that during multiple
replicative cycles (MOI 0.01) six of the mutants (G329D,
V516I, K572E, T438I, N443S, and E401K mutants) had signif-
icantly decreased infectivity at 4 h pi compared to parental
FCV-5 as determined by ANOVA (P 0.005) (Fig. 7B and
data not shown). Two of the eight mutants (K480Q and A168S/
F486L mutants) were similar to FCV-5, producing new infec-
tious virions by 4 h pi. By 8 h pi, all of the viruses had attained
similar increases in titer, and the final yields were similar. We
conclude that six of the panel of eight srr mutants have slower
growth kinetics at early times postinfection than the parental
FCV-5 in cells infected at a low multiplicity.
Binding of srr mutants to fJAM-A. In order to assess the
kinetics of binding of the srr mutants to fJAM-A, we purified
three mutants (G329D, V516I, and K572E mutants) and com-
pared their in vitro binding to plate-bound sfJAM-A by ELISA
to that of parental FCV-5 (Fig. 8A). All the viruses showed
saturable concentration-dependent binding to fJAM-A. The
binding kinetics of FCV-5 and the G329D mutant were similar,
and saturation of binding to fJAM-A occurred at similar con-
FIG. 5. Neutralization of srr mutants by sfJAM-A. A subpanel of
eight srr mutants and FCV-5 (1 105 PFU) was incubated in the
presence or absence of soluble GST–fJAM-A (55 M) at 37°C for 30
min. The infectivity of each sample was assayed by plaque titration.
The change in log titer was calculated by subtracting the titer of
samples incubated with receptor from that of samples incubated with-
out receptor. The mean change in log10 titer standard deviation
(SD) of three replicates of a representative experiment is shown.
VOL. 84, 2010
CALICIVIRUS CONFORMATIONAL CHANGE
5557
centrations of virus. The binding of V516I and K572E mutants
to plate-bound fJAM-A saturated at slightly higher virus con-
centrations than did those of FCV-5 and the G329D mutant.
One possible explanation for this difference is differential de-
tection by the polyclonal rabbit FCV-5 antisera. We conclude
that G329D, V516I, and K572E srr mutants and FCV-5 all bind
immobilized recombinant fJAM-A with relatively similar ki-
netics under in vitro conditions.
Relative affinities of srr mutants for fJAM-A. To determine
if the V516I, K572E, and G329D srr mutants differed from
FCV-5 in their affinity for fJAM-A, we performed a competi-
tion ELISA (Fig. 8B). Different concentrations of sfJAM-A
were used to compete for binding of virus to plate-bound
fJAM-A at 4°C. We found that binding of FCV to the plate-
bound receptor was strongly inhibited at the highest concen-
trations of sfJAM-A. The binding of FCV to plate-bound
fJAM-A correlated inversely to the concentration of soluble
receptor. A 50% decrease in binding of all the tested viruses to
plate-bound fJAM-A occurred at concentrations of sfJAM-A
between 0.33 and 0.66 M. All the viruses exhibited similar
competition curves; however, binding of the V516I mutant to
plate-bound fJAM-A was 50% inhibited at a slightly higher
concentration of sfJAM-A, as can be seen by the rightward
FIG. 6. Binding of srr mutants to CHO cells expressing fJAM-A.
(A) Nonpermissive CHO-S cells were transfected with a DNA con-
struct encoding fJAM-A. At 24 h posttransfection, FCV (MOI 5)
was adsorbed to the cells on ice for 30 min. After being washed with
cold PBS to remove unbound virus, the cells were fixed and bound
virus and cell surface fJAM-A were detected with mouse anti-FCV
MAb and rabbit anti-fJAM-A antibodies, followed by Alexa 488-con-
jugated goat anti-mouse IgG and Alexa 647-conjugated goat anti-
rabbit IgG. Virus binding and receptor expression were analyzed by
flow cytometry. Flow cytometry dot plots for four representative sam-
ples are shown. (B) Virus binding was assayed by determining the
percentage of fJAM-A-positive cells that bound virus. The means of
the results for three independent experiments (two samples of 1 104
cells per experiment) SE are shown.
