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11319941 | Complex trait analysis of the mouse striatum: independent QTLs modulate volume and neuron number
Abstract
Background
The striatum plays a pivotal role in modulating motor activity and higher cognitive function. We analyzed variation in striatal volume and neuron number in mice and initiated a complex trait analysis to discover polymorphic genes that modulate the structure of the basal ganglia.
Results
Brain weight, brain and striatal volume, neuron-packing density and number were estimated bilaterally using unbiased stereological procedures in five inbred strains (A/J, C57BL/6J, DBA/2J, BALB/cJ, and BXD5) and an F2 intercross between A/J and BXD5. Striatal volume ranged from 20 to 37 mm3. Neuron-packing density ranged from approximately 50,000 to 100,000 neurons/mm3, and the striatal neuron population ranged from 1.4 to 2.5 million. Inbred animals with larger brains had larger striata but lower neuron-packing density resulting in a narrow range of average neuron populations. In contrast, there was a strong positive correlation between volume and neuron number among intercross progeny. We mapped two quantitative trait loci (QTLs) with selective effects on striatal architecture. Bsc10a maps to the central region of Chr 10 (LRS of 17.5 near D10Mit186) and has intense effects on striatal volume and moderate effects on brain volume. Stnn19a maps to distal Chr 19 (LRS of 15 at D19Mit123) and is associated with differences of up to 400,000 neurons among animals.
Conclusion
We have discovered remarkable numerical and volumetric variation in the mouse striatum, and we have been able to map two QTLs that modulate independent anatomic parameters.
Background
The dorsal striatum is a massive nucleus in the basal forebrain that plays a pivotal role in modulating motor activity and higher cognitive function. Approximately 90% of all neurons in the striatum - 1.5 to 2.5 million in mice [1,2] and 110-200 million in humans [3,4] - belong to an unusual type 'of inhibitory projection cell referred to as medium spiny neurons [5-8]. Striatal neurons are divided into two major subpopulations (patch and matrix), that have somewhat different gene expression profiles and have different patterns of pre- and postsynaptic connections [9-13].
Numbers of medium spiny neurons and ratios of these and less numerous striatal interneurons are critical variables that influence motor performance and aspects of cognition. In the case of Huntington disease the loss of 15-30% of the normal complement of medium spiny neurons leads to distinct movement disorder in both humans and transgenic mouse models [14-17]. The subset of genes that normally control the proliferation, differentiation, and survival of striatal neurons [18-20] are therefore of considerable importance in ensuring adaptive behavior at maturity.
In this study we use a forward genetic approach [21, 22] to begin to map and characterize members of the subset of normally polymorphic genes that specifically modulate the production and survival of striatal neurons. Our analysis of the neurogenetic control of striatal cell populations relies on the combination of two complementary quantitative approaches. The first of these, complex trait analysis, is a comparatively new genetic method that makes it possible to map individual loci that underlie polygenic traits [22,23]. The second consists of a set of unbiased stereological procedures that can be used to obtain cell counts accurately and efficiently from large numbers of cases [24,25].
From a technical point of view the mouse striatum has several advantages that make it an excellent target for complex trait analysis of the mammalian CNS. First, it is a large, cytoarchitectonically distinct region comprising approximately 5-6% of the volume of the mouse brain. Second, the dorsal striatum has a comparatively homogenous cellular composition, potentially reducing the number of quantitative trait loci (QTLs) that affect striatal neuron number. Finally, recent experiments on the molecular control of telencephalic development have highlighted a number of genes that influence neuron proliferation and differentiation of the striatum and other neighboring forebrain structures [18][26-30].
We report here both neuroanatomic and genetic quantitative evidence that the size of the striatum and the number of neurons contained within it are modulated independently.
Results
The results are divided in two sections. The first is a biometric analysis of variation of size and neuronal populations in mouse striatum. The second section is a quantitative genetic dissection and QTL analysis of variation in the size and neuronal population of the mouse striatum.
Phenotypes
Strain differences
Brain Weight and Striatal Volume. Five strains were chosen to represent low, mid, and high brain weights (Fig 1A). Brain weights of the two high strains, BXD5 (540 ± 9 mg SEM) and BALB/cJ (527 ± 13 mg), are significantly greater (P < .001) than that of C57BL/6J (476 ± 5 mg). Similarly, the brain weights of A/J (395 ± 5 mg) and DBA/2J (403 ± 5 mg) are significantly lower than those of the other three strains (P < .001). As anticipated, differences in striatal volume correspond well with differences in brain weight and volume. The striata of BXD5 (31.0 ± 0.9 mm3) and BALB/cJ (32.5 ± 1.6 mm3) mice are significantly larger (P < .001) than those of C57BL/6J (23.6 ± 0.6 mm3), A/J (21.7 ± 1.0 mm3), and DBA/2J (22.8 ± 0.6 mm3) strains.
Figure 1
Histograms of histologic phenotypes for inbred strains of mice. A. Brain weight differences are apparent between the three categories (F4,28 = 82.1, P < .001). Asterisk indicates significant differences on post-hoc tests (P < .005). B. Striatal volume. There are significant differences in striatal volume between the low brain weight strains (A/J, DBA.2J) and the high brain weight strains (BALB/cJ, BXD5). C57BL6/J mice differ from the high, but not low brain weight strains (F4,28 = 28.9, P < .001). Asterisk indicates significant differences on post-hoc tests (P < .005). C. Neuron-packing density in the striatum. In general, brains with smaller striata have greater neuron-packing density (F4,28 = 17.6, P < .001). Asterisks indicate significant differences on post-hoc tests (P < .005). D. Neuron number in the striatum. There are no significant difference in striatal neuronal number among the five inbred strains (F4,28 = 2.0. ns).
Striatal Neuron-Packing Density. There is a significant difference among strains in the packing density of striatal neurons (P < .001 for all comparisons). A/J has a higher mean density (84,800 ± 3,500 neurons/mm3) than all strains other than DBA/2J (80,400 ± 2,700 neurons/mm3). BALB/cJ (57,700 ± 2,500 neurons/mm3) and BXD5 (62,700 ± 2,600 neurons/mm3) do not differ significantly from each other, but do differ from all other strains. C57BL/6J (73,100 ± 1,700) differs from all other mice with the exception of DBA/2J. Inbred strains with smaller striatal volumes have higher neuronal packing densities (Fig 1C).
Striatal Neuron Number. As a result of the reciprocal relation between volume and density, there is no significant difference in striatal neuron number among the five strains. Total striatal neuron numbers ranges over a very modest range - from a low of 1.72 ± .015 million in C57BL/6J to a high of 1.93 ± .035 million in BXD5.
Correlational Statistics
Our comparisons are based on five strains, and one consequence of this modest sample size is that sampling errors and intraclass correlations may bias the results [31]. We therefore also analyzed a larger sample of genetically heterogeneous ABF2 intercross animals (an F2 intercross between A/J and BXD5). We first determined that the distribution of all four dependent measures were normally distributed (Fig 2), before subjecting the data to correlational analysis. In this set of animals brain weight correlates significantly with striatal volume (r = .82, df = 42, P < .001, Fig 3A). Striatal volume is negatively correlated with neuron-packing density overall (r = -0.32, df = 42, P < .05), again indicating that the greater the striatal volume, the lower the neuron-packing density (Fig 3B). Despite this relationship, the total population of striatal neurons correlates positively with striatal volume in this larger sample (r = .60, df = 42, P < .001; Fig 3C). In this crucial respect, results from the genetically heterogeneous F2 animals differ from those of inbred strains.
Figure 2
Histograms of distribution of dependent measures in ABDF2 subjects. Brain weight (A: Χ2 = 2.91, df = 2, ns), striatal volume (B: Χ2 = 1.13, df = 2, ns), striatal neuron-packing density C: Χ2 = 1.64, df = 2, ns), and striatal neuron number (D: Χ2 = 0.73, df = 2, ns) are all normally distributed (Kolmogorov-Smirnov Normality Test).
Figure 3
Scatterplots of subjects from an F2 intercross between a BXD5 and A/J inbred strains (ABF2, N = 44) illustrating correlations of striatal volume with brains weight (A), striatal neuron-packing density (B), and striatal neuron number (C). There are significant positive correlations between striatal volume and both brain weight and neuron number. Striatal volume negatively correlates with neuron-packing density in these subjects. ***P < .001; *P < .05.
QTL Analysis
The analysis in this section is focused principally on two traits: striatal volume (absolute and residual values), and striatal neuron number (also absolute and residual values). Residuals for these traits were computed for both traits to minimize the influence of brain weight. Two additional traits were mapped to assess specificity of the effects of putative QTL-bearing intervals, namely brain weight and non-striatal brain weight (Table 1). This last parameter was estimated by subtracting the estimated bilateral striatal weight (assuming a specific gravity of 1.0) from that of the whole brain.
Table 1
Linkage Statistics for Striatal Volume and Neuron Number
** Alleles inherited from BXD5 that increase a value are defined as positive additive effects. † LRS values can be converted to LOD scores by dividing by 4.6. Previously described QTL for brain weight [56]. Column headings:Trait, the phenotype used in linkage analysis; Marker, the symbol of the microsatellite loci used to genotype mice; Chr, the chromosome on which the marker is located; LRS is the likelihood ratio statistic (4.6 x the LOD score); %Var is the percentage of the total phenotypic variance apparently accounted for by differences in genotype in the an interval defined by the marker; P, the point-wise probability that the linkage is a false positive. Add and Dom are estimates of the additive and dominance effects of genetic variation. Units are the same as those of the traits (volume in mm3 or numbers of cells). The two bold loci marked with asterisks achieve genome-wide significance in this sample population.
The strongest linkage was found between variation in striatal volume and an interval on chromosome 10 between the markers D10Mit194 at 29 cM and D10Mit209 at 49 cM. The likelihood ratio statistic - a value that can be read as a chi-square - peaks in this 20 cM interval at 17.5 and is associated with a P value of 0.00016 (Table 1), equivalent to a LOD score of 3.8 (Fig 4A). The genome-wide probability of achieving a linkage of this strength by chance is <0.05. This locus accounts for as much as a third of the total phenotypic variance in striatal volume and as much as 50% of the genotypic variance (h2 = 0.39). Alleles in this Chr 10 interval that are inherited from the BXD5 parental strain contribute to a significantly larger striatum than do alleles inherited from A/J. Mean bilateral volume corrected for shrinkage for the AA genotype at D10Mit186 (n = 11) is 25.3 ± 1.3 mm3 whereas that for AB and BB genotypes are 29.1 ± 0.7 mm3 and 30.0 ± 0.5 mm3 (n = 14 and 11), respectively. The insignificant difference between BB and AB genotypes suggests that the B allele is dominant. BXD5 is a recombinant inbred strain initially generated by crossing strains C57BL/6J and DBA/2J. Most of the proximal part of Chr 10 in BXD5 is derived from DBA/2J, but a short interval between 40 and 50 cM is derived exclusively from C57BL/6J. This C57BL/6J region corresponds to the peak LRS score. We estimate a single B allele inherited from the BXD5 parent increases striatal volume by approximately 2.0-2.5 mm3.
Figure 4
Plots of LOD scores and LRS for each of the two traits. A. Plot for Striatal volume on Chr. 10. Peak values for the LRS are around 30 cM. B. Plot for striatal neuron number on Chr. 19. Peak values for the LRS are around 50 cM.
The Chr 10 interval has an appreciable effect on brain size. Variance in brain weight minus that of the striatum is associated with an LRS of 14.2 in the same location between D10Mit106 and D10Mit186. Each B allele adds 20-30 mg to total brain weight. The Chr 10 locus clearly has pleiotropic effects on the CNS, but its effect on the striatum is more intense. Nonetheless, until we know more about the scope of effects, we have opted to give this Chr 10 locus a generic name, Brain size control 10a (Bsc10a). The specific striatal component of the Bsc10a was analyzed by mapping the residual striatal values that corrects for differences in brain volume. The specific effect of a B allele is reduced from 2.5 mm3 to 0.5-1.0 mm3, and the LRS is reduced to 6.9, a value which still has a point-wise probability of only 0.03, indicating a significant independent effect.
We identified a second strong candidate interval on distal Chr 19 that may modulate striatal neuron number. The LRS peaks at 15.0 (a LOD of 3.26) at one of our more distal markers (D19Mit123, 51 cM, p = .00055). In mapping neuron number we actually used the residual cell population as a trait, and we are therefore confident that this interval has a selective, although not necessarily exclusive, effect on numbers of neurons in striatum (Fig 4B). Each B allele increases the population by approximately 200,000 cells. The AA genotype has an average residual population that is 116,000 less than the mean (i.e., a residual of -116,000; n = 12), the AB heterozygotes have an average of -8,000 neurons (n = 18), and the BB homozygotes have an average of 290,000 neurons (n = 6). Corresponding absolute numbers of striatal neurons for the three genotypes are 1.8, 1.9, and 2.2 million. The two-tailed genome-wide probability of this locus is at the threshold for declaring a QTL (p = 0.035 ± 0.01 two-tailed for an additive model and 0.08 ± 0.2 for a free model). No other chromosomal interval has an LRS score remotely as high as distal Chr 19. The next highest LRS value is only 7.2 on Chr 1 near D1Mit65 and has a point-wise probability that is 50 times higher than the Chr 19 interval. Given these findings we have given the distal chromosome 19 interval the name Striatal Neuron Number 19a (Stnn19a). Allelic differences in this interval account for up to 30% of the total variance in striatal neuron number. As the heritability of this trait is 0.64, this trait can be said to account for over 80% of the genetic variance. Residual neuron counts have a higher LRS than the total neuron counts (LRS of 15.0 vs. 11.9). This indicates that the Chr 19 interval is likely to have selective effect on the striatum. Consistent with this hypothesis, the LRS for brain weight on distal Chr 19 is under 1.0, and weights of all three genotypes average 480 ± 5 mg. Linkage on Chr 19 is not affected at all by remapping with control for the striatal volume locus on Chr 10. Thus, Chr 10 and Chr 19 intervals do not interact or cooperate in controlling striatal volume or neuron number.
Discussion
Striatal volume correlates strongly to brain weight. Nonetheless, a significant fraction of the variation in both striatal volume and neuron number among inbred strains of mice can not be predicted on the basis of brain weight or volume. This non-predictable variation is of particular interest to us because it is generated in large part by genes that have more intense or even selective effects on the dorsal striatum than other brain regions. We have succeeded in mapping one QTL with somewhat more intense effects on the volume of the striatum than the rest of the brain to the proximal half of chromosome 10. We have also mapped a QTL with selective effects on number of neurons in the striatum to the distal end of chromosome 19.
Between-strain variability
Variation in the size of CNS regions and cell populations is already known to be substantial in the striatum and in many other regions of the CNS. The number of striatal cholinergic neurons, for example, varies 50% among 26 BXD RI lines [32]. Interestingly, this variation appears to be unrelated to susceptibility to haloperidol-induced catalepsy. The volume of the granule cell layer of the dentate gyrus varies as much as 40-80% among different inbred strains of mice [33-35]. More recent experiments using stereologic techniques have reported substantial variation in both neuron number and volume of the pyramidal and dentate cell layers of the hippocampus [36]. Granule cell numbers in NZB/BINJ and DBA/2 were 37-118% greater than C57BL/6J mice, and differences in volume were even larger (up to 150% larger in the DBA/2 as compared to the C57BL/6J mice). There is also substantial among-strain variation in other structures in the nervous system including the nucleus of the solitary tract [37], the spinal nucleus of the bulbocavernosus [38], and retinal ganglion cells [39, 40]. Taken together, these results point to a high level of variability in neuron number in the CNS of mice.
Based on these findings, we anticipated significant differences in the striatum of inbred strains. We did find large strain differences in volume. What was surprising was that in our set of five highly divergent strains the differences in volume were not matched by significant differences in neuron number. There was in fact a strong inverse relationship between striatal volume and neuron-packing density that led to a remarkably stabile neuron number. The variation in neuron-packing density with volume contrasts somewhat with the report of Abusaad and colleagues [36], who found no significant differences (P = .06) in neuronal packing density in the dentate gyrus cell layers of the hippocampus among the three mouse strains that they examined, but did see a significant difference in the pyramidal cell layers. A recent report demonstrates a 25% range of granule cell packing density among 35 BXD recombinant inbred lines [41], a finding supporting the notion that packing density varies significantly among mouse strains.
These data suggest that principles that govern the relationship between neuronal volume and neuron-packing density may differ between the striatum and the other CNS regions. The striatum may be a special case - a region in which numbers of medium spiny neurons is more tightly regulated than neuron populations in some other regions. It could also be that measuring specific neuronal subtypes would demonstrate a greater amount of variance than we currently report [32]. Given the relatively small number of strains that we have sampled, our hypothesis of lower variation in striatal cell populations requires a more extensive test, a problem which we are now pursuing using the large numbers of strains in the Mouse Brain Library .
Verification of QTL Results
We have quantified the population of striatal neurons on both sides of the brain in 77 cases total. This is a large sample from the perspective of stereological analysis of the mouse CNS, but from the perspective of gene mapping and quantitative genetics this is, of course, a modest-sized sample size and one that will need to be treated as a starting point for more refined genetic analysis. Nonetheless, we have succeeded in mapping one locus, Bsc10a, which modulates striatal volume with a genome-wide significance of P < 0.05. We have also discovered linkage on Chr 19 to variation in the total number of striatal neurons. These mapping data are concordant with our strain comparison and collectively suggest that there is apparently no significant genetic correlation between striatal volume and neuron number. To confirm and refine our genetic dissection of the striatum we plan to analyze the AXB and BXA recombinant inbred (RI) strains generated by crossing A/J with C57BL/6J. This large RI set has already been processed and regenotyped and is now part of the Mouse Brain Library (see and ). An analysis of RI strains can be quickly extended by generating F1 intercrosses between A/J and the subset of RI strains that have recombinations in the QTL intervals on Chrs 10 and 19. Isogenic sets of RI-backcross progeny can be used to test specific models of gene action, for example, the dominance of the B allele at Bsc10a.
A major goal of QTL mapping is to define loci that affect critical phenotypes with sufficient precision to generate short lists of candidate genes. Generating lists of candidates for QTLs will soon be greatly facilitated by more complete and better annotated mouse and human sequence databases combined with information on gene expression profiles of whole brain and striatum [42]. Once chromosomal positions of the QTLs have been determined to a precision of 1-3 cM, reducing the probability that a QTL actually represents a cluster of linked genes, it will become appropriate to assess strengths of candidates using transgenic animals and by sequence comparisons [43].
Interval mapping places the QTL for Bsc10a in the central portion of Chr 10 in proximity with a number of genes known to affect brain development. One of these is Grk2, a member of the family of ionotropic glutamate receptor genes that is thought to play a role in modulating Huntington disease [44]. In the mouse, members of this receptor type act to indirectly down-regulate synaptic activity in the striatum [45]. Another gene that falls into the Bsc10a interval is Macs, the gene encoding the myristoylated alanine-rich C kinase substrate (MARCKS protein). This molecule is important in cerebral development. MARCKS-deficient mice have a high incidence of exencephaly, agenesis of the corpus callosum, and abnormalities other forebrain structure including widespread neocortical ectopias [46, 47]. The MARCKS-related protein gene is expressed in the striatum during early brain development in the rat [48].
The location of the QTL modulating striatal neuron number to the distal part of chromosome 19 places it in proximity to a number of genes that have been recently been shown to be important factors in telencephalic development, particularly Vax1. Vax1 is a homeobox-containing gene and is a close relative of the Emx and Not genes. Vax1 is localized during development to the anterior ventral forebrain, and is expressed in the striatum during embryogenesis [28]. This molecule also has an important role in axon guidance: both the anterior portion of the corpus callosum and the optic chiasm are malformed or absent in Vax1 knockout mice [49]. In addition, Vax1 interacts with several molecules including sonic hedgehog,Pax2, Pax6, and Rx that are known to be important during development of the basal forebrain [27, 50].
Brain volume and neuron number
It has previously been shown that differences in brain weight are proportional to total brain DNA content and consequently to total CNS cell number [51, 52]. For this reason, brain weight has been suggested to be a good surrogate measure for total cell number in mice, as in humans [53]. Moreover, previous work has demonstrated a tight link between regional brain volume and neuron number [54, 55], which implies that volumetric measures reliably estimate neuron number. With this literature in mind, we expected that our measures of striatal volume would predict neuron number in this nucleus. With the inbred strains, however, we found that strains with small striata (A/J) had virtually the same number of neurons as those with large striata (BALB/cJ). This result indicates that at least for the striatum, volume is not a reliable indicator of neuron number, and that they may be two independent traits. This conclusion is bolstered at the genetic level by our report of two distinct QTLs for these two morphologic phenotypes. Taken together with previous reports [51-53], we speculate that while total neuron number in the cerebrum may relate to total brain weight, the relationship of these two variables is flexible at the regional level.
Materials and Methods
Subjects
Thirty-four of the 78 mice that we analyzed were common inbred strains that were selected to sample a wide range of brain weights, and by expectation, striatal volumes. Low brain weight strains included A/J (n = 5) and DBA/2J (n = 8). Mid and high brain weight strains include C57BL/6J (n = 10), BALB/cJ (n = 5), and BXD5 (n = 6, formally this recombinant inbred strain is known as BXD-5/Ty). One of the ten C57BL/6J subjects was removed from the analysis because values for striatal neuron number were anomalous with Z scores more than 2.5.