FIG. 7. Growth of srr mutants. CRFK monolayers were infected
with FCV-5 and the indicated srr mutants under single-cycle (MOI
5) (A) or multiple-cycle (MOI 0.01) (B) conditions. For clarity, the
data from only four mutants have been included in each graph. The
change in virus titer was determined by plaque assay. The mean log10
titer (log10 titer at each time point log10 titer at T 0) for each time
point is shown. The error bars shown at 4 and 8 h pi for the multiple-
cycle growth represent the standard deviation of four replicates; for
other data points, error bars have been omitted from the figure for
clarity. The asterisks indicate significant differences between viral iso-
lates as determined by ANOVA at these time points.
5558
OSSIBOFF ET AL.
J. VIROL.
shift in the competition binding curve (Fig. 8B). Statistical
analysis (ANOVA) at three concentrations of soluble receptor
(0.04, 0.33, and 2.7 M) showed significantly more viral bind-
ing of the V516I mutant than that of FCV-5 for all but the
lowest concentration of receptor (P 0.05). We found that
preincubation of virus with low concentrations of sfJAM-A
significantly enhanced binding to plate-bound fJAM-A com-
pared to virus alone. As fJAM-A can form dimers, and FCV
binding does not seem to require or occlude the fJAM-A
dimerization interface (1, 35), this enhanced binding may be
mediated by fJAM-fJAM interactions between immobilized
and virus-bound soluble receptor. An alternative explanation is
that the binding of sfJAM-A to FCV capsids enhances the
binding to the immobilized receptor allosterically. We con-
clude that the G329D, V516I, and K572E srr mutants demon-
strate similar affinities for fJAM-A.
Bis-ANS binding indicates a change in FCV surface hydro-
phobicity following preincubation with sfJAM-A. We hypoth-
esized that the interaction of soluble receptor with FCV caused
a change in conformation of the viral capsid. We, therefore,
used binding of bis-ANS as a probe to investigate changes in
hydrophobicity that occurred upon interaction of FCV with
fJAM-A in solution. Bis-ANS is a fluorophore that binds to
exposed hydrophobic regions of protein molecules (44). The
sequestration of bis-ANS in hydrophobic pockets of proteins is
accompanied by a large increase in the fluorescence quantum
yield of the probe. Thus, bis-ANS can be used to probe for
structural changes in viral proteins that are accompanied by
changes in surface hydrophobicity (10, 14, 15). We first prein-
cubated virus or receptor alone in a cuvette in the spectroflu-
orometer at 37°C for 10 min and then added bis-ANS and
collected fluorescence intensity readings over a period of 10
min at 37°C (Fig. 9A to C). To quantify differences in bis-ANS
fluorescence, we waited until the fluorescence emission read-
ings had stabilized and then averaged the fluorescence inten-
sity measurements over 1 min. Fluorescence intensity in-
creased during the first 5 to 10 min after addition of bis-ANS
to any of the protein samples, but it stabilized after 10 min.
This increase in fluorescence is likely caused by bis-ANS bind-
ing to surface-exposed hydrophobic pockets on the proteins
(Fig. 9A to C).
To determine if the interaction of FCV with fJAM-A trig-
gered a conformational change that would change the binding
of bis-ANS to the proteins, we preincubated FCV with either
hJAM-A (hJAM-A does not bind FCV [35; Fig. 9A and D]) or
fJAM-A (Fig. 9B and E) at 37°C for 10 min and then added
bis-ANS and monitored changes in fluorescence intensity over
10 min. We found that the average stabilized bis-ANS fluores-
cence intensity of samples containing FCV-5 plus hJAM-A was
equal to the sum of the intensities associated with incubation
of bis-ANS with FCV-5 and hJAM-A individually (Fig. 9D).
These findings were expected and indicate that the overall
binding of bis-ANS to either protein was unchanged when they
were mixed. In contrast, the average stabilized fluorescence
intensity of samples containing FCV-5 plus fJAM-A was
greater than the sum of the bis-ANS fluorescence intensities of
the two proteins when incubated individually (Fig. 9E). This
finding suggests that an interaction between virus and fJAM-A
changed the conformation of one or both proteins, resulting in
the exposure of more hydrophobic patches on the protein and
thus an increase in the amount of bis-ANS binding and result-
ing fluorescence.
As we found that FCV-5 neutralization by sfJAM-A was
temperature dependent (Table 2), we investigated if changes in
bis-ANS binding to the fJAM-A–FCV-5 mixture were affected
by temperature (compare Fig. 9E and F). We found that,
unlike our findings at 37°C (Fig. 9E), the fluorescence intensity
of samples of FCV-5 and fJAM-A preincubated together at
4°C was less than the sum of the bis-ANS intensities of virus
and receptor incubated at 4°C with the fluorescent probe alone
(Fig. 9F). Our interpretation of these findings is that binding of
FIG. 8. Binding of srr mutants to immobilized fJAM-A in the ab-
sence (A) or presence (B) of increasing concentrations of sfJAM-A.
ELISA plates were coated with 100 ng of sfJAM-A ectodomain. Var-
ious concentrations of purified FCV-5 and G329D, V516I, and K572E
srr mutants were incubated with the immobilized protein for 3 h on ice.
To investigate the effect of sfJAM-A on the binding of virus to immo-
bilized receptor, 750 ng of each virus and various concentrations of
soluble receptor were incubated together with plate-bound fJAM-A
for 3 h on ice. Bound FCV was detected with rabbit anti-FCV serum,
followed by HRP-conjugated goat anti-rabbit IgG. Colorimetric HRP
substrate was added, and the amount of bound FCV-5 was quantified
by absorbance at 595 nm. The means of three and four plates, respec-
tively (two replicates per plate), SE are shown. ANOVA was per-
formed on three concentrations of soluble receptor (0.04, 0.33, and 2.7
M) to determine statistical differences; significant differences are
indicated by asterisks.
VOL. 84, 2010
CALICIVIRUS CONFORMATIONAL CHANGE
5559
FIG. 9. Changes in bis-ANS binding upon mixing of FCV with sfJAM-A. At time zero, bis-ANS was added to the indicated purified virions,
soluble JAM-A (human [A] or feline [B and C]), or virions preincubated with JAM-A. Fluorescence was recorded every 2 s continuously for 10
min at excitation and emission wavelengths of 395 and 495 nm, respectively. (D to G) Stabilized fluorescence intensities measured during the last
minute for each sample were averaged. Broken lines indicate the additive average fluorescence intensity measured for both native virus and
receptor. The means SE (n 3) are shown.
5560
OSSIBOFF ET AL.
J. VIROL.
fJAM-A to FCV-5 at 4°C blocks hydrophobic patches previ-
ously available for bis-ANS binding on the virus and/or
fJAM-A. In contrast, the increase in binding of bis-ANS to the
fJAM-A–FCV mixture at 37°C indicates a change in confor-
mation of the viral capsid and/or the receptor.
The binding of bis-ANS at 37°C to V516A, K572E, and
G329D srr mutants and to field isolates Kaos and FCV-131 was
also investigated. Following incubation of virus and receptor at
37°C, these samples showed an increase in fluorescence inten-
sities similar to that seen for FCV-5 (Fig. 9C and G). In
contrast, incubation of the vaccine strain F9 with fJAM-A led
to a fluorescence intensity maximum that was less than what
would be expected from the sum of the intensities associated
with the receptor and virus alone; this pattern was similar to
that of FCV-5 incubated with receptor at 4°C. From these
findings, we conclude that interaction of fJAM-A with FCV at
37°C leads to changes in the binding of bis-ANS that are
indicative of changes in hydrophobicity of virus and/or recep-
tor. For FCV-5, these changes in bis-ANS binding are depen-
dent upon temperature, as virus and receptor incubated at 4°C
did not display the same increase in fluorescence intensity.
Not all FCV isolates are neutralized by sfJAM-A. The
FCV-5 isolate that we used to select for srr mutants was re-
covered from a cat suffering from severe systemic disease. As
this form of FCV disease is uncommon, we hypothesized that
not all isolates of FCV would be neutralized by incubation with
soluble receptor at 37°C. To test this hypothesis, we incubated
105 PFU of six additional FCV isolates with 45 M GST–
fJAM-A at 37°C; these virus isolates included the vaccine
strain F9, two additional isolates recovered from cats with
virulent systemic disease, the FCV-Urbana isolate, and three
other field isolates recovered from cats with mild- to moderate-
severity FCV disease. Several additional FCV isolates were
neutralized by preincubation with GST–fJAM-A at 37°C; how-
ever, the vaccine strain F9, FCV-Urbana, and the two isolates
recovered from cats with milder disease were not neutralized
under these conditions (Fig. 10). We conclude that there are
natural differences among FCV isolates in their responses to
incubation with sfJAM-A at 37°C.