To map QTLs that modulate variation in CNS size and cell populations we used an F2 intercross between a strain with low brain weight (A/J) and a strain with high brain weight (BXD5). A total of 518 of these ABDF2 progeny were generated, but for this study we selected a subset of 44 cases, of which 36 were fully genotyped (see below). The sample included 20 animals in the lowest and highest quartiles, and 24 cases within 0.5 SD of the mean brain weight. We therefore measured subjects representing the full range of brain weights (see Fig 2A). The ABDF2 intercross has been used previously to map QTLs that modulate total brain weight [56] and cerebellar volume [57]. The BXD5 strain used as the paternal strain in this intercross is a recombinant inbred strain that was derived by crossing C57BL/6J and DBA/2J lines of mice [58]. As a result, ABDF2 progeny are a mixture of three genomes (50% A/J, 25% C57BL/6J, and 25% DBA/2J). However, at any given locus there will be only two alleles, A and B, or A and D. All stocks of mice were obtained from the Jackson Laboratory . ABF2 mice were generated at the University of Tennessee by Dr. Richelle Strom [56] using Jackson Laboratory foundation stock. The F2 mice ranged in age from 35 to 143 days. The standard inbred strains ranged in age from 51 to 365 days. We studied approximately equal numbers of males and females.
Histological Preparation
All brains analyzed in this study are part of the Mouse Brain Library (MBL). The MBL is both a physical and Internet resource. High-resolution digital images of sections from all cases are available at .
Mice were anesthetized deeply with Avertin (1.25% 2,2,2-tribromoethanol and 0.8% tert-pentyl alcohol in water, 0.5-1.0 ml ip) and perfused through the left ventricle with 0.9% sodium phosphate buffered (PB) saline (pH 7.4) followed by 1.25% glutaraldehyde/1.0% paraformaldehyde in 0.1 M PB (pH 7.40) over a period of 2 to 4 min. An additional 10-ml of double-strength fixative (2.5% glutaraldehyde/2.0% paraformaldehyde) was injected for 1 to 2 min at an increased flow rate. The head with brain was placed a vial with the last fixative and stored at 4°C until dissection.
Following dissection, the brains were weighed immediately. Brains were subsequently shipped to Beth Israel Deaconess Medical Center. They were immersed in fresh 10% formalin for at least one week before being embedding in celloidin [59]. Brains were cut on a sliding microtome at 30 μm in either horizontal or coronal planes. Free-floating sections were stained with cresylechtviolett and four series of every tenth section were mounted on slides and coverslipped (see for further details).
Histologic Phenotypes
Total Brain Volume
To accurately estimate histological shrinkage for each brain in the sample, we determined the volume of the entire brain and took a ratio of this value to the original fixed brain weight. Brain volumes were determined from serial sections using point counting and Cavalieri's rule. High-resolution (4.5 μm/pixel) images of entire sections were taken from the Mouse Brain Library, and point counting was performed on these images using NIH Image 1.55 and an Apple Macintosh computer . If the criteria for using the Cavalieri's estimator were not met (due to missing or damaged sections), a measurement method involving piecewise parabolic integration was employed [60]. Subsequent measurements of striatal volume and neuron packing density were corrected for volumetric shrinkage. The average shrinkage was 62.2 ± 0.4% (a mean residual volume of 37.8%).
Striatal Volume
Volume of the striatum was also determined from serial section analysis using point counting and Cavalieri's rule. Images from the sections were captured at 12.5 x and were projected onto a video monitor. Point counting was performed as above. Volume was computed separately for the right and left sides and corrected for shrinkage.
Striatal Neuron-Packing Density and Neuron Number
Neurons were counted using the 3-dimensional counting software of Williams and Rakic [24]. A series of six contiguous counting boxes (each 40 x 65 x 20 μm) aligned in a 3 x 2 matrix were placed randomly within the striatum, and those neurons the nucleoli of which were in focus were counted as described previously [61, 62]. This large functional counting box (80 x 195 x 20 μm) was chosen to minimize sampling variance by ensuring an equitable sampling of striatal patch and matrix. Two of these large fields were counted in each of the hemispheres. Neuron-packing density was computed as the number of cells/mm3 corrected for shrinkage. Multiplying the volume of the striatum by its cell-packing density permitted estimation of the number of neurons in that nucleus.
Reliability
We determined test-retest reliability by having an observer blindly re-measure striatal volume on a subset of 10 brains from the collection. The observer not only re-measured the striatal volume from the same series of sections as the original measure, but also estimated volume from a second series of 1 in 10 sections offset by 5 sections from the previous series. The correlations among the three estimations ranged from .95 to .99 (P < .05), indicating a high degree of reliability for this dependent variable.
We assessed reliability of our estimates of neuronal numbers by having an observer blindly re-estimate neuron number in the same 10 brains above. The intra-observer correlation for this measure was .81 (P < .05), which is similar to the reliability seen in previous estimates of neuron number [39, 40].
Genotyping and QTL Mapping
Genomic DNA was extracted from spleens of F2 animals using a high-salt procedure [63]. A set of 82 microsatellite loci distributed across all autosomes and the X chromosome were typed in a set of ABDF2 animals using a standard PCR protocol [64, 65] as detailed in Zhou and Williams [66]. F2 genotypes were entered into a spreadsheet program and transferred to Map Manager QTb28 for mapping and permutation analysis [67]. Map Manager implements both simple and composite interval mapping methods described by Haley and Knott [68]. Two-tailed genome-wide significance levels were estimated by comparing the highest likelihood ratio statistic (LRS) of correctly ordered data sets with LRSs computed for 10,000 permutations of those same data sets [69]. LRS scores can be converted to LOD scores by dividing by 4.6. The 2-LOD support interval of linkage was estimated directly from interval maps. The approximate 95% support interval was estimated by application of equations in Darvasi and Soller [70]. With a modest sample size such as we have been able to examine using unbiased stereological methods, even a QTL responsible for 30 to 50% of the variance. is associated with a 95% interval of 20 to 30 cM.
Regression Analysis of Trait Values
The unadjusted striatal estimates vary to a large extent as a result of variation in total brain weight. However, one of our goals in this study is to map QTLs with relatively intense effects on the striatum. For this reason we also have corrected all of the parameters used in the mapping analysis for variation in brain weight using linear regression analysis. We have mapped data with and without compensation for variance in brain weight. The corrected values are referred to as residuals.
Analysis
All data were analyzed using regression, correlation, and ANOVA statistical tests (see StrAnatData.xls for original data used to perform this analysis). A Bonferroni/Dunn correction was used for post hoc examination of significant main effects in the ANOVA. This post-hoc test is functionally identical to a Fisher PLSD, but the alpha level is more conservative (.005).
Supplementary Material
StrAnatData.xls
This file contains anatomic data for each of the subjects used in the current experiment.
Click here to download StrAnatData.xls
StrMap.qtx
This file contains the genotyping data from the 82 markers used in the current experiment, in MapManager QTX format.
Click here to download StrMap.qtx
Acknowledgements
This work was supported, in part, by grants HD20806 and NS35485 from the Public Health Service of the USA. The authors wish to thank Dr. Jing Gu, Aaron Levine, Anna Ohlis, and Stefany Palmieri for technical assistance. We thank Richelle Strom for generating the F2 intercross mice. | [
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11604102 | Characterization of the mouse Dazap1 gene encoding an RNA-binding protein that interacts with infertility factors DAZ and DAZL
Abstract
Background
DAZAP1 (DAZ Associated Protein 1) was originally identified by a yeast two-hybrid system through its interaction with a putative male infertility factor, DAZ (Deleted in Azoospermia). In vitro, DAZAP1 interacts with both the Y chromosome-encoded DAZ and an autosome-encoded DAZ-like protein, DAZL. DAZAP1 contains two RNA-binding domains (RBDs) and a proline-rich C-terminal portion, and is expressed most abundantly in the testis. To understand the biological function of DAZAP1 and the significance of its interaction with DAZ and DAZL, we isolated and characterized the mouse Dazap1 gene, and studied its expression and the subcellular localization of its protein product.
Results
The human and mouse genes have similar genomic structures and map to syntenic chromosomal regions. The mouse and human DAZAP1 proteins share 98% identity and their sequences are highly similar to the Xenopus orthologue Prrp, especially in the RBDs. Dazap1 is expressed throughout testis development. Western blot detects a single 45 kD DAZAP1 protein that is most abundant in the testis. Although a majority of DAZAP1 is present in the cytoplasmic fraction, they are not associated with polyribosomes.
Conclusions
DAZAP1 is evolutionarily highly conserved. Its predominant expression in testes suggests a role in spermatogenesis. Its subcellular localization indicates that it is not directly involved in mRNA translation.
Background
Spermatogenesis is a complex developmental process in which male germ cells progress through mitotic proliferation, meiotic division and dramatic morphological changes to form mature sperm. This process is vital for the propagation of a species, and involves a large portion of the genome of an organism to ensure the quality and quantity of the final products. It is estimated that mutations in up to 11% of all genes in Drosophila might lead to male sterility [1]. This is likely to be true for humans also, considering the extremely high incidence (4–5%) of infertility in men [2]. Among the genes associated with male infertility is the DAZ (Deleted in Azoospermia) gene family. The family includes the Y-linked DAZ genes that are present only in great apes and old world monkeys [3], and the autosomal DAZL1 (DAZ-like 1) and BOULE genes [4,5] in all mammals. Deletion of the DAZ genes is found in about 10% of infertile males with idiopathic azoospermia [2], and disruption of Dazl1 causes infertility in both male and female mice [6]. Mutations in the DAZ family members of Drosophila[7], C. elegans[8], and Xenopus[9] also affect the fertility in either males, females, or both sexes.
The DAZ gene family encodes RNA binding proteins that are expressed specifically in germ cells. DAZ and DAZL are expressed in the nucleus and cytoplasm of primordial germ cells and spermatogonia, and in the cytoplasm of meiotic spermatocytes [6,10]. BOULE is expressed later, in the cytoplasm of pachytene spermatocytes [5]. Genetic and biochemical studies suggest a role for the DAZ family in the regulation of mRNA translation. Drosophila Boule mutants was defective in the translation of the meiosis-specific CDC25 homologue, Twine [11], and DAZL was found to be associated with polyribosomes in mouse testes [12]. More recently, DAZL was shown both in vitro and in a yeast three-hybrid system to bind specifically to oligo(U) stretches interspersed by G or C residues, including a U-rich segment in the 5' UTR of mouse Cdc25C mRNA [13].
In an attempt to elucidate the function of the DAZ gene family and to understanding the mechanisms of its action, we used a yeast two-hybrid system to isolate two human genes encoding DAZ associated proteins (DAZAPs) [14]. One of them, DAZAP1, is expressed predominantly in testes. It encodes a protein with two RNA binding domains and a proline rich C-terminal portion. The DAZAP1 protein interacted with both DAZ and DAZL in vitro. It also bound to RNA homopolymers. We now report our characterization of the mouse Dazap1 gene and its protein product. The subcellular localization of DAZAP1 suggests that it is not involved directly in mRNA translation.
Results
Characterization of the mouse Dazap1 cDNA
Mouse Dazap1 cDNA clones were isolated by library screening, and the 5' end of the cDNA was isolated by 5' RACE [15]. The near fall length cDNA consists of a 53 bp 5' untranslated region (UTR), an open reading frame for a protein of 405 amino acid residues, and a 362 bp 3' UTR (GenBank Accession No: AF225910). The coding region shares 89% similarity with that of the human orthologue. The 3' UTR sequence is remarkably conserved. It contains three segments of 35 bp, 133 bp and 90 bp that share 85%, 90%, and 97% similarity with segments in the human 3' UTR, respectively. These segments probably contain regulatory elements.
The DAZAP1 protein contains two RNA-binding domains (RBDS) and a C-terminal portion that is rich in proline (Figure 1). It is highly conserved evolutionarily. The mouse and the human proteins differ in 9 amino acids only, with 7 substitutions and two deletions/insertions. The mammalian proteins shares 89% similarity and 81% identity with Xenopus Prrp (for proline-rich RNA binding protein) [16]. The two RBDs are especially highly conserved. They share 98% and 97% similarity and 97% and 92% identity, respectively, between DAZAP1 and Prrp. These proteins may therefore have a similar RNA binding specificity. The C-terminal proline-rich portions of DAZAP1 and Prrp are less conserved (81% similarity and 71% identity). There is an insertion of a 58 bp segment in Prrp cDNA that causes a change of reading frame and results in a shorter Prrp with a different C-terminal end sequence.
Figure 1
Evolutionary conservation of the DAZAP1 proteins. The amino acid sequences of the human and mouse DAZAP1s and the Xenopus Prrp are compared. The two RNA binding domains are boxed. Differences between the human and the mouse sequences, and between the mouse and Xenopus sequences are marked by #'s and *'s, respectively.
Genomic structure of Dazap1 and chromosomal mapping
Several overlapping lambda clones containing mouse Dazap1 genomic sequences were isolated. The locations of exons were determined by PCR amplification across exon-intron boundaries following by sequencing. All but the first exon were isolated and mapped. The genomic structure of the human DAZAP1 gene was also determined by blasting the human genome sequence at National Center for Biotechnology Information with the human DAZAP1 cDNA sequence. The mouse and the human genes have very similar structures, consisting of 12 exons spanning about 28 kb. All intron insertion sites are conserved (Table 1). The two RBDs are encoded by exons 1–4 and 5–8, respectively.
Table 1
Exon-intron Organization of the Mouse and Human DAZAP1 genes
a: Introns are inserted after the indicated nucleotide positions of DAZAP1 cDNA sequences. GenBank accession numbers for mouse and human cDNAs are AF225910 and AF181719, respectively.
A pair of PCR primers was designed from Dazap1 intronic sequences that amplified mouse but not hamster genomic sequences. Using a panel of mouse-hamster radiation hybrids, the mouse Dazap1 gene was mapped to chromosome 10 placed 27.84 cR from D10Mit260 (lod > 3.0) (data not shown). This region is syntenic to human 19p13.3 where the human DAZAP1 gene is located [14,17]. It contains no known mutant alleles that are associated with infertility.
Expression of Dazap1
Northern analyses of adult mouse tissues showed the presence of two Dazap1 transcripts of 1.75 kb and 2.4 kb, respectively (Figure 2). Only the shorter transcript has been isolated in cDNA clones. Dazap1 was expressed most abundantly in the testis, much less in liver, heart and brain, and even less in other tissues. This pattern of expression is similar to that of the human DAZAP1 (14). RT-PCR analyses showed that Dazap1 mRNA was already present in fetal testes at embryonic day 15, similar to Dazl1 mRNA (Figure 3). The expression of both Dazl1 and Dazap1 persisted throughout testes development, in both the prenatal and postnatal periods. Dazl1 and Dazap1 transcripts were also present in the testes of Wv/Wv mutant mice which contained diminished number of germ cells [18]. However, only Dazap1 was expressed in a mouse germ cell line GCl-spg [19] and a Sertoli cell line MT4. The results suggest that Dazap1 is expressed in both somatic and germ cells in the testis.
Figure 2
Expression of Dazap1 in adult mouse tissues. A mouse multiple-tissue Northern blot was hybridized with a Dazap1 cDNA probe, stripped, and rehybridized with a β-actin probe. Dazap1 is expressed most abundantly in the testis.
Figure 3
Developmental expression of Dazap1 and Dazl in mouse testes. RT-PCR was performed on total testicular RNAs isolated from day 15 (El 5) and day 17 (El 7) embryos, new born mice (Day 0), and mice at various days after birth. Wv/Wv testes contain diminished germ cell population due to a mutated W (White spotted) gene. GC1 and MT4 are mouse germ cell and Sertoli cell lines, respectively, and gDNA is mouse genomic DNA. The PCR primers span over introns and produce much larger (if any) fragments from genomic DNA.
To study the expression of the DAZAP1 protein, two antibodies against mouse DAZAP1 were generated. The anti DAZAP1-C antibody was raised against a recombinant protein containing the C-terminal proline-rich portion, and the anti DAZAP1-P antibody was raised against an oligopeptide containing the last 19 amino acid residue at the C-terminus. Both antibodies recognized in vitro synthesized DAZAP1 in an immunoprecipitation assay (data not shown). Western blotting of mouse tissue extracts detected a 45 kD protein that was present most abundantly in the testis, and to a lesser degree in spleen, liver, lung and brain (Figure 4). The protein was also present in the ovary. The expression of DAZAP1 during germ cell development paralleled that of DAZL (Figure 5). It is present at a low level in the testes of 6 days old mice which contained only primitive type A spermatogonia. The expression of DAZAP1 increased afterward, as the testes contained increasing number of proliferating and meiotic germ cells.
Figure 4
Expression of the DAZAP1 protein in adult mouse tissues. Equal amounts of total protein from various tissue extracts were applied to a 10% SDS-polyacrylamide gel and western blotted with the anti-DAZAP1-P antibody.
Figure 5
Western blot analyses of the expression of DAZAP1 and DAZL in mouse testes during postnatal development.
Subcellular localization of DAZAP1
Our previous fractionation of mouse testis extracts showed that most DAZL were present in the post mitochondrial fraction, and some of them were associated with polyribosomes [12]. Similar analyses showed that a majority of DAZAP1 in adult mouse testes was also present in the cytoplasmic fraction (data not shown). However, sucrose gradient analyses of the post- mitochondria fraction showed that, unlike DAZL, DAZAP1 did not co-sediment with polyribosomes (Figure 6).
Figure 6
Sucrose gradient analyses shows that DAZAP1 is not associated with polyribosomes. The post-mitochondrial supernatant of mouse testis extracts was analyzed on a 15–45% sucrose gradient. Sedimentation was from left to right. The presence of DAZAP1 and DAZL in each fractions was analyzed by Western blotting.
Discussion
RNA-binding proteins have been found to participate in many cellular functions, including RNA transcription, pre-mRNA processing, mRNA transport, localization, translation and stability [20]. A role for the DAZ family in the regulation of mRNA translation is supported by lines of circumstantial evidence, including the association of DAZL with polyribosomes [12]. The absence of DAZAP1 from polyribosomes indicates that it is not directly involved in protein synthesis. This finding is different from two RNA-binding proteins, FXR1P and FXR2P, that were identified through their interaction with another polysomal-associated RNA-binding protein, the fragile X mental retardation protein [21]. Both FXR1P and FXR2P are associated with the polyribosomes [22].
The significance of the interaction between DAZAP1 and DAZL/DAZ remains to be defined. These proteins may act together to facilitate the expression of a set of genes in germ cells. For example, DAZAP1 could be involved in the transport of the mRNAs of the target genes of DAZL. Alternatively, DAZL and DAZAP1 may act antagonistically to regulate the timing and the level of expression. Such an antagonistic interaction between two interacting RNA-binding proteins is exemplified by the neuron-specific nuclear RNA-binding protein, Nova-1. Nova-1 regulates the alternative splicing of the pre-mRNAs encoding neuronal inhibitory glycine receptor α2 (GlyR α2) [23]. The ability of Nova-1 to activate exon selection in neurons is antagonized by a second RNA-binding protein, brPTB (brain-enriched polypyrimidine tract-binding protein), which interacts with Nova-1 and inhibits its function [24]. DAZAP1 could function in a similar manner by binding to DAZL and inhibiting its function. Comparing the phenotypes of Dazl1 and Dazap1 single and double knock-out mice may provide some clues to the significance of their interaction. Dazl1 knock-out mice have already been generated and studied [6]. The spermatogenic defect in the male becomes apparent only after day 7 post partum when the germ cells are committing to meiosis (H. Cooke, personal communication). The genomic structure of Dazap1, delineated here, should facilitate the generating of Dazap1 null mutation.
DAZAP1 was shown to bind RNA homopolymers in vitro, with a preference for poly U and poly G. Its natural substrates have not been identified. Recently, the Xenopus orthologue of DAZAP1, Prrp, was identified and characterized [16]. Prrp binds to a 340 nt sequence in the 3' UTR of Xenopus Vg1 mRNA. This Vg1 localization element (VLE) is sufficient for the migration and clustering of Vg1 mRNA to the vegetal cortex of mature oocyte. Prrp also interacts through its proline-rich domain with two microfilament-associated proteins profilin and Mena, which may facilitate the anchoring of Vg1 mRNA to the vegetal cortex. The Vg1 RNA encodes a member of the transforming growth factor-β family that is required for generating dorsal mesoderm at the blastula stage of Xenopus embryogenesis [25]. Sequence conservation between the RBDs of DAZAP1 and Prrp suggests that these proteins may bind to similar RNA sequences. However, a BLAST search of the GenBank for the 340 nt VLE sequence failed to identify any mammalian sequences with significant homology. Further mapping of the RNA sequence within VLE that binds Prrp, and possibly DAZAP1, may help to identify the natural substrates of DAZAP1.
Conclusions
DAZAP1 is an evolutionarily conserved RNA-binding protein. It is present at variable levels in many tissues. Its predominant expression in testes suggests a role in spermatogenesis. In mouse testes, DAZAP1 was found both in the nuclei and in the cytoplasm. Its absence from polyribosomes indicates that it is not directly involved in mRNA translation.
Materials and methods
Isolation of mouse Dazap1 cDNA clones
Dazap1 cDNA clones were isolated from a mouse testis cDNA library (#937308, Stratagene, La Jolla, CA) using a human DAZAP1 cDNA as a probe. The 5' end of the cDNA was isolated by 5' RACE [15], using prdap11 (ttgcgggccatatccttg, #749–732) as the primer for cDNA synthesis from mouse testis RNA, and prdap37 (ttgttgccacgtgggcg, #734–718) and an adaptor primer as the primers for PCR amplification. The PCR products were cloned into a TA cloning vector pCR2.1-TOPO (Invitrogen, Carlsbad CA). Dazap1 clones were identified by colony hybridization and sequenced. The 5' RACE clone with the longest 5' UTR region and the cDNA clone P21 were ligated together through a unique PmlI site at # 722 to generate a cDNA clone (Dazap1-C) with a near full-length insert.