DISCUSSION
Several nonenveloped viruses are neutralized by preincuba-
tion with their functional receptors (23, 31, 49). However, the
extent to which infectivity is lost at physiologic temperatures
depends upon the mechanism of receptor-mediated neutral-
ization. For polioviruses and some HRVs, preincubation with
their receptors induces irreversible inactivating conformational
changes in the virus (6, 17). We found that some but not all
isolates of FCV were neutralized by preincubation with a sol-
uble form of fJAM-A. Although it is possible that neutraliza-
tion was due to steric hindrance or occupation of receptor-
binding sites, the prominent temperature dependence of
neutralization strongly suggests that receptor-induced confor-
mational changes are involved. Receptor-induced conforma-
tional changes in both HRVs and poliovirus are temperature
dependent (16, 20). Polioviruses undergo an irreversible neu-
tralizing transition from a 160S particle to a 135S particle when
they are incubated with soluble receptor at 37°C, but this
transition does not occur at temperatures below 33°C (16, 61).
In contrast, although FCV-5 receptor-mediated neutralization
was clearly temperature dependent, no obvious temperature
threshold was observed. Rather, a gradual diminishment of the
neutralizing effect as the temperature decreased from 37°C to
4°C was found (Table 2). We propose that some FCV isolates
undergo a temperature-dependent, fJAM-A-mediated neutral-
ization following incubation at 37°C; however, as not all FCV
isolates undergo neutralization under these conditions, an ad-
ditional cellular factor or environmental condition is likely
required by some FCV isolates for neutralization.
Analysis of soluble receptor-resistant (srr) mutants in polio-
virus and murine coronavirus identified virus residues that
either directly or indirectly facilitated receptor binding (6, 23,
46). Of the 24 srr mutants of FCV-5, mutations to 20 unique
VP1 residues were identified; six mutations mapped to surface-
exposed residues on the P2 domain of the capsid that poten-
tially could directly interact with the receptor. Several mutated
residues were buried or inaccessible from the surface of the
structure and could possibly indirectly affect binding. Similarly,
poliovirus srr mutants contained mutations to both internal
and surface-exposed residues. The mutated residues for polio-
virus mapped to a hydrophobic pocket in the structure. It is
believed that changes to residues in this hydrophobic pocket
regulate receptor attachment and conformational changes
within the virus particle (6, 42).
One to two copies of the VP2 protein are found in FCV
virions, and VP2 is essential for the synthesis and maturation
of infectious FCV virions (52, 53). It was, therefore, important
to ensure that resistance to neutralization of the srr mutants
was not due to changes to VP2. Sequencing of the ORF en-
coding the minor capsid protein of FCV revealed that only two
mutants of the 24-mutant panel contained coding mutations to
VP2. As both mutants possessing VP2 mutations also had VP1
mutations, and the remainder of the srr mutants contained only
FIG. 10. Ability of sfJAM-A to neutralize select FCV isolates. A
panel of six FCV field isolates (FCV-5, Kaos, Deuce, FCV-127, FCV-
131, and FCV-796) and two tissue culture-adapted strains (F9 and
Urbana) (1 105 PFU) was incubated in the presence or absence of
soluble GST–fJAM-A (45 M) at 37°C for 30 min. The remaining
infectivity in each sample was assayed by plaque titration. The change
in log titer was calculated by subtracting the titer of samples incubated
with receptor from the titer of samples incubated without receptor.
The mean change in log10 titer SD (n 4) is shown.
VOL. 84, 2010
CALICIVIRUS CONFORMATIONAL CHANGE
5561
VP1 mutations, changes to VP2 cannot explain the resistance
of the mutants to receptor-mediated neutralization.
Our analysis of the FCV srr mutants indicates that residues
within critical regions of the capsid are necessary for structural
flexibility or receptor interactions. In particular, changes to
residue G329 (mutated in three srr mutants) suggest that
movement between the S and P domains of the capsid protein
occurs upon receptor interaction and is involved in receptor-
mediated neutralization. The glycine residue at position 329 is
100% conserved in the FCV capsid sequence and is located in
a short loop between the S and P domains of the capsid pro-
tein. A similar glycine residue is found in this hinge region of
the capsid in all caliciviruses. Cryo-EM reconstructions of mu-
rine norovirus in complex with neutralizing Fab fragments
show that this hinge region is extended, moving the P domain
radially away from the surface of the shell domain (24). In
addition, a cryo-EM structure of FCV-F9 complexed with the
fJAM-A ectodomain showed that the P domain of the capsid
moved with respect to the S domain when bound to fJAM-A,
indicating that movement in this region occurs (1). Our find-
ings further support the hypothesis that flexibility in this hinge
region is important to permit conformational changes of the
structural domains of the calicivirus capsid in relation to each
other and indicate that this region actively moves during re-
ceptor interactions. Moreover, our findings indicate that
changes at residue G329 substantially alter the susceptibility of
FCV to neutralization by sfJAM-A.
Several srr mutants had mutated residues at the P domain
dimer interface. These mutations could alter receptor binding
indirectly, or these residues could become accessible for re-
ceptor binding following a conformational change of the cap-
sid. The altered residue in the K572E srr mutant is in P1 and is
unlikely to be accessible from the surface of the capsid. This
mutant had decreased binding to fJAM-A on cells. However,
the K572E mutant had binding kinetics similar to that of pa-
rental FCV-5 in in vitro ELISA binding assays. A possible
explanation for this difference is that unlike fJAM-A purified
from bacteria, JAM-A on cells is N-glycosylated (32).
Poliovirus srr mutants had reduced binding affinity for their
receptor (6). It was, therefore, unexpected that only one mu-
tant (the K572E mutant) had decreased cell surface fJAM-A
binding and that none of the FCV srr mutants we examined in
detail had a substantive change in binding to fJAM-A in vitro.
In fact, the V516I mutant showed higher affinity for immobi-
lized fJAM-A (Fig. 8B). These findings indicate that resistance
of the srr mutants to receptor-induced neutralization cannot be
explained by changes in their binding affinity for fJAM-A.
Single-step growth kinetics of poliovirus srr mutants were
similar, but all mutants had lower titers than did wild-type
P1/Mahoney poliovirus at early time points postinfection. This
delay in replication was attributed to an assembly defect, as all
the srr mutants bound as efficiently as wild-type poliovirus to
cells at room temperature (6). In contrast, we found that the
FCV srr mutants and parental FCV-5 had similar single-step
growth kinetics and titers throughout the replicative cycle. We
found a difference during multiple-cycle growth experiments,
where six of the eight srr mutants investigated had significantly
decreased titers in comparison to parental FCV-5 at 4 h pi.
One explanation for these findings is decreased efficiency of
viral entry (receptor binding, cellular trafficking, and/or un-
coating) of the mutants with a consequent lag in the onset of
viral replication. As small differences in the efficiency of entry
may be masked when a high MOI is utilized, experiments using
low input titers of virus can be more sensitive. The K572E
mutant demonstrated decreased binding to cell surface-ex-
pressed fJAM-A and also had delayed multiple-cycle growth
kinetics. Therefore, the decreased early growth kinetics of this
mutant could be due to decreased receptor binding. However,
G329D and V516I srr mutants bound cell-expressed and plate-
bound fJAM-A to the same extent as did parental FCV-5 or
had increased affinity for plate-bound fJAM-A and also had
delayed early growth kinetics. We previously showed that low-
er-virulence-field or tissue culture-adapted FCV isolates had
delayed multiple cycle growth kinetics compared to more-vir-
ulent isolates (including FCV-5) (36). Although the virulence
of these srr mutants is currently unknown, it is possible that by
selecting for srr mutants, we may have also selected for viruses
with altered virulence.
Following attachment, nonenveloped viruses must undergo
conformational alterations that mediate membrane penetra-
tion and deliver their genetic material (60). The temperature
dependence of the sfJAM-A neutralization of FCV and the
identification of srr mutant residues buried in the capsid struc-
ture suggested that fJAM-A induces a conformational change
in the FCV capsid. Our finding that mixtures of FCV-5 and
sfJAM-A incubated at 37°C had bis-ANS fluorescence inten-
sities greater than the sum of the fluorescence intensities as-
sociated with virus and sfJAM-A individually provided further
support for this concept. Bis-ANS binding has previously been
used to detect conformational changes associated with changes
in hydrophobicity in the envelope glycoproteins of Newcastle
disease virus and HIV-1 upon incubation with gangliosides and
CD4, respectively (14, 22). Additionally, our finding that pre-
incubation of FCV-5 and fJAM-A at 4°C resulted in lower
bis-ANS fluorescence than the sum of the intensities of the two
components is consistent with the idea that binding of fJAM-A
to FCV-5 at 4°C occludes potential bis-ANS binding sites and
that exposure of new hydrophobic patches as a consequence of
conformational change does not occur or is decreased at 4°C
(Fig. 9E and F). An alternative explanation is that the in-
creases in hydrophobicity we observed were due to changes in
the receptor, not the virus. However, as there is substantial
evidence for viral protein conformational changes following
interaction with cellular receptors, and such conformational
changes are likely a requirement of all nonenveloped viruses,
this explanation is less likely. For those viruses that when
mixed with sfJAM-A showed an increase in total bis-ANS
fluorescence intensity, the intensity was 11 to 30% greater for
the mixtures of FCV and sfJAM-A than the sum of the bis-
ANS fluorescence intensities of each component incubated
separately. Although this is an in vitro assay, we believe that
taken together with our other data, these changes are biolog-
ically significant. Additional investigation will be required to
determine the nature of the conformational changes and their
functional significance.