Chromosomal mapping of Dazap1
Dazap1 genomic clones were isolated from a mouse 129SV genomic library (#946305, Stragagene), and sequences flanking each exons were determined. PCR primers (prdap25: cacctccaggatgtgttagc and prdazp26:gtcaccaagggtgtctgaag) were designed from intronic sequences flanking Dazap1 exon 8. These primers amplified a 271 bp fragment from mouse but not hamster genomic DNA. DNA samples of a panel of 100 radiation hybrids containing mouse chromosome fragments in a hamster background were purchased from Research Genetics (Huntsville, AL). The presence of mouse Dazap1 in the radiation hybrids was determined by PCR and the results were sent to the MIT server for computerized physical mapping of the gene.
Expression of Dazap1 transcripts
Northern hybridization was carried out according to standard procedures [26] using a mouse Multiple Tissue Northern Blot #7762–1 from Clontech (Palo Alto, CA). The blot was hybridized sequentially with DAZAP1 and β-actin cDNA probes, with stripping of the bound probes in between.
Reverse transcription-polymerase chain reaction (RT-PCR) was carried out as preciously described using an annealing temperature of 54°C [27]. The primers were prdap35: agctcagggagtacttcaaga and prdap24 :ggagcttgattcttgctgtcc for Dazap1 which generated a product of 211 bp, and prdaz71: atcgaactggtgtgtcgaagg and prdaz72: ggaggctgcatgtaagtctca for Dazl1 which generated a product of 245 bp. Both primer pairs annealed across intron insertion sites.
Generation of anti-DAZAP1 antibodies
Antibodies were generated against both a recombinant protein produced in E. coli and an oligopeptide synthesized in vitro. The insert of a Dazap1 cDNA clone P21, which encoded the C-terminal portion of DAZAP1 (starting from aa #197), was cloned in-frame into the EcoRI/XhoI sites of an expression vector pET32b (Novagen, Madison, WI). Sequences at the junctions were verified by DNA sequencing. Milligrams of fusion proteins between thioredoxin and DAZAP1 were prepared and purified on His-Bind metal chelation resins (Novagen, Madison, WI). The proteins were mixed with Freund's adjuvant and injected into rabbits to generate the anti-DAZAPl-C antibody. An oligopeptide containing the last 19 ammo acid residues of the mouse DAZAP1 was synthesized in vitro using the services of Bethyl Laboratories (Montgomery, TX). The peptide was conjugated to KLH as carrier and injected into a goat. The anti-DAZAP1-P antibody thus produced was purified on an affinity column containing the oligopeptide antigen.
Western blotting
Mouse tissues were homogenized in the RIPA lysis buffer (150 mM NaCl, 1.0% NP-40, 0.5% DOC, 0.1% SDS, 50 mM Tris, pH 8.0) at a concentration of 0.2 g tissue per ml of buffer. The homogenized samples were cleared of debris by centrifugation at 10,000 × g for 10 minutes. Protein concentration of the tissue extracts was determined by the Bradford method using the Bio-Rad Protein Assay system (Bio-Rad, Hercules, CA). About 50 μg of tissue extracts were separated on 10% SDS-polyacrylamide gels and blotted with either the anti-DAZAP1-C antibody (at a 1/2,000 dilution) or the anti-DAZAP1-P antibody (at a 1/5,000 dilution). After incubation with horseradish peroxidase-conjugated secondary antibodies, the binding of antibodies was detected using the ECL Western Blotting System (Amersham Pharmacia Biotech, Piscataway, NJ).
Fractionation of mouse testicular extracts
Adult mouse testes were homogenized in a buffer containing 20 mM Tris, pH 7.5, 100 mM KCl, 5 mM MgCl2, 0.3% NP-40, 40 U/ml of Rnasin ribonuclase inhibitor (Promega, Madison, WI), and a mixture of 10 protease inhibitors provided in the Protease Inhibitors Set (Roche Molecular Biochemicals, Indianapolis, IN). Homogenates were centrifuged at 1,000 × g for 10 minutes to pellet cell debris and nuclei. After an additional centrifugation at 10,000 × g for 10 minutes to pellet the mitochondria, aliquots of the supernatant were applied to 15–45% sucrose gradients in 20 mM Tris, 100 mM KCl and 5 mM MgCl2 and centrifuged in a Beckman SW41 rotor at 39,000 rpm for 2 hours at 4°C. Fractions of 0.5 ml were collected from the bottom of the tubes and analyzed by western blotting.
Acknowledgments
We thank Gary Kuo for his involvement in the construction of Dazap1 expression vectors, and Ron Swerdloff's group for helpful discussion. The work was supported by grants from the National Institutes of Health (HD28009 and HD36347). Y. Vera was supported by an NIH grant (GM56902) on Initiative for Minority Student Development. | [
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14624252 | Additive Effects of PDGF Receptor β Signaling Pathways in Vascular Smooth Muscle Cell Development
Abstract
The platelet-derived growth factor β receptor (PDGFRβ) is known to activate many molecules involved in signal transduction and has been a paradigm for receptor tyrosine kinase signaling for many years. We have sought to determine the role of individual signaling components downstream of this receptor in vivo by analyzing an allelic series of tyrosine–phenylalanine mutations that prevent binding of specific signal transduction components. Here we show that the incidence of vascular smooth muscle cells/pericytes (v/p), a PDGFRβ-dependent cell type, can be correlated to the amount of receptor expressed and the number of activated signal transduction pathways. A decrease in either receptor expression levels or disruption of multiple downstream signaling pathways lead to a significant reduction in v/p. Conversely, loss of RasGAP binding leads to an increase in this same cell population, implicating a potential role for this effector in attenuating the PDGFRβ signal. The combined in vivo and biochemical data suggest that the summation of pathways associated with the PDGFRβ signal transduction determines the expansion of developing v/p cells.
Introduction
Although signal transduction by receptor tyrosine kinases (RTKs) has been studied extensively, the roles of individual signaling proteins downstream of these receptors are a matter of debate. Some studies have shown that disruption of particular pathways leads to loss of specific cellular functions (Valius and Kazlauskas 1993). Others have suggested that it is the sum of the signals that results in the unique cellular outcomes directed by each receptor (Fambrough et al. 1999). Yet others have demonstrated that the interpretation of receptor signals is determined by the distinct cellular history (Flores et al. 2000; Halfon et al. 2000; Xu et al. 2000). Because many of these conclusions have been reached in diverse cell types and through the analysis of different RTKs, it is difficult to determine whether results from one receptor system can be used to generalize the functions of RTK signaling.
Recently, several labs have dissected the roles of RTK modular signaling components by generating point mutations in cytoplasmic domains of the receptors in mice (Partanen et al. 1998; Heuchel et al. 1999; Blume-Jensen et al. 2000; Kissel et al. 2000; Tallquist et al. 2000; Klinghoffer et al. 2001, 2002; Maina et al. 2001). These studies have revealed a unique requirement for individual signaling components in specific cell types (Partanen et al. 1998; Blume-Jensen et al. 2000; Kissel et al. 2000; Maina et al. 2001). In contrast, similar experiments on platelet-derived growth factor receptor α (PDGFRα) signaling mutants have demonstrated that phosphatidylinositol 3′-kinase (PI3K) and Src family kinase (SFK) signal transduction pathways play roles in oligodendrocyte development (Klinghoffer et al. 2002). These experiments suggest that requirements for signal transduction vary not only by the receptor under consideration, but also by the cell lineage that is receiving the signal.
The platelet-derived growth factor receptor β (PDGFRβ) has not only been studied physiologically, but also has been the focus of intensive biochemical analysis. Upon ligand binding, the PDGFRβ dimerizes and is autophosphorylated on as many as 13 cytoplasmic tyrosine residues. These phosphorylated tyrosines become binding sites for SH2 domain-containing proteins that initiate a number of signal transduction pathways (reviewed by Claesson-Welsh 1994; Heldin et al. 1998). The pathways downstream of the PDGFRβ control multiple cellular functions, including proliferation, migration, matrix deposition, and immediate early gene induction (reviewed by Heldin and Westermark 1999; Betsholtz et al. 2001). At least ten distinct SH2 domain-containing proteins can bind the phosphorylated PDGFRβ and activate downstream signal transduction cascades. These molecules include SFK (Kypta et al. 1990), PI3K (Kazlauskas and Cooper 1990; Kundra et al. 1994; Wennstrom et al. 1994a, 1994b), Shc (Yokote et al. 1994), RasGAP (Kaplan et al. 1990; Kazlauskas et al. 1990), signal transducers and activators of transcription (STATs) (Vignais et al. 1996), Grb2 (Arvidsson et al. 1994), Grb7 (Yokote et al. 1996), SH2-containing phosphotyrosine phosphatase (SHP-2, also known as SH-PTP2) (Kazlauskas et al. 1993; Lechleider et al. 1993), phospholipase Cγ (PLCγ) (Meisenhelder et al. 1989; Morrison et al. 1990), and Nck (Nishimura et al. 1993). While multiple downstream effects have been attributed to activation of these pathways, their relative importance downstream of the PDGFRβ has not been determined in vivo.
We have concentrated our present analyses on signal transduction by the PDGFRβ. Previous studies using a null allele of the receptor have demonstrated that PDGFRβ signal transduction is required for a subset of vascular smooth muscle cells and pericytes (v/p) (Levéen et al. 1994; Soriano 1994). These cells are the mesenchymal support cells that surround blood vessels (reviewed by Hungerford and Little 1999). Brain pericytes, kidney mesangial cells, retinal mural cells, and limb and skin pericytes have all been recognized as PDGFRβ-dependent cells (Lindahl et al. 1997a, 1998; Hellström et al. 1999; Enge et al. 2002). Studies have indicated that the PDGFRβ is likely to play a key role in the proliferation, migration, or both of a progenitor population (Hellström et al. 1999). These results explain why defective PDGF signal transduction results in a reduction of the v/p cell lineage and ultimately in perinatal lethality due to vessel instability (Hellström et al. 2001).
To examine the roles of PI3K and PLCγ downstream of the PDGFRβ, we have previously disrupted their binding sites in the receptor's cytoplasmic domain (Heuchel et al. 1999; Tallquist et al. 2000). Surprisingly, no overt phenotypes were detected in homozygous mutants lacking these two pathways, and deficiencies were observed only when the animals were challenged physiologically. To assess the roles of the remaining signal transduction pathways, we have created a PDGFRβ allelic series in mice (Figure 1). We refer to this series as the F series because it contains Y–F mutations at the known phosphorylated tyrosine residues. Using v/p cell number as a readout for PDGFRβ signal transduction, we have determined that the level of receptor expressed as well as the sum of signaling pathways induced by the PDGFRβ determines the number of v/p cells that form. These results provide an example of RTK signal transduction quantitatively controlling cellular development.
Results
Generation of the Allelic Series
Previous studies of the PDGFRβ have revealed an essential role for this receptor in v/p development, but attempts to identify essential biochemical signals thus far have demonstrated that loss of certain signaling pathways only diminishes PDGFRβ-driven responses (Heuchel et al. 1999; Tallquist et al. 2000). To study key signaling pathways, we have generated an allelic series of PDGFRβ mutants. Figure 1 illustrates the mutations that we have generated in the PDGFRβ locus and the signaling pathways that are disrupted by these mutations. Each mutant will be referred to by the number of tyrosines (Y) that have been mutated. For example, the mutation in the RasGAP-binding site is the PDGFRβF1/F1 or F1/F1 mutant. The truncation mutation of the PDGFRβ (βT) was created by the introduction of a frameshift and subsequent premature stop codon downstream of the RasGAP-binding site. Figure 2 illustrates the targeting events that were used to generate the series of mutants. The F1-, F2-, F3-, F5-, and βT-targeted mutations were generated by engineering Y–F, Y–I, or frameshift mutations in the same targeting vector (Figure 2A). The F7 mutation was generated by targeting the F5 heterozygous embryonic stem (ES) cells (Figure 2B; see Materials and Methods). Cells that contained all mutations on the same allele, as determined by Southern blotting, were used to generate the F7 line. All mutant mice were viable and fertile as homozygotes except the truncation allele, βT, which lacks the second half of the kinase domain and the SHP-2- and PLCγ-binding sites. Based on a similar mutation in the PDGFRβ, we assume that this receptor is kinase deficient and incapable of inducing DNA synthesis, but it still should bind ligand and undergo receptor downregulation (Escobedo et al. 1988). Embryos homozygous for the βT allele die perinatally with a phenotype identical to that of the PDGFRβ null embryos. E18 embryos exhibit edema and hemorrhaging in multiple tissues, including the kidney, brain, and skin (data not shown). These results suggest that PDGFRβ kinase activity is required for v/p development and that the receptor cannot function in the absence of kinase activity, unlike another RTK, vascular endothelial cell growth factor receptor 1 (Hiratsuka et al. 1998).
Identification of V/P Cells
We examined the blood vessels of F series homozygous mice by histology and detected no gross abnormalities (data not shown). To obtain a more global perspective of v/p cell populations, we introduced the XlacZ4 transgenic marker into our F series mutant mice. The XlacZ4 transgenic mouse expresses nuclear β-galactosidase in certain populations of differentiated, nonproliferating v/p cells in the embryo and the adult (Tidhar et al. 2001). As described below, using this marker in adult animals, we identified vascular defects in the F5 and F7 mice in the tissues of the eyes, hearts, and brains (see Figures 7, 8, and 9; data not shown). This observation suggests that both the F5 and F7 alleles function suboptimally in tissues known to require PDGFRβ signal transduction (Lindahl et al. 1997a; Hellström et al. 1999; Enge et al. 2002). Although both of these mutations cause notable phenotypes in some v/p populations, we have not observed pathologies in all populations of PDGFRβ-dependent v/p cells. V/p cell populations with no overt phenotype in the F5 and F7 mice include the kidney mesangial cells and pericytes in the skin and skeletal muscle (data not shown). We have observed a modest decrease in the number of nuclei present in F5/F5 and F5/− kidney glomeruli, but have not detected glomerulosclerosis with Masson trichrome stain (data not shown). The lack of any pathological phenotype in these tissues suggests either that the reduction in v/p cells is less severe than in the case of the PDGFRβ null mice, that the PDGFRα may be coexpressed in these same tissues, or that these tissues can function adequately even with reduced v/p cell numbers.
Because some populations of v/p cells appear to be more dependent on PDGFRβ signal transduction than others, we reasoned that the PDGFRα might be coexpressed in the less-affected v/p populations. Although PDGFRα has been reported in a variety of mesenchymal cell lineages (Schatteman et al. 1992; Lindahl et al. 1997b; Takakura et al. 1997; Zhang et al. 1998; Karlsson et al. 2000), we wanted to determine whether any v/p populations express the PDGFRα or whether it may be upregulated in any of the F series mice. We crossed the PDGFRαGFP line of mouse, which expresses a nuclear-localized green fluorescent protein (GFP) under the control of the PDGFRα promoter (Hamilton et al. 2003), with the F5 mutant mice and compared the GFP expression pattern to the pattern of v/p cells in the kidney, eye, and brain (Figure 3; data not shown). We have used three independent markers to designate v/p cells: smooth muscle actin α (αSMA), desmin, and the XlacZ4 transgene. Although PDGFRα-expressing cells are found in the same tissues as v/p cell markers, there is no overlapping expression of GFP with any of the v/p cell markers in the arteries or veins in the vessels of the eye and brain. PDGFRα-expressing cells are also absent from the larger vessels of the kidney, but a population of GFP+ cells is detected within the kidney glomerulus (Figure 3A). These may be either kidney mesangial cells or vascular adventitial fibroblasts. Both are populations of cells that are known to express the PDGFRα (Seifert et al. 1998). These data indicate that PDGFRα is not expressed or upregulated in two of the most affected tissues of the mutant mice, the eye and the brain, but could be functioning as a surrogate coreceptor with the PDGFRβ.
V/P Development
To determine whether the reduction in v/p was caused by a gradual loss or a developmental defect, we examined pericyte populations in wild-type and mutant embryos. The XlacZ4 mouse marker can be used to identify specific v/p cell populations as early as E12.5. We chose to observe pericytes at E14.5 because at this timepoint v/p are abundant in wild-type animals in several tissues, including the developing spinal cord and intercostal vasculature. Figure 4 demonstrates whole-mount visualization of the v/p cell populations in E14.5 wild-type embryos and the most severe F series mutant embryo (F7/−). After examining several litters of F series mutant embryos bearing the XlacZ4 marker, it was clear that the entire panel of F series homozygous mutant embryos could be distinguished from wild-type embryos simply by the degree that blood vessels had acquired v/p (data not shown).
To obtain a quantitative view of these results, we chose to focus on the spinal cord pericyte population. These cells begin to form at E10.5 in a rostral-to-caudal fashion in the embryo and require PDGFRβ signals for development (Levéen et al. 1994). Cross-sections through the developing spinal cord (neural tube) provide a relatively uniform area for quantitation. We can consistently identify a particular maturation stage of the developing vasculature based on its axial level within the embryo, and the pericytes can often be found as isolated cells (Figure 5). Using the entire panel of PDGFRβ mutant mice, we compared pericyte numbers between the different F series mutants (Figures 5 and 6). In all mutants examined, with the exception of the F1 mutation, we observed a decreased incidence of pericytes when compared to the wild-type embryos. The reduction in pericyte numbers ranged from 42% to 77%. This reduction was present at the more mature axial level of the heart as well as at the axial level of the kidney.
The F7/F7 mutant embryos are the only embryos that exhibited a difference between the number of pericytes at the heart level versus the number at the level of the kidney. All other mutants demonstrated similar numbers at both levels, indicating that pericyte development is disrupted and does not reach homeostasis as the tissue matures. Because the F7 is the most severely affected allele, it is possible that the difference between the heart and kidney levels is due to a developmental delay in v/p formation. Pericyte development may still be proceeding at the level of the kidney in these embryos. At the more mature level of the heart, the F7/F7 pericyte populations have reached a steady-state level and resemble v/p numbers more similar to those observed in the F5/F5 embryos.
Previously, chimeric analysis had demonstrated that PDGFRβ heterozygous cells do not contribute extensively to the smooth muscle cell compartment, suggesting that heterozygous cells may have reduced v/p developmental potential (Crosby et al. 1998). To find out whether receptor levels had any impact on v/p cells in our system, we crossed animals bearing the PDGFRβ null allele to our mutant series (Figures 5B and 6B). We observed an even further reduction in pericyte levels, resulting in a 70%–92% decrease in pericytes when compared to wild-type embryos. Interestingly, even the PDGFRβ+/− embryos, which exhibit a approximately 50% reduction in PDGFRβ messenger RNA (Soriano 1994), demonstrate a nearly 40% decrease in pericytes. This result suggests that the quantity of receptor impacts the number of pericytes. Another observation from this data is that even the F7/− embryos can induce cell development at levels greater than the null. In fact, the F7/− animals survive, whereas the PDGFRβ nulls do not. This is a rather surprising result given that most of the downstream signal transduction molecules that directly interact with the receptor have been dissociated.
While most of the F series alleles demonstrate a decrease in v/p cells, the F1 allele results in an apparent increase in spinal cord pericytes. Although the increase is most pronounced when we compare the F1 hemizygote to the PDGFRβ heterozygote (Figure 6), an increase is also observed when comparing F1/F1 embryos and wild-type embryos. In fact, the level of pericytes in the F1/− embryos is very similar to those in the wild-type. These data demonstrate two interesting findings. One is that RasGAP may play a role in PDGFRβ signal attenuation and that loss of this pathway results in increased PDGFRβ signals. The second is that v/p numbers may not be tightly controlled and that PDGFRβ signaling can result in more cells.
To determine whether the signaling pathways affected other v/p populations in the same manner, we have examined the v/p population in the retina. It has been shown previously that PDGFB and PDGFRβ signaling controls pericyte development in the eye (Benjamin et al. 1998; Klinghoffer et al. 2001; Enge et al. 2002). Adult mice transheterozygous for one null allele and one F5 or F7 allele exhibited severe eye defects. These defects were first observed as an opacity and sometimes as visible hemorrhage in the eye (Figure 7A), as previously described for PDGFRβ and PDGFB signaling mutants (Klinghoffer et al. 2001; Enge et al. 2002). The F5 and F7 hemizygous mutant mice possessed fewer discontinuous blood vessels and overgrowth of retinal cells. This phenotype occurred with 100% penetrance, but with variable severity (Figure 7C), and was detectable sometimes as early as 4 d after birth. The presence of a pathological condition suggests that the F5 and F7 alleles have compromised receptor function when compared to the wild-type, F1, F2, and F3 alleles and demonstrates that retinal pericytes are also dependent on the PDGFRβ signaling pathways that we have disrupted.
To examine the retinal pericytes in the entire F series, we again used mice bearing the XlacZ4 transgene. At 4 wk of age the retinal vasculature is mature and can be isolated from the lens and pigmented epithelium for visualization. Figure 8 illustrates that homozygotes for the F1, F2, and F3 mutant alleles are indistinguishable from wild-type eyes; however, F5/F5 and F7/F7 eyes exhibit reduced numbers of pericytes. Even without the ability to quantitate these differences, it is clear that the PDGFRβ+/−, F5/F5, F7/F7, F5/−, and F7/− mutant retinas have a reduction in v/p when compared to wild-type eyes, reinforcing the requirement for multiple PDGFRβ signal transduction pathways in v/p development.
A final tissue where we have examined v/p formation is the heart. F2 and F3 homozygotes and transheterozygotes were indistinguishable from wild-type and heterozygous hearts, respectively. Consistent with our observations in the eye and the nervous system, the F5 and F7 mutant alleles display abnormalities in the vascular coating of their coronary arteries and veins (Figure 9; data not shown). F5/− and F7/− mice often exhibited a variety of heart abnormalities, including enlarged ventricles, increased heart:body mass ratio, dilated atria, and fibrosis (data not shown). In contrast, the F1/+ mice appeared to have more extensive v/p coating on their coronary arteries (Figure 9). In agreement with the data from the nervous system and the eye, the F5 and F7 mutant alleles have a significant reduction in v/p cells.