Upon incubation with soluble receptor, the srr mutants and
parental FCV-5 demonstrated similar increases in hydropho-
bicity. These findings raise the possibility that FCV neutraliza-
tion by soluble receptor does not involve a conformational
change in the capsid. However, a more likely possibility is that
5562
OSSIBOFF ET AL.
J. VIROL.
more than one intermediate capsid conformer forms upon
interaction of FCV with fJAM-A. Following interaction of
picornaviruses with their functional receptors, three interme-
diate conformers of the capsid are formed—the 135S altered
particle and two forms of the 80S empty particle (7, 9, 13, 27,
28). Therefore, our working model is that mutants resistant to
neutralization undergo an initial conformational change upon
interacting with fJAM-A that either is reversible or results in a
stable, infectious intermediate.
A cryo-EM reconstruction of the FCV-F9 strain complexed
with fJAM-A at 18 Å revealed that fJAM-A interacted with the
top of the P2 subdomain of the F9 capsid, causing an 13°
counterclockwise rotation of the P dimers about their 2-fold
axes of symmetry (1). Interestingly, we found that in contrast to
the other FCV isolates we investigated, the F9–sfJAM-A com-
plex bound less bis-ANS than did the F9 virus alone (Fig. 9G).
Given the structural evidence for a conformational change in
the F9 capsid, one possible explanation is that fJAM-A binding
induces a conformational change that decreases capsid surface
hydrophobicity. In the face of our observations of increased
bis-ANS binding to other FCV isolates when preincubated
with fJAM-A, it is possible that the capsid conformational
changes reported by Bhella et al. (1) for the vaccine isolate F9
differ from changes that occur in the capsids of other FCV field
isolates.
We found that some FCV isolates, including low-passage
field isolates and tissue culture-adapted lab strains, were not
susceptible to fJAM-A neutralization (Fig. 10). A natural iso-
late that was resistant to receptor neutralization, FCV-131,
also exhibited increased hydrophobicity following incubation
with soluble receptor. These findings suggest that similar to the
srr mutants, field isolates undergo a conformational alteration
following interaction with fJAM-A but do not lose infectivity.
Other isolates, however, may undergo more drastic, global
conformational alterations that result in either a complete loss
of or significant decrease in infectivity. Interestingly, we found
that sensitivity of different FCV isolates to neutralization cor-
related with their virulence in cats. FCV is a common pathogen
of cats that typically causes mild or unapparent disease, with
fatal disease being unusual. However, highly virulent isolates
of FCV cause systemic signs of disease and result in mortality
rates as high as 50% (21, 38, 39, 47). In our panel, the most
virulent isolates were susceptible to fJAM-A neutralization,
while low-virulence isolates were resistant. It is possible that
the in vitro assay of receptor neutralization is highlighting an
important determinant of in vivo virulence.
ACKNOWLEDGMENTS
We thank Christian Nelson, Meg Crapster-Pregont, Sarah Caddy,
and Rachel Mays for excellent technical assistance. We thank Terry
Dermody and Kristen Guglielmi for the generous gift of reagents and
Stanislav Sosnovtsev for technical advice. We acknowledge the SBC-
CAT 19ID beamline at the Advanced Photon Source (supported by
the U.S. Department of Energy, Basic Energy Sciences, Office of
Science, under contract no.W-31-109-Eng-38) and its staff for their
help during data collection.
This work was supported by grants from The Cornell Feline
Health Center and the Winn Foundation to J.S.L.P., National In-
stitutes of Health (PO1 AI057788), and R. Welch Foundation (Q-
1279) to B.V.V.P. R.J.O. is the recipient of a scholarship from
Cornell University.
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