Taken together, these results demonstrate several important findings for PDGFRβ signal transduction. First, the number of pericytes formed directly correlates with the number of signaling pathways transducing PDGFRβ activity. Second, a reduction in pericytes is observed even when only the amount of receptor is affected. Finally, although SH2 domain-containing proteins impact v/p numbers, the intrinsic kinase activity of the receptor may play a role in transmitting the PDGFRβ signal because the truncation mutation does not exhibit any rescue of v/p development, while the F7 mutant allele that transmits primarily through kinase activity (owing to loss of the SH2 domain-containing protein-binding sites) still supports v/p development sufficient for viability.
Downstream Signal Transduction
Because F2, F3, and F5 mutant receptors have been previously studied biochemically (Valius and Kazlauskas 1993; Heuchel et al. 1999; Tallquist et al. 2000), we have focused our biochemical analysis on the F1 and F7 mutant receptors' signal transduction to verify the effects of these particular mutations on downstream signal transduction cascades. We have used mouse embryo fibroblasts (MEFs) for these analyses. All lines of MEFs that we generated expressed the PDGFRβ at similar levels (Figure 10D) as well as the PDGFRα (data not shown). To avoid stimulation of the PDGFRα by PDGFBB, we downregulated PDGFRα surface expression by pretreatment with PDGFAA 2 h before PDGFBB stimulation. In all cell lines examined, we observed an increase in tyrosine phosphorylation in response to ligand (Figure 10A). The most evident phosphorylated bands are around 200 kd, which are likely to be the PDGFRα and PDGFRβ. Although we have mutated seven of the 13 tyrosines, a significant amount of phosphorylation is observed in all cell lines, albeit at lower levels in the F7/− cell line (Figure 10A and 10B). In the whole-cell lysate phosphotyrosine blot, the phosphorylated protein detected at 200 kd is likely cytoplasmic PDGFRα, as it is reduced in F7 cells after downregulation of the PDGFRα.
Because we have disrupted only one of the potential Src-binding sites, we examined the level of Src activation downstream of our F7 cell line (Figure 10B). Upon PDGFBB addition, there was an increase in the amount of Src phosphorylated on tyrosine 418 (a site whose phosphorylation is required for full catalytic activity; Johnson et al. 1996). In contrast, in the F7/F7 MEFs, we did not observe any increase in Src activation. These results are in agreement with other reports that demonstrate that a mutation at amino acid 578 of the PDGFRβ is sufficient for reducing the level of Src binding and activation (Mori et al. 1993; Twamley et al. 1993; Vaillancourt et al. 1995; Fanger et al. 1997).
Two potential downstream targets of PDGFRβ activation are activation of extracellular signal-related kinases 1 and 2 (ERK1/2) and AKT (Franke et al. 1995). As expected, phosphorylation of ERK1/2 and AKT is reduced or absent in the F7 homozygous and hemizygous mutant cell lines, but cells expressing at least one copy of the wild-type receptor are capable of inducing the activation of these downstream molecules (Figure 10C). These data demonstrate that loss of seven tyrosine residues on the PDGFRβ results in a severe loss of downstream signal transduction. In contrast, cells bearing even just one copy of the F1 receptor show increased phosphorylation of ERK1/2. These data are in concordance with the in vivo data, which show that lack of the RasGAP-binding site on the receptor results in an increase in the downstream signaling events and a subsequent increase in v/p. Therefore, the F1 mutant receptor has increased activity while the F7 receptors have decreased activation of these same pathways, despite having apparently normal levels of kinase activity.
Discussion
RTK signal transduction plays an important role in directing many cellular activities. We have used an in vivo system to analyze how cellular development relates to the signaling pathways downstream of the PDGFRβ. Examination of the v/p population demonstrates a quantitative relationship between the extent that signals are being transduced and the number of v/p that form. Several other studies have demonstrated that combinations of signal transduction pathways may dictate cellular outcomes. Examples of these are T cell development in the immune system and gradients of morphogens in developmental systems (Heemskerk and DiNardo 1994; Nellen et al. 1996; Zecca et al. 1996; Ferrell and Machleder 1998; Gong et al. 2001).
In our system, the signal can be affected in two ways. The first is by the amount of receptor expressed at the cell surface. V/p numbers are significantly lower in PDGFRβ+/− embryos when compared to wild-type controls. In this situation there is a decrease in overall signal, but no specific, directly associated pathway is disrupted. This demonstrates a quantitative role for receptor activity. The second influence on PDGFRβ signal transmission is by the number of associated SH2 domain-containing proteins. Loss of even a single pathway results in reduction of v/p, and as the number of disrupted pathways increases, there is a concomitant decrease in v/p. There is no significant difference between the F2 and the F3 mutant alleles, but there is a noticeable difference when the number of mutations is further increased. These signaling differences as illustrated by v/p number and the presence of vascular pathologies can be categorized in the mutant alleles by the following hierarchy: F1 > wild-type > F2 = F3 > F5 > F7 > null. In addition, hemizygotes show an even further reduction in v/p when compared to the F series homozygotes. This suggests that specific effector pathways may play more of a role in fine-tuning PDGFRβ signals.
In total, our results demonstrate that PDGFRβ signal transduction is regulated not only by direct binding of signal transduction molecules, but also by receptor expression levels, possibly reflecting inherent kinase activity. In addition, no particular signaling pathway that we have analyzed is absolutely required for transmission of PDGFRβ signals, because even the F7 allele has a phenotype less severe than the null. In support of the observation that receptor levels and kinase activity may have a direct role in signal transduction, we have observed that a chimeric PDGFR that has the extracellular domain of the PDGFRβ but the intracellular domain of the PDGFRα exhibits a more severe phenotype than the F5 allele (Klinghoffer et al. 2001). This chimeric receptor can signal through all of the same downstream components as the PDGFRβ except for RasGAP, suggesting that the more severe vascular defects in these chimeric receptor mice may be due to reduced kinase activity and/or expression levels of the chimeric receptor.
The PDGFRβ's signaling pathways appears to dictate the absolute numbers of v/p that form, but how the individual pathways contribute to this phenotype remains to be tested. In fact, very little difference in numbers of v/p is observed between the F2 and the F3 mutants, suggesting that PLCγ signals may be somehow redundant with or dependent on the PI3K pathway. In contrast, loss of additional pathways leads to an incremental loss of v/p. The difference between the F3 and the F5 mutations is the ability to bind SHP-2 and RasGAP, and it has been proposed that both of these molecules play roles in downregulating the PDGFRβ signal (Klinghoffer and Kazlauskas 1995; Ekman et al. 1999). Our results demonstrate that loss of these signaling pathways is detrimental to PDGFRβ signal transduction and that both may have positive and negative influences on receptor activity.
There are several potential ways that loss of these signaling pathways leads to v/p reduction. One mechanism would be that each pathway contributes to a specific cellular outcome. For example, SFK's predominant role could be to promote proliferation (Roche et al. 1995; Hansen et al. 1996), whereas PI3K activity could be more important for migration (Kundra et al. 1994; Wennstrom et al. 1994b). Therefore, the combined loss of these pathways results in a net reduction in v/p, albeit for entirely different cellular reasons. A second scenario would be that all pathways lead to a single or few specific cellular conclusions. Thus, loss of any one pathway only reduces the outcome but does not ablate it. Evidence from immediate-early gene expression analysis suggests that this mechanism may occur (Fambrough et al. 1999), although this possibility does not require that all pathways contribute equally. Last, some pathways may play a primary role downstream of the receptor, while others may be more secondary.
Our data suggest that PI3K may be a principal pathway, while the other pathways may be less significant. The F2/F2 mutant mice have a significant reduction in v/p numbers when compared to the wild-type and the heterozygous mice, and additional mutations have less of an effect than PI3K on v/p numbers. A similar situation has been observed with the PDGFRα (Klinghoffer et al. 2002): the phenotype of mouse embryos with loss of the PDGFRα–PI3K pathway was just as severe as that of embryos expressing a PDGFRα F7 allele (which is similar to the F7 allele of the PDGFRβ). The pathological difference observed between the F5 and F7 alleles versus all of the other mutations suggests that SHP-2, SFK, Grb2, and RasGAP also impact pericyte development. Assessing the importance of these pathways would require generating additional alleles with different combinations of mutant sites. The only signaling differences between the F3 mutation and the F5 mutation are the SHP-2 and RasGAP pathways. This suggests that one or both of these pathways promote PDGFRβ signaling, in contrast to the F1 mutation, which demonstrates that absence of RasGAP leads to a potential increase of PDGFRβ signaling. This apparent contradiction may indicate that RasGAP plays both a positive and a negative role in signal transduction or that SHP-2 may have mainly a positive role in modulating this response.
Although we find that overall loss of downstream pathways attenuates receptor actions as demonstrated by v/p formation, it is surprising that the F7/F7 mice do not phenocopy the null animals. The F7 allele possesses disruptions at seven of the 13 known phosphorylated tyrosine residues. These mutations should disrupt a majority of the signal relay molecules downstream of the receptor. The remaining tyrosines are capable of binding SFKs, STATs, and Grbs. Based on several previous reports, disruption of Y578 affects the majority of SFK binding (Mori et al. 1993; Twamley et al. 1993; Vaillancourt et al. 1995; Fanger et al. 1997), and we have shown that SFKs do not become activated after stimulation of the F7 receptor. As for the signaling roles of STAT and Grb2 downstream of the receptor, little direct function has been demonstrated for these remaining effector molecules in PDGF-induced cellular responses (Heldin et al. 1998). Therefore, F7 signal transmission must use some other means than direct binding by SH2 domain-containing proteins. The receptor should still have full kinase activity, unlike the lethal βT mutation, which is lacking half of the kinase domain and the SHP-2- and PLCγ-binding sites. Possibly, the receptor is phosphorylating molecules that are only transiently associated. Another possibility is that other receptors may function as surrogates. The most likely surrogate is the PDGFRα, but we have demonstrated that in several of the v/p cell populations the PDGFRα is not expressed. Other candidate molecules for such a mechanism are integrins, ephrins, and the low-density lipoprotein receptor-related protein LRP (Miyamoto et al. 1996; Schneller et al. 1997; Woodard et al. 1998; Boucher et al. 2002; Loukinova et al. 2002). Although these proteins are known to cross-talk with the PDGFRβ, it is unclear whether they have the capability to substitute for the PDGFRβ's own signaling components.
The F1 mutant allele is an interesting corollary to the F mutant series. While all of the other mutations appear to have a detrimental effect on PDGFRβ signal transduction, the F1 mutation results in an apparent increase in PDGFRβ activity as determined by v/p incidence. These data are in agreement with previous observations that RasGAP function decreases the Ras/MAP kinase pathway activity and migration (Kundra et al. 1994; Ekman et al. 1999). In addition, an add-back mutation of the RasGAP-binding site induced a different gene profile from the PDGFRβ immediate-early gene profile (Fambrough et al. 1999). This suggests that RasGAP may have different signaling capabilities from the other PDGFRβ signal transduction components.
The mutant mice not only uncover the role of RTK signal transduction in vivo, but they also reveal some interesting information regarding v/p cell development. For example, although v/p cell development is impaired when PDGFRβ signal transduction is disrupted, a basal level of cells forms, in agreement with previous observations that propagation, not initiation, of v/p cell development is directed by the PDGFRβ (Lindahl et al. 1997a; Hellström et al. 1999). Even in the null embryos, v/p cells can be found. It has been proposed that PDGFRβ signals are required for the expansion of v/p cells (Lindahl et al. 1998). While this may be the case, it is curious that in the F5 and F7 animals, the pericyte numbers never reach wild-type levels, resulting in vascular pathologies.
There are two explanations for the observation that v/p cells never reach wild-type levels. The first is that there is constant turnover in the v/p population and that the rate of replacement in the mutant mice is below the rate of loss, resulting in a net reduction in the v/p population. Evidence against this mechanism is the failure to observe any significant proliferation in the adult wild-type animals under normal conditions or significant apoptosis in the mutant panel of mice (data not shown). The second possibility is that there is a specific window during development when v/p cells can expand. After a specified time, v/p cell number expansion could be limited, perhaps related to the ability of endothelial cells to secrete the PDGF ligand (Benjamin et al. 1998). Support for this model is the inability of nascent endothelial tubes to recruit v/p cells in tumors (Abramsson et al. 2002). The inability to develop sufficient numbers of v/p cells also appears to be recapitulated in the eye vasculature, suggesting that the maturation of the vessel is more dependent on the local environment than on the chronological age of the embryo.
Our findings demonstrate that the combination of signaling pathways downstream of PDGFRβ determines the total number of v/p cells. These can be modulated not only by the amount of receptor expressed at the cell surface, but also by the number of specific downstream signaling pathways activated by the receptor. Whether these results are unique to PDGFRβ signal transduction in v/p cells or whether they can be extrapolated to other RTK remains to be demonstrated.
Materials and Methods
Mice
Point mutations that disrupt the designated signal transduction pathways were generated by changing the tyrosine residue to phenylalanine. The exception was Y1020, which was mutated to encode an isoleucine, thus generating a unique restriction site that facilitated identification of homologous recombinants. Mouse mutants F2 and F3 have been previously described (Heuchel et al. 1999; Tallquist et al. 2000). The targeting vector for the F1, F5, and βT mutations utilized the same arms of homology as the F3 vector. The exons containing the point mutations were introduced in the arms of homology of the targeting vector by site-directed mutagenesis and verified by sequence data of PCR-amplified genomic DNA from homozygous mutant mice. The F7 mutation was generated by creating a targeting vector that incorporated the 5′ arm of the F5 targeting vector with 5′ genomic sequences that included the exons containing the Src- and Grb2-binding sites. Tyrosines 578 and 715 were mutated to phenylalanine to disrupt Src and Grb2 binding, respectively. This targeting vector was transfected into F5 heterozygous ES cells and screened for homologous recombination. The truncation mutation possesses a frameshift at amino acid 780, resulting in a premature stop codon after amino acid 801, 11 amino acids downstream of the RasGAP-binding site. ES cell colonies were screened initially by PCR, and positive clones were further verified by Southern blot analysis for the correct recombination at the 5′ and 3′ arms. The PGK–neo cassettes were removed by crossing mice to Meox2Cre (Tallquist and Soriano 2000) and ROSA26FlpeR (Farley et al. 2000) deleters.
The majority of analyses have been carried out on a mixed 129S4 × C57BL/6 background, except where indicated. The XlacZ4 transgenic mouse (Tidhar et al. 2001) was kindly provided by Moshe Shani and crossed into the F series. We also crossed the F5 and wild-type mice to the PDGFRαGFP line (Hamilton et al. 2003).
Histology, Immunohistochemistry, and Pericyte Quantitation
Embryos and tissues were processed and embedded for sectioning according to standard protocol. We have not examined the vasculature of all PDGFRβ-dependent tissues in the F series mutant animals. Those tissues not examined are lung, brown adipose tissue, and the adrenal gland.
Immunohistochemistry
Kidneys were removed and fixed for 20 min in 4% paraformaldehyde, 200 μm sections were then obtained by vibratome sectioning, and immunofluorescence was performed. For eye immunohistochemistry, the pigmented epithelium was removed from the mouse retinas and fixed for 10 min in 4% paraformaldehyde. Retinas and kidney slices were then blocked and subjected to immunohistochemistry for the indicated v/p marker. Antibodies were β-galactosidase (55976; Cappel, Costa Mesa, California, United States), αSMA (1A4; Sigma, St. Louis, Missouri, United States), and desmin (D33; Dako Cytomation, Glostrup, Denmark). Photographs were obtained on a Zeiss Axiophot microscope (Carl Zeiss MicroImaging, Thornwood, New York, United States).
Pericyte quantitation
E14.5 embryos were divided into quarters at the following levels: head–neck, neck–liver, liver–kidney, and kidney–tail. Quarters were rinsed with PBS and fixed for 20 min in 2% formaldehyde, 0.2% glutaraldehyde. They were then washed three times in PBS, stained overnight with X-Gal, transferred to PBS, photographed, post-fixed in 10% formalin, and then processed and embedded. Sections (7 μm) were generated and X-Gal-positive nuclei quantitated in the neural tube at the level of the heart and the kidney. Seven to ten samples were counted for each level; the mean of these data is represented in Figure 6. Pericytes surrounding the exterior of the neural tube were excluded from the sample. Positive nuclei were counted at 20× magnification and photographed at 10× magnification on Zeiss Axiophot microscopes. Retinas were prepared in a similar manner. The pigmented epithelium was removed prior to the initial fixation step, and the lens was not removed until after the final fixation to maintain retina shape. Images were obtained on a Nikon SMZ1000 with a Coolpix 900 camera (Nikon Corporation, Tokyo, Japan).
Immunoprecipitation and Western Blotting
MEFs were generated from E9-d-old or E14.5-d-old embryos. Embryos were isolated, decapitated, and eviscerated. The remaining tissue was then treated with trypsin and plated. Cells were frozen down at passages 2 and 3. Most experiments were completed on cells at passages 3–6, except for the wild-type line, which was spontaneously immortalized.
Cells were plated at 1 × 105 to 3 × 105 cells per well and starved for 48 h. Receptor downregulation was achieved by treating starved cells for 2 h with 100 ng/ml PDGFAA (R&D). Cells were then stimulated with PDGFBB (R&D) for 5 min and lysed.
Immunoprecipitation and Western blotting were executed as previously described (Tallquist et al. 2000). Antibodies were obtained from the following sources: PDGFRβ (06-498; Upstate Biotechnology, Lake Placid, New York, United States); PDGFRα (sc-338; Santa Cruz Biotechnology, Santa Cruz, California, United States); Akt (9272; Cell Signaling Technology, Beverly, Massachusetts, United States); Phospho-AKT (9271; Cell Signaling Technology); RasGAP (05-178; Upstate Biotechnology); Grb2 (610111; BD Transduction Laboratories, San Jose, California, United States); ERK1/2 (06-182; Upstate Biotechnology); c-Src (SRC2 and sc-18; Santa Cruz Biotechnology); Phospho-Src Y418 (44-660; Biosource International, Camarillo, California, United States); Phospho-ERK1/2 (9101; Cell Signaling Technology); PLCγ (05-163; Upstate Biotechnology); SHP-2 (sc-280; Santa Cruz Biotechnology); and phosphotyrosine (4G10) (05-321; Upstate Biotechnology). PDGFRβ 97A (kinase insert domain) was a kind gift from Andrius Kazlauskas.
Supporting Information
Accession Numbers
The LocusLink ID numbers discussed in this paper are PDGFRα (Locus ID 18595) and PDGFRβ (Locus ID 18596).
Acknowledgements
We thank Moshe Shani for the XlacZ4 transgenic line; Andrius Kazlauskas for antibodies; Elizabeth Behler, Philip Corrin, Triston Dougall, Jason Frazier, and Tara Swamy for excellent technical support; and our laboratory colleagues for discussions and critical comments on the manuscript. MDT was supported in part by a fellowship from the American Cancer Society (PF-98–149-01). This work was supported in part by research grant 0330351N from the American Heart Association and grant 5-FY03–14 from the March of Dime Birth Defects Foundation to MDT and by grants HD24875 and HD25326 from the National Institute of Child Health and Human Development to PS.
Abbreviations
ERK - extracellular signal-related kinase
ES - embryonic stem
GFP - green fluorescent protein
MAP kinase - mitogen-activated protein kinase
MEF - mouse embryonic fibroblast
PDGF - platelet-derived growth factor
PDGFR - platelet-derived growth factor receptor
PI3K - phosphatidylinositol 3′-kinase
PLCγ - phospholipase Cγ
RTK - receptor tyrosine kinase
SFK - Src family kinase
SH2 domain - Src homology domain 2
SHP-2 - SH2-containing phosphotyrosine phosphatase
αSMA - smooth muscle actin α
STAT - signal transducer and activator of transcription
v/p - vascular smooth muscle cell/pericyte
VSMC - vascular smooth muscle cell
Figures and Tables
Figure 1
PDGFRβ Allelic Series
This figure depicts the mutant alleles generated in the mouse PDGFRβ genomic locus. X represents a mutation in the tyrosine-binding site(s) for a particular signal transduction molecule. The F7 allele contains a disruption in one SFK-binding site because loss of both sites results in diminished kinase activity (Mori et al. 1993). The truncation allele (βT) was created by deletion and subsequent frameshift that results in a stop codon 32 amino acids past the RasGAP-binding site.
Figure 2
Targeting Strategy and Southern Blot
(A) Targeting vector used to create F5 mutant allele. Two exons contain all five mutated tyrosines.
(B) Targeting vector containing mutations in 5′ exons used to generate the F7 mutant allele.
(C) Wild-type allele.
(D) Targeted allele with PGK–neo removal. Restriction enzyme abbreviations: Sp, SpeI; A, Asp718; S, SacI; RV, EcoRV; H, HindIII; X, XhoI; and RI, EcoRI. Green boxes indicate probes used in Southern blotting for F7-targeted ES cells. The blue arrow indicates the exon where point mutation causes frameshift in truncation mutation. Black boxes indicate wild-type exons. Red boxes indicate exons containing targeted mutations. Restriction enzymes in red indicate sites introduced by mutagenesis to verify proper homologous recombination by Southern blotting. Circles denote FRT sites. Triangles denote loxP sites.
(E) Southern blot results from various ES cell lines. SpeI digest using P1 probe. Blot with P2 probe gave expected results (data not shown). Lanes 1 and 3, F5 allele targeted. Lanes 2 and 4, wild-type allele targeted. Lanes 5 and 7, F5 mutant ES cells after and before Cre activity, respectively. Lane 6, wild-type ES cell clone.
Figure 7
Eye Defects in F5/− Mice
(A) Eyes from a P4 F5/− mouse demonstrating severe hemorrhaging. (B and C) H/E-stained sagittal sections through eyes of wild-type and F5/− 3-mo-old mice, respectively. The absence of the lens of the F5/− eye is a histological defect and not a phenotype of the F5/− eye. L, lens; R, retina.
Figure 8
V/P Populations in P28 Retinas
Whole-mount retinal preparations from wild-type and mutant eyes. Pigmented epithelium was removed for visualization of β-galactosidase. Note F7/F7 and F7/− had extensive thickening of the retinal layers that resulted in a contraction of the entire retina and apparent reduction in size. The far right panel shows a close-up of the artery and vein of three homozygous eyes.
Figure 9
Vascular Smooth Muscle Cells of the Coronary Arteries
(Top) Whole-mount views of P21 hearts from littermates of the F5 alleles of mutant mice. Hearts were sliced coronally, and the ventral surface was photographed. The F5/− heart was sliced disproportionately and therefore appears to be smaller.
(Bottom) P28 hearts from wild-type and F1 littermates. Hearts were sliced sagittally. Both the left and right views are shown.
Figure 3
Tissue Localization of V/P Cell Markers and PDGFRα Expression
Tissue preparations from P21 PDGFRαGFP/+;PDGFRβF5/F5 mutant mouse. Immunofluorescence was used to detect αSMA (A and B), desmin (C), β-galactosidase (D), and GFP expression for PDGFRα (A–D). (A) Kidney (200 μm vibratome section). The arrow indicates glomerulus. The asterisk indicates an arteriole. (B–D) Retina (whole-mount preparation). Arrowheads point to β-galactosidase-positive nuclei.
Figure 4
Reduction in V/P Cells in the Thoracic Region of E14.5 Embryos
Ventral view of E14.5 wild-type and F7/− littermates with the XlacZ4 mouse marker background. β-Galactosidase-positive nuclei represent v/p cells. Th, thymus.
Figure 5
Pericytes within the E14.5 Nervous System of F Series Mutant Embryos
(A) Representative sections through neural tubes of embryos from the homozygous F allelic series. (B) Representative sections of embryos from the hemizygous allelic series (F series mutant with one copy of the null allele). Sections are from the rostral level between the heart and kidney. Pericytes are visualized by nuclear-localized β-galactosidase staining in cells committed to the v/p lineage. Sections are 7 μm.
Figure 6
Quantitation of Pericytes in Nervous System
β-Galactosidase-positive nuclei were counted within the neural tube. Each datapoint represents a mean of seven to ten sections from a single embryo. Data were gathered at two rostral levels in each embryo. The genotypes are ordered by the predicted strength of the signal, depending on the number of copies of the receptor being expressed and the signal transduction pathways remaining downstream of the receptor. A two-tailed Student's t-test was performed comparing the number of spinal cord perictyes in mutants to those in wild-type, and all homozygous mutants and hemizygous mutant values were statistically different (p < 0.005) from wild-type values. The F1/− was also statistically different from the PDGFRβ+/− (p < 0.001).
Figure 10
Biochemistry of MEFs from F7 and F1 Mice
(A) Whole-cell lysates were generated from MEFs that were unstimulated or stimulated with PDGFAA and/or PDGFBB alone, 100 ng/ml and 30 ng/ml, respectively. Lysates were then subjected to SDS–PAGE and Western blotting accomplished with the anti-phosphotyrosine antibody (4G10).
(B) Immunoprecipitation of tyrosine-phosphorylated proteins from wild-type and the F7 series of mutant MEFs. The precipitates were then run on SDS–PAGE, and a Western blot was performed using anti-Src [pY418] antibody and anti-PDGFRβ 97A.
(C) Whole-cell lysates from unstimulated or stimulated MEFs. Lysates were then subjected to SDS–PAGE and Western blotting accomplished with the indicated phosphorylated-specific antibodies. Blots were stripped and blotted with antibodies to the corresponding unphosphorylated proteins to demonstrate protein loading. Data are representative of multiple experiments from at least two independently derived cell lines.
(D) Western blots of whole-cell lysates from 5 ×104 cells per lane blotted with antibodies to PDGFRβ (06-498) and ERK as a loading control.
Footnotes
Conflicts of Interest. The authors have declared that no conflicts of interest exist.
Author Contributions. MDT and PS conceived and designed the experiments. MDT and WJF performed the experiments. MDT analyzed the data. MDT wrote the paper.
Academic Editor: Roel Nusse, Stanford University School of Medicine | [
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14675480 | Knockout of ERK5 causes multiple defects in placental and embryonic development
Abstract
Backgroud
ERK5 is a member of the mitogen activated protein kinase family activated by certain mitogenic or stressful stimuli in cells, but whose physiological role is largely unclear.
Results
To help determine the function of ERK5 we have used gene targeting to inactivate this gene in mice. Here we report that ERK5 knockout mice die at approximately E10.5. In situ hybridisation for ERK5, and its upstream activator MKK5, showed strong expression in the head and trunk of the embryo at this stage of development. Between E9.5 and E10.5, multiple developmental problems are seen in the ERK5-/- embryos, including an increase in apoptosis in the cephalic mesenchyme tissue, abnormalities in the hind gut, as well as problems in vascular remodelling, cardiac development and placental defects.
Conclusion
Erk5 is essential for early embryonic development, and is required for normal development of the vascular system and cell survival.
Background
Mitogen activated protein kinase (MAPK) cascades play important roles in many cellular processes including cell proliferation, differentiation, survival and apoptosis. They are also important for many physiological functions in several systems, including in developmental, immune and neuronal systems. At least 12 isoforms of MAPKs exist in mammalian cells, and these can be divided into 4 main groups, the 'classical' MAPKs (ERK1 and ERK2), JNKs (also referred to as SAPK1), p38s (also referred to as SAPK2, SAPK3 and SAPK4) and atypical MAPKs such as ERK3, ERK5 and ERK8. With the exception of ERK3, MAPKs are activated by dual phosphorylation on a Thr-Xaa-Tyr motif by a dual specificity MAPK kinase (MKK). MKKs are in turn activated by a MAPK kinase kinase (MKKK), which are activated in response to appropriate extracellular signals.
ERK5 is an atypical MAPK that can be activated in vivo by a variety of stimuli, including some mitogens such as EGF, and some cellular stress such as oxidative and osmotic shock [1-3]. These stimuli activate a cascade in which the MAPK kinase kinases MEKK3 or MEKK2 activate MKK5, which in turn activates ERK5 [4,5]. Interest in the ERK5 pathway has been fuelled by reports that the activation of ERK5 by MKK5 can be blocked in vivo by the kinase inhibitors PD184352, PD 98059 and U0126. These inhibitors were developed as inhibitors of the classical MAPK cascade, and have been used extensively to study this cascade in vivo. The discovery that they can also block ERK5 activation, although at higher concentrations than are required to block the activation of ERK1 and ERK2, raised the possibility that ERK5 and ERK1/ERK2 may have some overlapping functions in vivo [6,7].
The physiological roles of ERK5 are still largely unclear. Overexpression of a constitutively active MKK5 in mice results in cardiac hypertrophy and death of the mice by 8 weeks of age [8]. This is suggestive of a role of ERK5 in the heart, possibly related to cardiac development. ERK5 has also been implicated in the development of smooth muscle, as ERK5 antisense oligonucleotides [9] or dominant negative ERK5 constructs [10] have been reported to block the differentiation of smooth muscle cells in cell culture models. At present little is known about the substrates for ERK5 in vivo, however it has been suggested to phosphorylate connexin 43 [11] and the transcription factor MEF2C [12-14]. Mouse knockouts of MEF2C are embryonic lethal, and MEF2C-/- embryos die due to a failure of the developing heart to undergo normal looping at E8.5-9 [15]. Knockout MEKK3 also results in embryonic lethality at E11, MEKK3-/- embryos show problems with myocardium formation, angiogenesis and placental formation [16]. While this could be consistent with a role for ERK5 in linking MEKK3 signalling to MEF2C during cardiac development, it should be noted that MEKK3 can activate other MAPK isoforms, particularly p38α (also referred to as SAPK2A) [17-20]. Knockout of p38α has been reported by several groups, and p38α-/- embryos have also been reported to show problems in cardiac development, angiogenesis and placental formation at E10-11 [21-23].
In order to further examine the role of ERK5 we carried out expression and gene targeting studies in mice. ERK5 knockout was found to be lethal during embryogenesis at E10.5 to E11, and here we report a detailed analysis of these embryos. While this work was in progress, both Regan et al [25] and Sohn et [26] also reported ERK5 knockouts, and the effects of these different ERK5 knockouts are considered in the discussion.
Results
Generation of ERK5 knockout mice
Sequencing of the mouse ERK5 gene showed that it comprised of 7 exons spanning 5.4 kb of genomic sequence. Of these, exons 2 to 7 encoded the sequence of ERK5, while the 5' untranslated region was located in exons 1 and 2, and the 3' untranslated region in exon 7. Based on this sequence a targeting vector was designed to delete exons 4 and 5 of ERK5 in ES cells (Fig 1). Correct incorporation of this vector was confirmed by Southern screening of ES cells (Fig 1A and 1B). Germline transmission was obtained from two independent targeted ERK5 ES clones, and the ERK5+/- mice were of similar size and morphology to wild type littermates. Breeding of ERK5+/- mice gave the expected numbers of wild type and ERK5+/- mice, however no ERK5-/- mice were obtained, indicating that the ERK5 knockout is lethal during embryogenesis. To determine the point of lethality, embryos were genotyped by a PCR based method (Fig 1A and 1C) from timed matings. The expected Mendelian numbers of homozygous knockout mice were found at E9.5 and 10.5 (table 1) and at this point knockout embryos were still alive, as judged by a beating heart. In contrast, at E11.5 all homozygous ERK5 knockout embryos found had died and were undergoing reabsorption. Similar results were obtained from crosses of the ERK5 mutation onto either Balb/C or C57/Bl6 backgrounds, and from two independent ES cell clones. The deletion made in the ERK5 gene removes the sequence encoding for amino acids 133 to 712, and introduced a neomycin resistance cassette, including a polyadenylation sequence into the ERK5 gene. While exons 1 to 3 remain in the targeted gene, insertion of the neomycin cassette would be expected to interfere with normal transcription and splicing after exon 3. Should exon 3 be able to splice onto exon 6 in the targeted gene, this would result in a frame shift mutation. To confirm that the knockout blocked the production of ERK5 protein, extracts from E9.5 embryos were analysed by immunoblotting using a polyclonal antibody raised against the whole ERK5 protein. ERK5 was detected in wild type embryos and in ERK5+/- embryos, however the levels of ERK5 protein were reduced in ERK5+/- embryos compared to wild type embryos. As expected no protein was seen for ERK5 in the ERK5-/- embryos. No evidence for the production of truncated forms of ERK5 in the ERK5-/- embryos could be seen in the immunoblots (data not shown). Expression of MKK5 and other MAPK kinases (ERK1, ERK2 and p38) were unaffected by the knockout of ERK5 (Fig 1d).
Figure 1
Generation of ERK5 knockout mice. A) ERK5 knockout mice were made using a targeting vector to delete exons 4 and 5 of the murine ERK5 gene through the addition of a neomycin selection cassette. A thymidine kinase cassette acts as a negative selection marker during ES cell selection. B) ES cell DNA was digested with both Hind III and Mfe I, and a Southern blot performed using a probe 3' to the targeting vector. The position of the wild type 9.5 kb fragment and targeted 3.3 kb fragment are indicated. C) DNA was isolated from E9.5 embryos and digested with Hind III and Mfe I. Southern blots were then probed with the 3' probe as described in (B)D) Soluble protein from homogenates of E9.5 embryos was run on 4–14% acrylamide gels. Immunoblotting was then carried out using antibodies which recognised ERK5, MKK5, ERK1/2 or p38.
Table 1
Ratios of ERK5 adults and embryos
* Of the E10.5 knockout embryos, 12 were class I and 16 were class II ** -/- embryos found at E11.5 were dead
At E9.5 the appearance of homozygous ERK5 knockout embryos was similar to that of the wild type. However, between E9.5 and E10.25 some differences between the knockout embryos and wild type embryos started to become apparent (Fig 2). By E10.25 knockout embryos were clearly growth retarded compared to littermate controls, and clear morphological differences could be seen. All ERK5 knockout embryos had problems in placental and blood vessel development, and in addition to this, two distinct morphologies could be seen in knockout embryos by E10.25. The first morphology, referred to as 'class I', was characterised by severe retardation of growth, especially in the head and lower trunk region. In contrast, 'class II' embryos were less growth retarded than class I embryos, however development of the head and lower trunk was abnormal. Development of the first and second branchial arches was reduced, and the embryos developed an abnormal head shape. In addition, development of the cephalic mesenchyme appeared abnormal (Fig 2C). Changes seen in the ERK5 knockout are discussed in more detail below.
Figure 2
Phenotypes of ERK5 -/- mutant embryos. Embryos were isolated from timed matings of ERK5+/- mice and genotyped by PCR analysis of the isolated yolk sac. At E9.5 (A) little difference could be observed between wild type and ERK5-/- littermates. At E9.75 (B) differences could be seen between the WT and ERK5-/- embryos in the head regions, particularly in the cephalic mesenchyme of class II embryos (red arrowhead). At E10.25 (C) ERK5-/- embryos were growth retarded compared to wild type embryos. ERK5-/- embryos showed an abnormal head shape, compared to wild type embryos (green arrow) and in class II ERK5-/- embryos also showed severe abnormalities in the cephalic mesenchyme and 1st and 2nd branchial arches (yellow arrows). Development of the hind limb buds (star) and lower trunk was also retarded in the ERK5-/- embryos.
Expression of ERK5 and MKK5 during embryogenesis
Analysis of the expression of ERK5, and its upstream kinase MKK5, by whole mount in situ hybridisation using antisense RNA probes showed that the expression of these kinases was dynamically regulated during embryonic development (Fig 3A,3B,3C,3D,3E). At E8.5 ERK5 expression was low and occurred mainly in the cephalic neural fold and primitive gut. At E9.5 ERK5 expression was seen in the first and second branchial arch, cephalic region, somites and lateral ridge along the body wall. By E10.5 and 11.5 ERK5 expression was also seen in the developing limb buds. As would be expected for the upstream activator of ERK5, the expression pattern of MKK5 was found to be similar to that of ERK5 from E8.5 to E12.5. Interestingly, ERK5 was found not to be highly expressed in the developing heart as judged from whole mount immunosatining. To examine this further, sections of ERK5 whole mount in situ hybridisations were taken. At E 9.75 and E10.5 strong ERK5 expression was seen in the branchial arch, cephalic mesenchyme and neuropethelial regions (Fig 4A to 4C), as well as in the limbs, hind gut, septum transversum, dorsal root ganglion, somites and tail. Only weak expression of ERK5 was seen in the heart in sections at E9.5 and E9.75, however a slightly stronger expression of ERK5 could be seen in the atrial chamber of the heart at E10.5 (Fig 4). Strong expression of ERK5 was however apparent in the sinus venous below the heart. Expression of ERK5 was also examined at E10.5 in the placenta by whole mount in situ hybridisation and sectioning. ERK5 expression was found to be highest in the chorionic plate and labyrinthine layers. As protein expression does not always exactly mirror mRNA expression, E9.5 and E10.5 embryos were also dissected and ERK5 expression examined by western blotting (Fig 5A). This showed that ERK5 expression was high in the head and lower trunk of the embryo, intermediate in the heart region and low in the placenta. In adult mice, high levels of ERK5 and MKK5 were found in brain, thymus and spleen, with lower levels present in lung, stomach, adrenal gland, adipose tissue, pancreas and heart (Fig 5B).
Figure 3
Expression of ERK5 and MKK5 during embryonic development. Whole mount in situ hybridisation was carried out on wild-type embryos as described in the methods using antisense RNA probes against ERK5 or MKK5 or with no RNA probe. Expression of ERK5 and MKK5 was analysed at E8.5 was seen in the cephalic neural fold and primitive gut. At E9.5 expression was also seen in the branchial arch, cephalic region and somites and lateral ridge of the body wall. From E10.5, E11.5 to E12.5 expression of ERK5 and MKK5 increases with high expression seen in the branchial arch, head and limb buds.
Figure 4
Analysis of ERK5 expression in embryo sections. Whole mount in situ hybridisation was carried out on wild-type embryos (A-C) or placentas (D) as described in the methods using antisense RNA probes against ERK5 at E9.5, E9.75 and E10.5. After staining embryos were sectioned on a vibrotone. Strong ERK5 expression was seen in the cephalic mesenchyme (star) branchial arch and neuroepithelium. Weak expression was seen in the heart at E9.5, however this increases by E10.25 (diamond). Strong ERK5 expression was seen in the sinus venous below the heart (arrow). In the placenta (D) strongest expression was seen in the chorionic plate and labyrinthine layers.
Figure 5
Immunoblotting of ERK5 and MKK5. Wild type embryos (E9.5 and 10.5) were dissected and head, heart, gut and placenta were isolated. The placenta was then subdivided into upper (mainly embryonic) and lower (mainly maternal) regions. Whole wild type and ERK5-/- embryos were also isolated at E9.5. Tissues were also isolated from adult wild type mice. Samples were homogenised, insoluble material removed by centrifugation, and the concentration of soluble protein in the extract determined by a Bradford assay. Soluble protein (30 μg) were run on 4–14% acrylamide gels. Immunoblotting was then carried out using antibodies which recognise ERK5 or MKK5 for both embryonic (A) and adult (B) samples. Levels of ERK1/2, p38 and actin were also determined in the adult tissue samples (B).
ERK5 is required for normal angiogenesis and placental development
One of the most apparent problems in the ERK5 knockout embryos was a defect in the formation of blood vessels in the yolk sac. At E9.5 blood islands could be seen in the membranes of both WT and knockout embryos. However, by E10.25 the ERK5 knockout embryos failed to develop the highly branched network of blood vessels seen in the WT or heterozygous embryos (Fig 6). We therefore also examined blood vessel formation in the knockout embryos. At E9.75 CD31 staining of endothelial cells showed little difference between wild type and ERK5-/- embryos, and a clearly defined network of large blood vessels could be seen in both genotypes. In the wild type embryos these blood vessels continued to develop, giving rise to large blood vessels which branched down into networks of smaller vessels. This network of vesicles was especially apparent in the head regions of the embryo. In contrast much less branching of the blood vessels was apparent in the head region of ERK5-/- embryos. It should however be noted that the formation of other head structures, as well as blood vessels, was also retarded by E10.25 in the ERK5-/- embryos (Fig 7).
Figure 6
Morphology of wild type and ERK5 -/- mutant yolk sacs. Embryos were isolated from timed matings of ERK5+/- mice and photographed. Embryos were then genotyped by PCR analysis of the isolated yolk sac. At E9.75 (A) wild type and ERK5-/- mutant yolk contain blood islands (arrow). By E10.25 (B), the blood vessels found in wild type yolk sacs formed distinct large vessels which branched down into smaller vessels (arrow). In contrast, the surfaces of the ERK5-/- yolk sacs became pale, and did not show the branched blood vessels. At E11.5 (C), ERK5-/- yolk sacs appeared intermittent with diffuse patches of red blood cells (arrow). The ERK5-/- embryos showed were pale and apparently devoid of blood circulation without a beating heart.
Figure 7
CD31 whole-mount immunohistochemistry of embryos at E9.75 and E10.25. Embryos were isolated from timed mating and stained using an CD31 antibody as described in the Methods. At E9.75 (A) networks of large blood vessels were seen in the head regions of both wild type and ERK5-/- embryos (red arrow), and intersomitic vessels (blue arrow) were also apparent in both genotypes. By E10.25 however the blood vessels in the head region of wild type embryos had started to undergo angiogenesis to give rise to branched networks of smaller vessels. This was not seen in the ERK5-/- embryos (compare red arrows in B). Similarly more branching was seen in the intersomitic vessels in the wild type than ERK5-/- embryos at E10.25 (blue arrows). Results are representative of three independent experiments.
As knockouts of proteins upstream of ERK5 have been reported to cause problems in cardiac development, the development of the heart was also examined. ERK5-/- embryos underwent normal looping of the heart and were able to establish the basic heart pattern. At E9.75 however, the myocardium wall of ERK5-/- was thinner than in WT embryos, and some bleeding was seen in a proportion of ERK5-/- embryos (Fig 8).
Figure 8
Histological sections of the heart at E9.75. E9.75 wild type and ERK5-/- embryos were isolated from timed matings and TS paraffin sections were taken and stained with haematoxylin and eosin as described in the methods. Normal patterning of the heart was observed in the ERK5-/- embryos, with both atrial (At) and ventrical (V) chambers. The thickness of the atrial wall (arrow) in ERK5-/- embryos was thinner that in wild type hearts (A). The average thickness of the atrium wall was quantified from 4 wild type and 4 ERK5-/- embryos (B). ERK5-/- embryos had a significant decrease in atrial wall thickness (P < 0.01)
The cardiac defects and changes in vascular remodelling seen in the ERK5-/- embryos suggested that ERK5 may also play a role in placental development, so we therefore studied the morphology of ERK5-/- placentas at E9.75 and E10.25. Haematoxylin and eosin staining of paraffin sections from E9.75 placentas (Fig 9A and 9B) showed little difference between wild type and ERK5-/- embryos. Chorioallantoic fusion was able to occur in the absence of ERK5, and the placenta of ERK5-/- mice formed chorionic plate, labyrinth and spongiotophoblast layers. At this stage, the laryrinth of the ERK5-/- placenta resembled that of the wild type placentas, with both embryonic and maternal blood vessels present. However at E10.25 (Fig 9C and 9D) the labyrinth layer in ERK5-/- placentas was thinner than in wild type and there was less intermixing between embryonic and maternal blood vessels in the labyrinthine region. Development of trophoblast giant cells did not appear to be affected by ERK5 knockout, and staining with the spongiotrophoblast layer maker 4311 [24] suggested that this layer developed normally (Fig 10A). Staining of placental sections at E10.25 with an antibody against the cleaved form of caspase 3 showed that at E10.25 more apoptosis was occuring in the labyrinth of ERK5-/- embryos compared to wild type embryos (Fig 10B). Apoptosis was seen in endothelial cells, diploid trophoblast cells and some embryonic blood cells. No cleaved caspase 3 staining was observed in E9.75 placentas from either wild type or ERK5-/- placentas.
Figure 9
Histological sections of placenta at E9.75 and E10.25. Placentas were isolated from E9.75 and E10.25 wild type and ERK5-/- mice. Transverse paraffin sections were taken of the placentas and stained with haematoxylin and eosin as described. At E9.75, low (A) and high magnification (B) pictures of the TS sections are showed little difference between wild type and ERK5-/- embryos. At E9.75 both maternal (white arrowhead) and embryonic (green arrow) blood vessels, could be seen in both wild type and ERK5-/- placentas. At E10.25 low (C) and high magnification (D) showed that chorionic plate (CP), labyrinth (L) and spongiotrophoblast (S) layers are were present in both wild type and ERK5-/- embryos. At E10.25 intermixing of maternal and embryonic blood vessels (arrows) was seen, however in the ERK5-/- placentas many fewer maternal blood vessels were apparent in the labyrinthine layers. Scale bars are 0.1 mm and results are representative of three independent experiments.
Figure 10
4311 in situ and caspase 3 staining of E10.25 placentas. A) Wax sections of E10.25 wild type and ERK5-/- placentas were analysed by in situ hybridisation with an antisense RNA probe against 4311. The spongiotrophoblast layer is indicated by an arrow. B)Wild type and ERK5-/- placentas were isolated from E10.25 embryos, paraffin sections taken and then stained with an antibody which recognised cleaved caspase 3. Higher numbers of cleaved caspase 3 postitive cells were seen in ERK5-/- embryos compared to wild type controls. In the ERK5-/- placentas, apoptosis was seen in endothelial cells (red arrow), trophoblast cells (green arrow) and some embryonic blood cells (yellow arrow) within the labryinth. No apoptosis of giant cells was seen.
ERK5 is required for normal development of the head and lower trunk regions
By E10.25 all ERK5-/- embryos, and particularly class II embryos, showed problems with the development of the head and lower trunk regions of the embryo. Superficially, between E9.5 and E10, these differences were much less apparent, however more detailed analysis of serial sections of E9.75 embryos revealed problems in these regions in ERK5-/- embryos (Figs 9,10). The timing of this is significant, because at this developmental stage relatively little difference was seen between the vasculature and placentas of ERK5-/- and wild type embryos (Figs 6,7,8,9,10). In the head region E9.75 sections, development of the lumen was retarded in ERK5-/- embryos compared to littermates. The cephalic mesenchyme tissue was less dense with larger spaces in between the cells in ERK5-/- embryos than in wild type embryos. There was also less contact between the cephalic mesenchyme and neuroepithelial tissue in the ERK5-/- embryos (Fig 11A and 11B). This phenotype was seen in all ERK5-/- embryos examined by sectioning at E9.75 when compared to wild type littermates (n = 5 for LS sections and n = 5 for TS sections), and was seen in all of the serial sections for each ERK5-/- embryo. This defect may in part explain the abnormal head shape of the embryos (compare whole embryo pictures in Fig 2B). Problems with the cephalic mesenchyme tissue were even more apparent in class II embryos at E10.25 (Fig 2C), and an apparent absence of cephalic mesenchyme could be seen clearly in the whole embryo. To confirm this, E10.25 ERK5-/- embryos were also sectioned and the cephalic mesenchyme compared to that of wild type littermates (Fig 11C and compare to Fig 2C). In wild type embryos the cephalic mesenchyme was present, however in ERK5-/- embryos the cephalic mesenchyme was almost completely absent. In addition, the thickness of the neuroepithelial layer surrounding the area were the cephalic mesenchyme should have been was much thinner in the ERK5-/- embryos when compared to wild type controls. Analysis of TS sections of E9.75 ERK5-/- embryos showed that cephalic mesenchyme did contain major blood vessels, similar to those in wild type embryos (Fig 11D and 11E). The blood vessels in the ERK5-/- embryos were however frequently ruptured giving rise to bleeding into the mesenchyme tissue. The sites of bleeding occurred where the surrounding mesenchyme tissue was absent, suggesting that the reason for the rupturing of the vessels may have been due to the lack of support provided to the blood vessels. These results suggested that while the cephalic mesenchyme was able to form in early ERK5-/- embryos (pre E9.75), it was unable to proliferate and survive through the developmental stages from E9.75 to E10.25. We therefore used whole mount TUNEL analysis of these embryos to examine levels of apoptosis. While little apoptosis was seen in the head region of wild type embryos in E9.75 embryos, high levels of apoptosis were observed in the head of ERK5-/- embryos (Fig 12).
Figure 11
Analysis of cephalic mesenchyme. Wild type and ERK5-/- embryos were isolated from timed matings and LS or TS paraffin sections were taken and stained with haematoxylin and eosin as described in the methods. Analysis of LS sections at low (A) and high (B) magnification showed that there was less contact between the neuroepithelium and cephalic mesenchyme (black arrows) in ERK5-/- embryos, and that the cephalic mesenchyme was less dense with larger spaces between the cells in the ERK5-/- embryos (green arrows). Sections shown are representative of 5 wild type and 5 ERK5-/- embryos. LS sections were prepared from E10.25 embryos and stained with haematoxylin and eosin (C). The cephalic mesenchyme was almost completely absent in ERK5-/- embryos and the neuroepithelium was thinner. In ERK5-/- embryos at E9.75 the TS sections (D and E) through the cephalic mesenchyme again showed less mesenchymal tissue in the ERK5-/- embryos, however major blood vessels (arrows) were seen in both wild type and ERK5-/- embryos. In contrast to wild type embryos, where little bleeding was seen in the cephalic mesenchyme, in ERK5-/- embryos bleeding into the cephalic mesenchyme was frequently seen, especially where the mesenchymal tissue was absent (arrows in ERK5-/- embryos in D and E).
Figure 12
Analysis of apoptosis in the head of ERK5-/- embryos. The level of apoptosis in E9.75 embryos were analyses by whole mount TUNEL staining. These showed that there was greatly increased apoptosis in the ERK5-/- embryos compared to wild type littermate controls. (A) Fluorescent images of the head region of whole mount TUNEL stained embryos. (B) Light microscope pictures of the same regions at the same magnification. The cephalic mesenchyme is indicated by an arrowhead. Results are representative of 3 experiments.
In the lower trunk, the development of the region below the heart appeared abnormal in ERK5-/- embryos. In several embryos, the development of the septum transverum region was retarded in most ERK5-/- embryos (data not shown). Detailed analysis of transverse sections showed that while development of the foregut appeared normal, development of the mid and hind gut was not. In ERK5-/- embryos at E9.75 there appeared sites of overproliferation of cells in some areas of the hind to mid gut wall (Fig 13).
Figure 13
Analysis of hind gut at E9.75. E9.75 wild type and ERK5-/- embryos were isolated from timed matings and TS paraffin sections were taken and stained with haematoxylin and eosin as described in the methods (A). TS sections through the guts of ERK5-/- mice showed several areas were there appeared to be hyperproliferation of the gut endothelium (arrow). This was not observed in wild type embryos. Similar results were seen in 3 wild type and 3 ERK5-/- embryos.
Discussion
In this report, we show that knockout of ERK5 results in embryonic lethality at around E10.25 and show that ERK5-/- embryos have problems with placental development, changes in angiogenesis and problems with the development of the head, (especially the cephalic mesenchyme and neuroepithelium), and lower trunk of the embryo. While this work was in progress, two other groups reported ERK5 knockouts. Regan et al reported that the ERK5 knockout was lethal between E9.5 to E11.5 [25], while Sohn et al reported lethality between E10.5 and E11.5 [26]. Similar effects on placental development and angiogenesis were found in both reports, and this phenotype is consistent with the effects described here. While both Sohn et al and Regan et al reported that ERK5-/- embryos were growth retarded by E10, neither study reported characterisation of the head and trunk regions of these embryos. It is therefore not possible to say if the defects we report in the cephalic mesenchyme and gut were present in these knockouts. Differences in the targeting strageties between both Sohn et al and Regan et al, and that used here may explain why some differences were seen in the phenotypes observed, as it is not possible to rule out the possibility that truncated fragments of the ERK5 protein were expressed in any one of those knockouts, which may give rise to a dominant negative effect. Interestingly however the most severe phenotype reported was that of Regan et al, and in this study the targeting used here deleted the smallest region of the ERK5 gene of all the knockouts. It should also be stressed that other differences, such as the strain and source of mice and ES cells used, may also explain differences between the phenotypes of the three knockouts.
Interestingly we found two distinct morphologies of ERK5-/- embryos at E10.25, however the reason for this was not clear. Class I embryos were characterised by severe growth retardation compared to wild type embryos, while class II embryos were larger but had severe abnormalities in the development of the head and lower trunk. One possible explanation may relate to the degree of severity of the placental phenotype. Placental defects are a common cause of lethality at this developmental stage in knockout mice [27,28]. If the severity of these placental defects varied between individual ERK5-/- embryos due to other genetic or environmental factors, then as a result, the problems in placental development may be sufficient to kill some embryos (class I) before E10.25, but other embryos (class II) survive longer, allowing other phenotypes to become more pronounced. A similar effect has also been observed in a knockout of p38α, in which some embryos died at E11.5, presumably due to placental defects, while some embryos survived past this stage [22]. We found that the basic structures of the placenta, including the chorionic plate, labyrinthine trophoblast, labyrinth and spongiotrophoblast layers were able to form in the absence of ERK5. Histological sections did not show significant differences between ERK5-/- and wild type placenta at E9.75 (Fig 9). By E10.25 however the thickness of the labyrinth was reduced in ERK5-/- placentas, and there was less intermixing of the embryonic and maternal blood vessels in the placental labyrinth. This correlated with increased apoptosis in this region in ERK5-/- placentas (Fig 10). This is suggestive of a role for ERK5 in development of the labyrinth and chorioallantoic branching. As the labyrinth is the major site of exchange between the embryonic and maternal blood, the problem seen in the ERK5-/- placentas is likely to be sufficient to cause embryonic lethality. ERK5 is not the only MAP kinase signalling protein whose knockout affects labyrinth development. Knockout of MEKK3, an upstream activator of ERK5 and p38 [23], resulted in embryonic lethality due in part to a failure of the labyrinth development. Also knockouts of p38 and MEK1 [29] result in problems with the labyrinth, as do knockouts of several receptors known to activate MAPK signalling including LifR [30], EgfR [31], PdgfR [32,33], Met [34], and the GDP/GTP exchange factor Sos1 [35].
Consistent with the findings of Regan et al and Sohn et al, we also found that ERK5 knockout resulted in problems in angiogenesis in the embryo. Analysis of ERK5 expression by in situ hybridisation however showed that expression of ERK5 was not restricted to the developing blood vessels, but was instead expressed more widely in the embryos. Both Sohn et al and Regan et al report that expression of various signalling molecules important in angiogenesis were normal in the ERK5 knockout. The reason for the reduction in angiogenesis in the knockout mice is unclear, but may be related to general growth retardation seen in ERK5-/- embryos compared to wild type litermates at E10.25.
Knockout of ERK5 also affected cardiac development. Using both whole mount in situ hybridisation and immunoblotting of dissected embryos, we show that expression of ERK5, and its upstream activator MKK5, is expressed in the heart at E9.5 to E10.25, although their expression level was low compared to other regions of the embryo (Figs 2,3,4,5). Consistent with this, Sohn et al also reported that ERK5 expression was highest in the heart and trunk of the embryo at E9 to E9.5. Using only RNA in situ hybridisation Regan at al however reported the opposite, with high levels of ERK5 expression localised to the developing heart and little expression in the rest of the embryo. The reasons for this difference between the report of Regan et al, and both our findings and those of Sohn et al is unclear. We found development of the embryonic heart was retarded compared to wild type embryos. Similar to the report of Sohn et al, we observe that basic patterning of the heart can occur in the absence of ERK5. Once formed however, the heart does not develop past it's basic patterning. In particular the thickness of the atrial wall at E9.75 was reduced in ERK5 knockout embryos (Fig 8). Interestingly, the knockout of ERK5 had much less severe effects on heart development compared to the knockout of its potential substrate MEF2C, in which embryos die at E8.5-9 due to failure to undergo normal looping. This suggests that either MEF2C has functions which are independent of its phosphorylation by ERK5 in vivo at this developmental stage, or that other kinases such as p38 can also phosphorylate the same sites on MEF2C as ERK5 in vivo. In this respect it is interesting to note that knockout of p38 resulted in similar problems in cardiac development to the ERK5 knockout. In contrast to this report, and that of Sohn et al, Regan et al reported that the heart did not undergo normal looping at E9.5. The reason for this discrepancy is not clear, but may be due to differences in targeting or the genetic strains of mice used. The knockout of ERK5 has been previously observed to have a similar phenotype to knockouts of receptor tyrosine kinase Tie-2 [46] and its ligand Ang-1 [47], which may suggest that ERK5 could function downstream of these receptors in the heart. There is however no direct evidence to demonstrate this link and further work would be needed to establish if this were true. In isolated cell lines ERK5 has been reported to be activated by the neuregulin receptors erbB2 and erbB3 [48], raising the possibility that erk5 may mediate some to the effects of neuregulins in the heart. A further possible reason for the cardiac phenotype is that ERK5 has been reported to inhibit the activity of the VEGF promoter [26], so that increased VEGF levels in the ERK5-/- embryos may affect cardiac development. A third possibility is that the cardiac defects observed in ERK5-/- embryos may not be directly due to the lack of ERK5 in the heart, and that these phenotypes may be caused wholly or in part by stress induced by the placental defects in the knockouts. It has been shown in other knockout models that cardiac phenotypes can be secondary to other problems in the embryo (for examples see [21,36]). Further work, including the use of placental rescue or cardiac specific ERK5 knockouts, will be required to fully resolve these issues.
We also observed defects in the development of the cephalic mesenchyme and gut in the ERK5-/- embryos. In ERK5-/- embryos problems were seen in the cephalic mesenchyme from E9.75 onwards. At E9.75 the cephalic mesenchyme appeared less dense with larger spaces between the cells and less contact between the cephalic mesenchyme and the neuroepithelium. However as the embryos developed, this gradually worsened and by E10.25 the cephalic mesenchyme was essentially absent (Fig 11). Several factors suggest that the defects seen in the cephalic mesenchyme are primary phenotypes directly caused by the loss of ERK5 protein in this region. First, in stiu hybridisation showed that ERK5 was expressed in the cephalic mesenchyme from E9.75 (Fig 5). Secondly, these problems could be seen in E9.75 ERK5-/- embryos, while at this stage blood vessel and placental development appeared relatively normal in the knockouts (Fig 6,7,8,9,10), suggesting that the cephalic mesenchyme and gut defects were not secondary to a lack of angiogenesis. Consistent with this, blood vessels were present in the cephalic mesenchyme of E9.75 ERK5-/- embryos, suggesting that the problems with this tissue were not due to a lack of blood supply. The defect in the cephalic mesenchyme appeared to be due to increased apoptosis causing the tissue to be lost, rather than a problem with its initial development. Consistent with the normal initial development of this region, expression patterns of sonic hedgehog and Six3 (L. Yan, unpublished data) at E9.5 were unaffected by the knockout of ERK5. The increase in apoptosis in the ERK5-/- embryos suggests that ERK5 may be involved in regulating cell survival or poliferation. Consistent with this, overexpression of ERK5, or its upstream activator MKK5, has been shown to promote proliferation in some cell types in response to some mitogenic stimuli [1,37-39].
In summary these results are consistent with a role for ERK5 in angiogenesis and placental development, and show new functions for ERK5 in the survival of the cephalic mesenchyme and regulation of survival and apoptosis. Further work however will be required in order to determine the molecular details of these ERK5 functions.
Methods
Materials
Antibodies against ERK1/2, p38/SAPK2 and cleaved caspase 3 were from Cell Signalling. The MKK5 antibody was from Stressgen and the CD31 antibody from Pharmhigen. The ERK5 antibody has been described previously [7].
Generation of ERK5 knockouts
A genomic clone for ERK5 was obtained by screening a 129SvJ mouse BAC genomic library using a mouse ERK5 EST. Regions of the BAC corresponding to the ERK5 were subcloned by either restriction digestion or random fragmentation and sequenced. A targeting vector was designed based on this sequence to delete exons 4 to 5 of the ERK5 gene. The vector consisted of a first arm of homology (generated by cloning of a Sal I / Eco RI fragment ligated to a PCR product generated using the primers GAATTCAGATCTGTGTAAGG and AAGCTTCTGAAAATGGGAAG) then a neomycin resistance cassette, followed by a second arm of homology (generated by using the primers CATATGAGAAGAGGAAAGCCTGGGA and GCGGCCGCAGCAGGGATCAATATGT) and a thymidine kinase cassette (Fig 1). The targeting vector was linearised using Not I before transfection into mouse ES cells.
Mouse embryonic stem cells were grown and transfected as described previously ([40]), using embryonic fibroblasts from MTK-neo mice as a feeder layer. Colonies resistant to both G418 and ganciclovir were expanded and screened for correct incorporation of the ERK5 targeting vector. A probe external to the targeting vector was generated by PCR using the primers CAAGTAGGGGACCAAGTCAAC and GGCCCAATGGAAAGGCTTCTAT. This probe was used to screen DNA double digested with Hind III and Mfe I from ES cell colonies. Positive cell lines were injected into blastocysts from a C57Bl/6 × BALB/c cross, which were then reimplanted into recipient female mice [41]. Chimeric male offspring were then bred to BALB/c or C57Bl/6 mice as indicated and transmission identified by a combination of coat colour and genotyping by Southern and PCR analysis.
Routine gentoyping of the ERK5 mice was carried out by PCR on tail biopsies. PCR was carried out using the primers AACTAACCAACCCACCTTCCAAGAC and CACTAGTACTCCTACTGGCCCCGTA to identify wild type and AACTAACCAACCCACCTTCCAAGAC and ACCACCAAGCGAAACATCGCATCG to identify targeted alleles.
Isolation of embryos
Male and female mice of known genotype were placed together and time of fertilisation determined by observation of copulation plugs, and noon of that day defined as E0.5. Embryos were dissected from pregnant females at the times indicated, and the yolk sacs separated and used to genotype the embryos by PCR.
Whole mount in situ hybridisation, immunohistochemistry and TUNEL staining
Embryos were harvested and fixed in 4% paraformaldehyde. In situ hybridisation was carried out as described previously [42]. Probes for ERK5 (corresponding to the last 207 amino acid and first 165 bp of the 3' utr) and MKK5 (corresponding to the last 71 amino acid and first 295 bp of the 3' utr) were generated by PCR using the primers ACTAGTACTCCTACTGGC and GCTCAGGTGGCTGCTTAAG or ACTAGTAGGATTCGCCGGTCCTTC and ATCAGTGCTGCTGATAGGGCCTGAC respectively. PCR products were cloned into pBluescript to give antisense sequence when transcribed from the T7 promoter.
Whole mount immunohistochemical analysis of embryos using a CD31 antibody as described [43]. Whole mount terminal deoxynucleotidyl transferase-mediated UTP end labelling (TUNEL) was carried out using the in situ cell death detection kit from Roche.
Sectioning
Embryos placenta were fixed in formaldehyde, then dehydrated in ethanol, cleared in chloroform and then embedded in paraffin as described [44]. Sections were cut and stained using haematoxylin and eosin.
The atrial wall thickness was determined using a modified Cavalieri method [45]. Both the inner and outer areas of the atrial chamber were measured and the average wall thickness was defined as the difference between the average radius of the inner and outer areas of the atrial chamber. Between 6 and 9 sections were analysed per embryo, and 4 wild type and 4 ERK5-/- embryos were analysed.
Immunoblotting
Tissue was homogenised in 50 mM Tris-HCl pH 7.5, 1 mM EGTA, 1 mM EDTA, 1 mM sodium orthovanadate, 50 mM sodium fluoride, 1 mM sodium pyrophosphate, 0.27 M sucrose, 1% (v/v) Triton X-100, 0.1% (v/v) 2-mercaptoethanol and complete proteinase inhibitor cocktail (Roche). Insoluble material was removed by centrifugation at 13000 g for 5 min at 4°C. Soluble lysate (30 μg) was then run on 4–12% polyacrylamide gels (Novex, Invitrogen) and transferred onto nitrocellulose membranes. Primary antibodies against ERK1/2, p38 and MKK5 were used as described by the supplier, and the ERK5 antibody was used at 0.8 μg/ml. Secondary antibodies conjugated to horseradish peroxidase were from Pierce, and detection was performed using ECL (Amsersham).
Authors contributions
LY was involved in all aspects of this study and was responsible for most of the experimental work. JC was responsible for genotyping and management of mouse breeding, PRA assisted with analysis of the embryos, VMT was responsible for ES cell culture and blastocyst injection and CT carried out histological and caspase 3 staining. JSCA was responsible for co-ordinating the study and drafting the paper.
Acknowledgements
We would like to thank Philip Cohen for many helpful discussions and critical reading of the manuscript. We would also like to thank Janet Rossant for the 4311 clone. This research was supported by grants from the UK Medical Research Council, Astra-Zeneca, Boehringer-Ingelheim, GlaxoSmithKline, NovoNordisk and Pfizer. | [
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14691534 | Pten Dose Dictates Cancer Progression in the Prostate
Abstract
Complete inactivation of the PTEN tumor suppressor gene is extremely common in advanced cancer, including prostate cancer (CaP). However, one PTEN allele is already lost in the vast majority of CaPs at presentation. To determine the consequence of PTEN dose variations on cancer progression, we have generated by homologous recombination a hypomorphic Pten mouse mutant series with decreasing Pten activity: Ptenhy/+ > Pten+/− > Ptenhy/− (mutants in which we have rescued the embryonic lethality due to complete Pten inactivation) > Pten prostate conditional knockout (Ptenpc) mutants. In addition, we have generated and comparatively analyzed two distinct Ptenpc mutants in which Pten is inactivated focally or throughout the entire prostatic epithelium. We find that the extent of Pten inactivation dictate in an exquisite dose-dependent fashion CaP progression, its incidence, latency, and biology. The dose of Pten affects key downstream targets such as Akt, p27Kip1, mTOR, and FOXO3. Our results provide conclusive genetic support for the notion that PTEN is haploinsufficient in tumor suppression and that its dose is a key determinant in cancer progression.
Introduction
The PTEN (phosphatase and tensin homolog deleted on chromosome 10) tumor suppressor gene is located on chromosome 10q23, a genomic region frequently lost in human cancers. Somatic deletions or mutations of this gene have been identified in a large fraction of tumors, frequently in prostate cancer (CaP), placing PTEN among the most commonly mutated tumor suppressor genes in human cancer (Cantley and Neel 1999; Di Cristofano and Pandolfi 2000).
As dictated by Knudson's “two-hit” hypothesis (Knudson 1971), however, the analysis of cancer samples for mutations in PTEN has been performed searching for biallelic inactivation of the gene, which pointed at complete PTEN inactivation as a late event in cancer progression. The consequence of loss or mutation in one PTEN allele in carcinomas in situ or in primary cancers may have been underestimated. It could be postulated that if PTEN were to be haploinsufficient for some of its tumor-suppressive functions, loss of one PTEN allele or reduction in its expression may be playing a key role in tumor initiation, while further reduction of its function/expression may favor invasion and possibly tumor metastasis in advanced cancers.
In agreement with this hypothesis, it has been reported that primary tumors often show loss or alteration of at least one PTEN allele (e.g., 70%–80% of primary CaPs; Gray et al. 1998; Whang et al. 1998), while homozygous inactivation of PTEN is generally associated with advanced cancer and metastasis (Cantley and Neel 1999; Di Cristofano and Pandolfi 2000), supporting a possible key role for progressive PTEN functional loss in tumor progression.
The elucidation of the molecular basis for tumor initiation and progression in most epithelial neoplasms has been hindered by the lack of suitable laboratory and preclinical models that accurately reflect the genetic and histopathological progression of these cancers. Furthermore, the outcome of a progressive dose reduction in tumor suppressor function has been rarely assessed in vivo in the mouse. Small interfering RNA (siRNA) technology has more recently allowed testing the consequence of knockdown of a tumor suppressor such as p53 in specific cell types such as hemopoietic stem cells, by generating epi-allelic series of hypomorphs created by stable RNA interference (RNAi) transduction (Hemann et al. 2003). In the case of Pten, this analysis is further complicated since complete inactivation of the gene results in early embryonic lethality and aberrant developmental programs (Di Cristofano et al. 1998, 2001a; Suzuki et al. 1998; Podsypanina et al. 1999). Thus, a further unrestricted reduction of the Pten dose could still result in embryonic lethality. On the other hand, the consequence of complete Pten inactivation, even when restricted to a specific organ/tissue, could still affect the developmental program of that organ, while complete somatic loss of Pten in an adult organ would better approximate what is normally observed in human cancer. We have addressed these issues for Pten, as we describe in this article, through the generation of (i) hypomorphic mouse mutants and (ii) conditional mutants for complete prostate-specific Pten inactivation after puberty.
We previously reported that Pten heterozygous (Pten+/−) mutants are prone to develop neoplasms of various histological origins, including prostatic intraepithelial neoplasias (PIN) after a long latency ( >14 mo) and at incomplete penetrance (40%) (Di Cristofano et al. 2001a). Pten+/− mutants, however, never develop invasive CaPs. Invasive CaPs were, by contrast, observed in compound Pten+/−/p27Kip1+/− or Pten+/−/p27Kip1−/− mutants (Di Cristofano et al. 2001a). These lesions appeared to originate in PIN lesions, which then become invasive. However, no metastatic CaP was observed in this model, also in view of early lethality due to the occurrence of concomitant tumors of various histogenesis. Analysis of tumors, PIN, and CaP in Pten+/− mice and Pten/p27Kip1 compound mutants strongly suggested that Pten may be haploinsufficient in prostate tumor suppression since Pten protein expression was never lost in these lesions. If this was indeed the case, a further reduction in the Pten dose with respect to Pten+/− mutants should impact on tumorigenesis and tumor progression. By contrast, in case Pten needed to be completely lost for phenotypic consequences to be manifest, only a complete Pten inactivation would hasten the neoplastic process.
The discovery that phosphatidylinositol 3,4,5-trisphosphate (PIP3) is the main in vivo substrate of PTEN (Maehama and Dixon 1998) placed this phosphatase into a well-defined pathway (reviewed in Vivanco and Sawyers 2002). PIP3 levels are very low in quiescent cells, but rapidly increase upon stimulation by growth factors, through phosphoinositide 3-kinase (PI3K) activation. The role of PTEN is to keep the levels of PIP3 low by dephosphorylation at the D3 position. Loss of PTEN function results in increased PIP3 levels and subsequent Akt hyperactivation/phosphorylation (Stambolic et al. 1998; Di Cristofano et al. 1999; Backman et al. 2002; Vivanco and Sawyers 2002). It may therefore be proposed that a progressive reduction in the Pten dose could simply result in a concomitant progressive dose-dependent increase in Akt activation and its downstream molecular biological consequences. Alternatively, Pten expression levels may constitute discrete biochemical thresholds below which qualitative functional changes would occur, contributing to tumor progression and invasion.
CaP is the most common non-skin malignancy among Western adult males. It is estimated that in 2003, approximately 220,900 new cases and 28,900 CaP-related deaths will occur in the United States. Approximately 70% of CaP patients will harbor mutation or display loss of at least one allele of PTEN ensuing in the constitutive activation of the PI3K/mTOR pathway (Gray et al. 1998; Whang et al. 1998; Di Cristofano and Pandolfi 2000). As aforementioned, those mutations are in positive correlation with tumor stage and grade, PTEN being completely lost in approximately 40% of metastatic CaPs (Gray et al. 1998; Whang et al. 1998). We therefore decided to focus on the prostate as an important model system to determine genetically whether and how the dose of Pten dictates tumor initiation and progression.
We demonstrate in vivo, in a hypomorphic mouse mutant series, that Pten plays a crucial dose-dependent role in CaP tumor suppression and that Pten progressive inactivation leads to both quantitative and qualitative molecular and biological changes towards full-blown tumorigenesis.
Results
Generation of a Hypomorphic Pten Mutant Series
We have previously reported that Pten heterozygous mutants are tumor prone and viable, while complete Pten inactivation results in embryonic lethality (Di Cristofano et al. 1998, 2001a). We therefore decided to generate and characterize a mouse mutant in which the dose of Pten would be reduced further below the levels observed in a Pten+/− mutant. To this end, we at first engineered by homologous recombination an allele of Pten that would express a wild-type Pten gene at reduced levels, taking advantage of the well-known phenomenon of transcriptional interference (McDevitt et al. 1997; Morita et al. 2003). We therefore targeted within Pten intron 3 the neomycin (Neo) cassette under the control of the strong CMV promoter (Figure 1A; see also Figure 3A) in mouse embryonic stem (ES) cells. Transcription of the Neo cassette in the recombined allele was expected to interfere with the transcription of the Pten gene, in turn resulting in lower expression levels of a wild-type Pten protein by this allele. Correctly recombined ES clones were obtained (see Figure 3B; see Materials and Methods) and injected into blastocysts for germline transmission. Mutants harboring the Ptenhy allele were obtained from recombined ES cells and were found viable, thriving, and fertile. We next intercrossed Ptenhy/+ mice with Pten+/− mice to generate a hypomorphic series (Figure 1B).
Partial Rescue of Embryonic Lethality in Pten Hypomorphic Mutants
The further reduction of the Pten dose could indeed result in embryonic lethality; we therefore assessed whether compound Ptenhy/− mice would be born at all. In the analyzed cohort (n = 190), viable male and female Ptenhy/− mutants were in fact obtained. However, the frequencies among various genotypes did not follow Mendelian ratios. Out of the 25% Ptenhy/− mutants expected in these crosses, only 10% were born. In particular, approximately 40% of the Ptenhy/− males and 75% of the Ptenhy/− females were lost during gestation. This strongly suggested that a further reduction of the Pten dose had occurred in Ptenhy/− mutants and that this reduction still results in embryonic lethality, albeit at incomplete penetrance (the embryonic phenotype of the Ptenhy/− will be described elsewhere; L. C. Trotman and P. P. Pandolfi, unpublished data). Enough viable Ptenhy/− males were obtained, however, and monitored for tumorigenesis in the prostate. As predicted, a significant reduction in Pten levels and the consequent increase in Akt activation (phospho-Akt/Akt ratio) were observed in organs and primary cells from Ptenhy/− mutants when compared to Pten+/−, Ptenhy/+, and wild-type littermates. In particular, we analyzed mouse embryonic fibroblasts (MEFs), peripheral blood mononuclear cells (PBMCs), and prostates of various genotypes (Figure 1C; Figure 2A and 2B; data not shown). Semiquantitative RT–PCR analysis confirmed the reduction of Pten mRNA expression in Ptenhy/− MEFs when compared to Pten+/− and wild-type cells (see Figure 1D).
Massive Prostate Hyperplasia and Invasive CaP in Ptenhy/− Mutants
The levels of Pten were indeed further reduced in the prostates from Ptenhy/− mutants when compared to wild-type, Ptenhy/+, and Pten+/− mice (Figure 2A and 2B). Conversely, Akt activation was increased in Ptenhy/− mutants (Figure 2B). On this premise, we therefore studied prostate tumorigenesis in Ptenhy/+ (n = 26), Pten+/− (n = 24 in this study plus n = 18, as previously reported; Di Cristofano et al. 2001a), and Ptenhy/− (n = 14) mutants on a comparative basis. Mice were followed throughout their lives according to the following experimental scheme: (i) all cohorts were subjected to serial monthly (in selected cases, biweekly) nuclear magnetic resonance imaging (MRI) (see Materials and Methods) for detection of morphological changes in the size and shape of the prostate; (ii) mice were sacrificed at or prior to tumor detection for postmortem pathological analysis or when manifest sign of distress were observed (owing to tumor codevelopment; Ptenhy/− mutants, as Pten+/− mice, in fact, developed tumors of multiple histologic origins; data not shown). Striking differences were observed in the prostates of these three cohorts over time. The prostates of Pten+/+ and Pten+/− mice were never found enlarged by MRI analysis at any age (Figure 2C; see Figure 5D). Pten+/− mutants developed moderate/low-grade PINs at incomplete penetrance (35%–40% approximately, after a long latency of >12 mo) that were only detected by serial postmortem analysis, as previously reported (Figure 2F; Di Cristofano et al. 2001a). Strikingly, however, MRI analysis immediately revealed a profound difference in the Ptenhy/− cohort. The prostates of these mice were found markedly enlarged, and prostate growths were detected starting at 3 mo (see Figure 5D). By 6–8 mo, these growths typically surpassed the size of seminal vesicles (Figure 2C and 2D). Interestingly, prostate enlargements were not accompanied by an overall increase in total body weight or size in Ptenhy/− mutants (data not shown). Postmortem pathological analysis of such mutants confirmed the marked epithelial hyperplasia of the prostate (Figure 2F). All lobes were found to be hypercellular, with prominent macroscopic enlargements of anterior and dorsolateral lobes. Microscopically, the regular and round prostatic glands observed in wild-type mice were replaced by large, irregular, and complex glands that contained multiple intraluminal papillary projections already at age 2–3 mo. Furthermore, many cells had sizeable nuclei, clumped chromatin, and prominent nucleoli, forming foci of epithelial dysplasia.The phenotype observed in the Ptenhy/− mice was characterized by a wide spectrum of levels of cellular differentiation, which was not typically observed in the Pten+/− mice. As we previously reported in Pten+/− and Pten+/−/p27Kip1−/− mutants (Di Cristofano et al. 2001a), Pten protein expression was also still retained in the hyperplastic prostates from Ptenhy/− mutants, and Southern blot analysis confirmed retention of the hypomorphic Pten allele (Figure 2E; data not shown). Importantly, approximately 25% of the Ptenhy/− mice analyzed at more than 6 mo of age showed histological signs of local invasion, with tumor cells disrupting the basement membrane of the gland and growing into the surrounding stroma (Figure 2F; see Figure 5E). Thus, strikingly, a further reduction in the Pten dose as observed in Ptenhy/− mutants when compared with Pten+/− mutants does lead to massive prostate hyperplasia at complete penetrance and accelerates tumor progression from high-grade PIN (grade 3–4 tumor as per the CaP grading system for genetically engineered mice; Park et al. 2001) to locally invasive CaP in this mouse model.
Generation of Conditional Prostate-Specific Pten Mutants
We next determined whether complete Pten inactivation in the prostate would further affect CaP tumor progression. To this end, we generated mouse mutants in which Pten exons 4 and 5 are flanked by loxP sites (Figure 3A; see Materials and Methods). The Neo cassette was excised from the locus in vivo by intercrossing Ptenhy/+ with EIIA-Cre transgenic mice (Lakso et al. 1996). In these mice, the Cre recombinase is expressed transiently, early in embryogenesis, allowing the production of a mosaic progeny that harbors in the germline Pten alleles in three different configurations: targeted unmodified (still retaining the Neo cassette), Pten floxed, or Pten deleted (Figure 3A). Breeding of these mosaic mutants allowed us to generate mice heterozygous for the Pten floxed allele (PtenloxP mutants; see Figure 3B).
As expected, PtenloxP mice were born following Mendelian frequencies, viable and fertile, and were utilized for the conditional inactivation of the Pten gene in the prostate. To this end, we made use of two different Cre transgenic lines, PB-Cre and PB-Cre4, in which the Cre gene is under the control of two distinct versions of the rat Probasin (PB) gene promoter (Maddison et al. 2000; Wu et al. 2001). The PB gene is expressed in the prostatic epithelium postpuberty since the gene is androgen responsive (Matusik et al. 1986). Hence, using both PB-Cre transgenic lines, excision of the Pten gene would occur in the prostatic epithelium postpuberty. This is of relevance because it avoids any possible developmental effect due to complete inactivation of Pten during prostate organogenesis. The main difference in the two lines essentially resides in the strength of the promoter. In the PB-Cre4 line (Wu et al. 2001), the Cre gene is driven by a composite promoter, ARR2 PB, which is a derivative of the rat PB promoter from which the PB-Cre line (Maddison et al. 2000) was originally generated. PB-Cre4 mice express Cre at high levels and at high penetrance, while PB-Cre mice express Cre at lower levels and in fewer cells. This could in turn result in a more focal or more diffuse inactivation on Pten in the prostatic epithelium, as we could indeed document (Figure 4A). Also of note, PB-Cre and PB-Cre4 mice were crossed with PtenloxP mutants in order to generate PB-Cre/PtenloxP/loxP (hereafter referred to as Ptenpc1 mutants) or PB-Cre4/PtenloxP/loxP (hereafter referred to as Ptenpc2 mutants) (see Figure 3C).
The Dose of Pten Dictates Tumor Progression in the Prostate
We next studied the impact of complete Pten inactivation on prostate tumorigenesis in Ptenpc1 and Ptenpc2 mutants. Mice were followed over time exactly as described for the Pten hypomorphic series by serial MRI and pathological analysis over a period of approximately 18 mo (Figure 5D and 5E). Strikingly, we observed a dramatic difference in Ptenpc1/2 mutants when compared to Ptenhy/− mutants, but also, importantly, when comparing Ptenpc1 with Ptenpc2 mutants. MRI and macroscopic postmortem analysis of the prostate revealed a rapid massive enlargement of all the prostatic lobes already manifested from 2–3 mo of age in Ptenpc2 mutants at complete penetrance (see Figure 4B; Figure 5D). In Ptenpc1 mutants, the enlargement was less pronounced at early stages and undetectable by MRI, but still apparent (see Figure 4B; Figure 5D). As aforementioned, the pattern of Pten inactivation in the prostatic epithelium was different in Ptenpc1 versus Ptenpc2 mutants (see Figure 4A; Ptenpc1 focal versus Ptenpc2 widespread). This difference also correlated with a clear difference in both the morphology and proliferative rates of the prostatic epithelium in these two models (see Figure 4A; see also the following paragraph). Thus, Ptenpc2 mutants displayed a much more severe prostate enlargement than did Ptenpc1 mutants (see Figure 4B; Figure 5D). Ptenpc1 mice displayed focal areas of epithelial hyperplasia, with enlarged glands formed by relatively regular cells, while Ptenpc2 mice presented disorganized hyperplastic glands in all lobes with signs of cellular dysplasia, containing large, irregular cells, forming at times cryptic glandular formations (see Figure 4A).
Even more strikingly, however, pathological analysis of prostates from Ptenpc1 and Ptenpc2 mice revealed in both mutants, at complete penetrance, the development of invasive and diffuse CaP after a variable latency (see Figure 4B; Figure 5E). Tumors were in both cases made of Pten null cells (data not shown). Continuity of local invasive disease with that of intraepithelial lumens was often observed. The tumors were composed of rather large, mature epithelial cells. Neuroendocrine features, such as cytoplasmic granularity and small round cells, were not seen in the tumors analyzed (see Figure 4B). Although the tumors were clearly invasive in both Ptenpc1 and Ptenpc2 mutants with disruption of the basement membrane and clear signs of organ infiltration, once again we observed differences in the CaPs from these two models. In many instances, in Ptenpc2 mutants more than one prostate lobe was completely effaced by the infiltrating neoplastic cells, suggesting a multifocal tumor origin, while invasive tumors in Ptenpc1 mutants were often affecting only one lobe at a time and the extent of the infiltration was less diffuse (see Figure 4B). In addition, tumors in the Ptenpc1 animals were well-to-moderately differentiated while tumors in the Ptenpc2 mice were in general high-grade, undifferentiated lesions. In contrast, Ptenhy/− mutants displayed a spectrum of phenotypes, from well to undifferentiated and occasionally to locally invasive phenotypes, as previously shown.
Thus, Pten conditional inactivation in the prostate leads to invasive and diffuse CaP at complete penetrance.
Effects of the Pten Dose on the Biology of the Prostatic Epithelium
We studied the molecular and biological consequences of Pten dose variations in the prostatic epithelium of our various mutants. We included in the analysis age-matched (8-wk-old) Pten+/+, Pten+/−, Ptenhy/−, and Ptenpc2 males. The latter mutants were used instead of Ptenpc1 mice in view of the more widespread pattern of Pten inactivation observed in their prostates.
First, we determined whether reduction in the Pten dose would affect proliferation of epithelial cells. Upon Ki-67 staining, we observed a progressive increase in the proliferative rates of prostate epithelial cells, which correlated with the reduction in Pten expression levels (Figure 5A). For instance, approximately 10-fold more cells were found in active proliferation in the prostates from Ptenpc2 mutants when compared with Pten+/+ mice. Thus, Pten inactivation, in a dose-dependent manner, lends to epithelial cells a dramatic growth advantage in the prostate. By contrast, MEFs of all genotypes from the hypomorphic series did not show noticeable differences in their growth rates under standard culture conditions, while showing progressive activation of the Akt signaling pathway (data not shown; see Figure 1C). This may reflect an intrinsic differential cell-type sensitivity to Pten inactivation, also in agreement with the fact that Pten hypomorphism does not result in overall increased body size and weight.
We next assessed whether and how the progressive reduction of the Pten dose would affect the functions of key downstream targets such as Akt, p27Kip1, mTOR, and FOXO3 by immunohistochemistry (IHC) and Western blot (WB) analysis (see Materials and Methods). As aforementioned, we studied the status of Akt phosphorylation, the major known biochemical effect of Pten loss (Vivanco and Sawyers 2002), using an anti-phospho-Akt-specific antibody on cells and extracts from the various Pten mutants. Staining of prostates from various genotypes with this antibody increased proportionally to the decrease in Pten levels as confirmed by WB analysis (see Figures 2B, 5B, and 5C; data not shown). In addition, IHC analysis revealed a drastically different staining pattern in prostate cells from Ptenpc2 mutants when compared with cells from other genotypes in that phospho-Akt was found to overtly accumulate at the plasma membrane (Figure 5C). A similar staining pattern was observed both in Pten null epithelium prior to tumor occurrence and in cells from overt tumors from both Ptenpc1 and Ptenpc2 mutants (data not shown). Thus, Akt is progressively activated by a reduction in the Pten dose, which in turn results in its recruitment to the plasma membrane.
We next studied the effects of Pten inactivation on p27Kip1, FOXO3, and mTOR, known targets of Akt kinase activity. Phosphorylation of p27Kip1 and FOXO3 by Akt results in their functional inactivation through multiple mechanisms such as nuclear export, in the case of FOXO (Brunet et al. 1999), or inhibition of nuclear import and downregulation, in the case of p27Kip1 (Mamillapalli et al. 2001; Liang et al. 2002; Shin et al. 2002; Viglietto et al. 2002). By contrast, Akt activation results in mTOR phosphorylation and increased protein translation and translation initiation (Gingras et al. 2001). We previously reported that the localization or the expression levels of p27Kip1 were not affected in Pten+/− and Pten+/−/p27Kip1−/− compound mutants (Di Cristofano et al. 2001a). By contrast, a progressive downregulation in p27Kip1 staining was observed in Ptenhy/− and Ptenpc2 mutants (Figure 5C; data not shown). p27Kip1 was apparently expressed at lower levels, and fewer cells were found to express the protein (Figure 5C). We also studied the status of FOXO3 in the various Pten mutants, analyzing both its localization and threonine phosphorylation with a specific anti-phospho-FOXO3 antibody (see Materials and Methods). While no noticeable differences were observed in Pten+/− and Ptenhy/− mutants (data not shown), increase in the levels of anti-phospho-FOXO3 cytoplasmic staining were observed in prostate epithelial cells from Ptenpc2 mutants when compared to wild-type mice (Figure 5C). Complete Pten inactivation also resulted in an increase in mTOR phosphorylation in the prostatic epithelium of Ptenpc2 mutants (Figure 5C).
Thus, the progressive reduction of the Pten dose (and the extent of Pten inactivation in Ptenpc1/2 mutants) affects the proliferative rate of the prostatic epithelium and results in molecular changes, which in turn dictates the natural history of these lesions and tumor progression (Figure 5D and 5E).
Discussion
Our analysis allows a detailed deconstruction the molecular genetics underlying cancer progression in the prostate and the assessment of the key relevance of Pten and subtle variations in its dose in controlling this process. Based on our findings, we can now attribute distinct preneoplastic or malignant pathological entities to distinct molecular states. Furthermore, this new knowledge allows the reclassification, on the basis of their true molecular nature, of pathological lesions that at a superficial analysis appeared very similar (Figure 6).
In this supposedly linear and multistep process, which separates two extremely different anatomical and pathological entities, a normal prostatic epithelium from an invasive CaP, there are, in between, a discrete number of anatomically distinct intermediary steps, such as prostate hyperplasia > displasia/low-grade PIN > high-grade PIN > locally invasive CaP > diffused CaP. These entities have been recognized before on the basis of their pathological features. The fact that we are now able to correlate these anatomical stages with specific molecular events, such as the level of expression of a single tumor suppressor gene, not only represents an integration of anatomical, descriptive findings with biological data, but also offers the opportunity of therapeutic interventions.
We show in this study that inactivation of one Pten allele in the mouse may lead to prostate epithelial hyperplasia, as complete inactivation of p27Kip1 in the mouse leads to what resembles human benign prostate hyperplasia (BPH). However, these two lesions are very different in nature and outcome: the first, in fact, impacts only on epithelial elements and over time evolves to low-grade PIN, while the second impacts on both epithelial and stroma elements, representing hyperplasia of the whole organ, but does not evolve toward malignancy (Figure 6; Cordon-Cardo et al. 1998; Di Cristofano et al. 2001a). This obviously does not exclude that loss of p27Kip1 expression when accompanied by additional genetic events (e.g., loss of PTEN) may lead to a completely different outcome, as is also supported by our previous in vivo analysis in the mouse (Figure 6; Di Cristofano et al. 2001a).
Moreover, our data demonstrate that a further reduction in the Pten dose, as observed in the Ptenhy/− mutants, accelerates tumor progression dramatically, eventually resulting in high-grade PIN and locally invasive carcinoma (Figure 6). This fact has very important therapeutic implications. As a large number of CaP patients (more than 80%) display at presentation loss of one PTEN allele, but do retain the other normal PTEN allele, it could be proposed that high-grade PIN or locally invasive prostate carcinomas could be prevented or treated by pharmacologically modulating the expression of the remaining PTEN allele or by antagonizing the downstream consequences of PTEN downregulation or partial inactivation (e.g., PI3K/mTOR activation; Figure 6).
While the massive prostate hyperplasia/dysplasia observed in the Ptenhy/− mutants is not, as expected, accompanied by complete loss of Pten expression (see Figure 2E), the low penetrance of the invasive CaP in these mutants strongly suggest that additional events have to occur for this pathological transition to occur. Nevertheless, it is important to underscore that Pten+/− mice, unlike Ptenhy/− mutants, do not develop invasive CaP, but only low-grade PIN lesions (Figure 6). Thus, a further reduction of the Pten dose seems to be essential for this process. Several possibilities, not mutually exclusive, could be entertained: (i) a more severe reduction in the Pten dose could facilitate cooperative tumorigenesis by rendering the cells permissive and sensitive to the activation of additional oncogenic pathways; (ii) the massive prostate hyperplasia could facilitate the accumulation of additional genetic hits (including complete Pten loss) by simply expanding the pool of actively dividing cells; (iii) a more severe reduction in the Pten dose could protect cells from apoptotic programs that are normally triggered by the occurrence of damaging genetics events.
The fact that Ptenhy/− mutants develop massive prostate hyperplasia not accompanied by an overall increase in body weight and size underscores the fact that different tissues and organs are differentially sensitive to Pten dose variations, the prostatic epithelium being one of the most sensitive. In agreement with this notion, MEFs from Ptenhy/− mutants do not display proliferative advantage in standard culture condition over wild-type MEFs. In this respect, it is also important to remember that, unlike p27Kip1−/− mice, which display hyperplasia of both stromal and parenchymal components in the prostate (as well as an increase in body mass; Fero et al. 1996; Kiyokawa et al. 1996; Nakayama et al. 1996), in Ptenhy/− mutants it is specifically the prostatic epithelium that actively proliferates, leading to prostate hyperplasia (Figure 6). Interestingly, this does not parallel what is observed in other model systems, such as Drosophila, where dose variations in the expression of Pten or downstream signaling components (e.g., TOR) do result in variations in body size and mass (reviewed in Oldham and Hafen 2003).
While we and others had previously implicated Pten heterozygous loss in prostate tumorigenesis when in cooperation with additional oncogenic events (such as loss of p27Kip1 or Nkx3.1 or the expression of the large T oncogene; Di Cristofano et al. 2001a; Kwabi-Addo et al. 2001; Kim et al. 2002; Abate-Shen et al. 2003), we demonstrate in this article that complete inactivation of Pten alone in the prostate has already catastrophic consequences leading to invasive, diffuse, and highly aggressive malignancies (Figure 6). Although it cannot be formally excluded that complete Pten loss would still cooperate with additional oncogenic events toward full-blown transformation, this finding underscores once more the exquisite sensitivity of the prostatic epithelium to loss of Pten. This information should be factored in when tailoring the treatment of CaP or high-grade PIN lesions that have completely lost PTEN function. These lesions may be extremely sensitive to PI3K or mTOR inhibitors, while modulation of PTEN expression would not be a therapeutic option in these cases (Figure 6). On the basis of these findings, it would seem therefore of paramount importance to routinely assess the status of the PTEN gene and its expression in human CaPs and precancerous lesions as a key biomarker to opt for the most appropriate therapeutic or chemopreventive intervention modalities. It remains to be seen whether complete Pten inactivation in the mouse prostate also influences the metastatic potential of these invasive CaPs alone or in combination with additional oncogenic events. In this respect, while MRI and postmortem analyses have indeed revealed the presence of lung neoplastic lesions in our conditional Pten mutants, morphological and molecular analyses have so far excluded the metastatic nature of these tumors (M. Niki et al., unpublished data).
The stepwise reduction in the Pten dose results in clear progressively quantitative changes in the prostate epithelial cells prior to tumor development. Increased cellular proliferation correlates with increased phosphorylation of Akt. In Pten null epithelial cells, both in the preneoplastic stage or in overt CaPs, phospho-Akt is found to accumulate almost exclusively at the plasma membrane (see Figure 5C). Whether this represents the extreme consequence of Akt superactivation or rather reflects a genuine qualitative change in Akt biology and function in Pten null prostate epithelial cells remains to be determined. It also remains to be resolved whether the reduction in p27Kip1 expression is also associated with its cytoplasmic relocalization, as previously reported in human breast cancer cells suffering AKT hyperactivation (Liang et al. 2002; Shin et al. 2002; Viglietto et al. 2002). Taken together, this analysis supports a Pten dose-dependent model with progressive changes at the molecular level, arguing against a Pten threshold model for tumor suppression.
By contrast, clear qualitative morphological changes were observed in the prostate epithelium when analyzing the Pten hypomorphic series. While Pten+/− mice never display massive prostatic enlargement, further reduction to Ptenhy/− levels leads to a sharp increase in prostate mass. In addition, Pten+/− mice never develop invasive CaP, whereas Ptenhy/− mice do (Figure 6). These data clearly support a threshold model for Pten tumor suppression at the physiopathological level.
Similarly, qualitative morphological changes are also caused by the number of cells suffering complete Pten inactivation, as observed when comparing Ptenpc1 versus Ptenpc2 mice. In Ptenpc2 mice, signs of cellular dysplasia and irregular glandular formations were frequently observed, but they were mostly absent from Ptenpc1 mutants. Along the same tenet, profound mass increase was always observed at early timepoints in Ptenpc2 mice, but not in Ptenpc1 mutants. This is likely due to the marked difference in the number of Pten null cells present in the parenchyma of these two models at puberty, reflecting the different levels of Creexpression in the two models. These observations are also consistent with a threshold model for Pten tumor suppression as a function of the number of Pten null cells in a given organ.
From a biological standpoint, it will be challenging to determine in the future whether this “field effect” would also play a role in the natural history of human CaP. On the one hand, it could be speculated that human CaP likely arises from a single transformed cell and that therefore that the field effects observed in mouse models will not be critical determinants in the natural history of the human disease. On the other hand, the multistep nature of the neoplastic process (and hence the genetic heterogeneity of the lesion) and the fact that the tumor will eventually grow to form “fields” of neoplastic cells may suggest that these effects, as well as stromal and parenchymal interactions, are key aspects of the human disease that the mouse model can recapitulate.
Taken together, our findings demonstrate the key importance of Pten in CaP tumor suppression and the dramatic impact that subtle changes in Pten levels and extent of Pten inactivation may have on tumor initiation and progression in the prostate.
Materials and Methods
Mice
Generation and characterization of Pten+/− mice have been described perviously (Di Cristofano et al. 1998, 2001a). Mice were genotyped by PCR on tail DNA or by Southern blotting as shown. Hypomorphic mice were generated as described in Figure 1 and Figure 3 and tail DNA genotyped by PCR for presence of the Pten− and Ptenhy alleles (using primers 1 and 2 described below) or by Southern blotting.
Targeting vector and generation of prostate-specific Pten-deficient mice
A 129/Sv mouse genomic library (Stratagene, La Jolla, California, United States) was screened with a mouse Pten probe containing exons 4–6. To generate the targeting construct, a 4.1 Kb KpnI–BamHI fragment containing 5′ Pten genomic DNA and a 2.0 Kb XbaI fragment containing 3′ genomic DNA were cloned into pPNT. The targeting construct was linearized with NotI and electroporated into CJ7 ES cells. Transfectants were selected in G418 (350 μg/ml) and gancyclovir (2 μM) and expanded for Southern blot analysis using a 3′ probe. Chimeric mice were produced by microinjection of two independently generated targeted ES cell clones with normal karyotypes into E3.5 C57BL6/J blastocysts, then transferred to pseudopregnant foster mothers. Chimeric males were mated with C57BL6/J females (Jackson Laboratory, Bar Harbor, Maine, United States), and germline transmission of the mutant allele was verified by Southern blot analysis of tail DNA from agouti coat-colored F1 offspring. Next, PtenloxP-neo/+ mice were mated with EIIA-Cre transgenic mice (Lakso et al. 1996), and tail DNA from offspring was subjected to Southern blot analysis using probe 6.1. Through these crosses, mosaic mice harboring a Pten wild-type allele, a Pten targeted allele (PtenloxP-neo), and a floxed allele (Ptenloxp) in their germline were generated. These mosaic mutants were mated with wild-type mice and tail DNA from offspring subjected to Southern blot analysis using probe 6.1 and to PCR analysis using primer 1 (5′-AAAAGTTCCCCTGCTGATGATTTGT-3′) and primer 2 (5′-TGTTTTTGACCAATTAAAGTAGGCTGTG-3′). PCR conditions were 35 cycles (30 sec at 95°C, 1 min at 55°C, and 1 min at 72°C) using HotStarTaq Master Mix (Qiagen, Valencia, California, United States), primer (0.25 μM), and DNA (50 ng). To detect the deleted allele, primer 3 (5′-CCCCCAAGTCAATTGTTAGGTCTGT-3′) was used. PtenloxP/loxP mice were next mated with PB-Cre transgenic mice (Maddison et al. 2000) or male PB-Cre4 transgenic mice (Wu et al. 2001) for conditional prostate-specific Pten inactivation.
Autopsy and histopathology
Animals were autopsied and all tissues were examined regardless of their pathological status. Normal and tumor tissue samples were fixed in 10% buffered formalin and embedded in paraffin. Sections (5 μm) were stained with hematoxylin and eosin (H&E) according to standard protocols.
Cells
Primary MEFs derived from ten littermate embryos were produced as follows. MEFs were obtained by crossing Ptenhy/+ and Pten+/− animals, embryos were harvested at 13.5 dpc, and individual MEFs were produced and cultured as previously described (Di Cristofano et al. 2001b) and genotyped as described above. At passage 2, cells were harvested for WB analysis (see below) and mRNA extraction by the TRIZOL method (GIBCO–BRL, Carlsbad, California, United States) and cDNA produced using the SuperScript First-Strand Kit for RT–PCR (Invitrogen, Carlsbad, California, United States) according to the manufacturers' instructions. Semiquantitative PCR (yielding a 102 bp amplicon) was performed with a Pten exon 3-specific forward (5′-TGGATTCAAAGCATAAAAACCATTAC-3′) and an exon 4- and exon 5-specific reverse primer (5′-CAAAAGGATACTGTGCAACTCTGC-3′) using 30 cycles of 1 min at 95°C, 1 min at 55°C, and 1 min at 72°C) with the PCR optimizer kit (Invitrogen) and buffer A according to the manufacturer's protocols. PCR amplification of the cDNAs was performed for 25, 30, and 35 cycles, and the amplified products were found to be in the linear range at 30 cycles. PCR products from hypoxanthine phosphoribosyltransferase (HPRT) control amplification with 5′-CCTGCTGGATTACATTAAAGCACTG-3′ and 5′-GTCAAGGGCATATCCAACAACAAAC-3′ primers were used as standards (352 bp amplicon). For measuring proliferative rates of all MEF genotypes, cells were seeded in 12-well plates at 3,000 cells per well in medium containing 10% FBS and fixed at 2-d intervals. Analysis and quantification were performed as previously described (Di Cristofano et al. 2001b).
WB analysis and densitometry
Prostates from dissected animals of all genotypes were briefly homogenized (using a Polytron homogenizer; Polytron Vertrieb, Bad Wildbad, Germany), and primary MEFs were harvested and directly lysed in 50 mM Tris (pH 7.5), 150 mM NaCl, 1 mM EDTA, 0.1% NP-40, 1 mM sodium ortho-vanadate (Na3VO4), 10 mM NaF, and protease inhibitor cocktail (Hoffmann–La Roche, Basel, Switzerland) and cleared by centrifugation; concentrations were determined by the Bio-Rad Protein Assay (Bio-Rad Laboratories, Hercules, California, United States); and samples were taken into an SDS sample loading buffer.Blood was taken from the retro-orbital cavity of anesthetized mice, red cells were lysed in lysis buffer (0.155 M NH4Cl, 10 mM KHCO3, 1 mM EDTA), and intact cells harvested by centrifugation. PBMCs were lysed in lysis buffer as previously described (Di Cristofano et al. 2001b), and protein concentrations were determined as above. Equivalents of 50 μg of lysate per lane were used for SDS–PAGE and WB analysis. Proteins were detected using a rabbit polyclonal anti-Pten antibody (anti-PTEN/MMAC-1 Ab-2; NeoMarkers, Lab Vision Corporation, Fremont, California, United States), anti-actin monoclonal antibody (mAb) AC-74 (Sigma, St. Louis, Missouri, United States) for normalization, and the Phospho-Akt Pathway Sampler Kit antibodies against Akt and phospho-Serine 473 of Akt (Cell Signaling Technology, Beverly, Massachusetts, United States) following the manufacturers' instructions. After visualization with the ECL system (Amersham Biosciences, Little Chalfont, United Kingdom), films were scanned and band density was quantified using MacBas v2.5 (Fuji Photo Film Company, Tokyo, Japan) on a Macintosh computer (Apple Computer, Sunnyvale, California, United States).
IHC
Tissues were fixed in 4% paraformaldehyde and embedded in paraffin, and 8 μm-thick sections were prepared. Anti-p27 mouse mAb (catalog #610242; BD Transduction Laboratories, San Diego, California, United States) was used at 0.5 μg/ml concentration. The IHC detection was performed with the MOM kit from Vector Laboratories (Burlingame, California, United States). Rabbit polyclonal phospho-Akt (Ser 473) antibody (catalog #9270 from Cell Signaling Technology) was used in 1:50 dilution, and rabbit polyclonal antibody PTEN/MMAC1 Ab-2 (catalog #RB-072-PO from NeoMarkers) was used at 1:200 dilution. The IHC detection was performed with the ABC kit from Vector Laboratories. These staining procedures were carried out by the automated staining processor Discovery from Ventana Medical Systems (Tucson, Arizona, United States).
Anti-Ki-67 antibody (Novocastra Laboratories Ltd., Newcastle-upon-Tyne, United Kingdom), anti-Cre antibody (Novagen, Madison, Wisconsin, United States), and polyclonal anti-phospho-mTOR antibody (Cell Signaling Technology) were purchased. Anti-phospho-FOXO3 (Thr 32) was a kind gift from Anne Brunet (Harvard University, Cambridge, Massachusetts, United States). Paraffin sections (5 μm thickness) were fixed in ice-cold acetone and incubated in 0.1% H2O2 for 15 min to inactivate internal peroxidase. Antigen retrieval was performed in 10 mM citric acid for 15 min using microwave. IHC staining was performed using biotinylated secondary antibodies followed by the Vectastain ABC Elite kit (Vector Laboratories).
MRI analysis
Mice were assessed individually by MRI for tumor establishment. Images were obtained on a 1.5T General Electric LX Echo Speed Signa scanner (General Electric Medical Systems, Milwaukee, Wisconsin, United States) using homemade foil solenoid coils (22 mm or 27 mm inner diameter). The mice were anesthetized with isofluorane. Images were acquired using a two-dimensional spin-echo pulse sequence. Initially, the animal was positioned in the coil in a holder, and sagittal scout (field of view, 120 mm) images obtained (TR = 300 ms, TE = 14 ms, one excitation per phase-encoding step, 256 × 128 matrix, 3.0 mm-thick slice, 1.5 mm slice separation) to localize the region of interest. Subsequently, high-quality T2-weighted fast spin-echo transverse images were obtained (TR = 3,000–5,000 ms, TEeff = 102 ms, echo train length = 12, 6–12 excitations per phase-encoding step, 1.5 mm thick slice, field of view = 40 mm, 0.5 mm slice separation, in plane resolution of 156 ×156 μm or 156 × 208 μm).
Statistical analysis
Survival was calculated using the Kaplan–Meier method. Log rank tests were used to compare groups by using the SPSS 10 software package (SPSS Incorporated, Chicago, Illinois, United States).
Acknowledgements
We thank Peter Scardino and Howard Scher for advice and discussions; Zhenbang Chen for help with genotyping and characterization of the Pten hypomorphic mutants; Lisette Anne Maddison for the genotyping of the PR-Cre transgenic mice; Katia Manova and Craig Farrell for assistance with the automated IHC procedures; Maria Socorro Jiao for assistance with pathological analysis; and Mihaela Lupu and Cornelia Matei for assistance with the MRI analysis. This study was supported, in part, by National Cancer Institute (NCI) grants (SPORE 92629 in Prostate Cancer, MMHCC CA-84292, and RO1 CA-82328) and by the I.T. Hirschl/M. Weill–Caulier Foundation to PPP and CC-C. Imaging analysis was supported by NCI grant R24CA83084 and an MSKCC NCI Core Grant to JAK.
Abbreviations
AP - anterior prostate
BPH - benign prostatic hyperplasia
CaP - cancer of the prostate
ES - embryonic stem
H&E - hematoxylin and eosin
HPRT - hypoxanthine phosphoribosyltransferase
IHC - immunohistochemistry
mAb - monoclonal antibody
MEF - mouse embryonic fibroblast
MRI - magnetic resonance imaging
Neo - neomycin
PB - Probasin
PBMC - peripheral blood mononuclear cell
PI3K - phosphoinositide 3-kinase
PIN - prostatic intraepithelial neoplasia
PIP3 - phosphatidylinositol 3,4,5-trisphosphate
PTEN - phosphatase and tensin homolog deleted on chromosome 10
RNAi - RNA interference
siRNA - small interfering RNA
WB - Western blot
Figures and Tables
Figure 1
Production and Analysis of a Pten Hypomorphic Mouse Series
(A) Schematic representation of the wild-type (+), hypomorphic (hy), and null (−) alleles of Pten. (For a detailed view, see Figure 3A.)
(B) Breeding scheme used to produce a Pten hypomorphic mouse series and predicted hierarchy of Pten expression levels.
(C) WB analysis of MEF lysates from ten littermate primary cell cultures obtained from a single cross (indicated in [B]) confirms predicted Pten expression hierarchy in the hypomorphic series and inversely related Akt phosphorylation status (top), both verified by densitometric analysis and plotting of the Pten/actin and phospho-Akt/Akt ratios (bottom).
(D) MEF cDNA was amplified by semiquantitative PCR with exon 3 (forward) and exon 4–5 spanning (reverse) primers for Pten. Consistent with the concept of transcriptional interference at the Pten locus, Pten mRNA levels in Ptenhy/− MEFs are clearly below the level observed in heterozygosity. Lower panels show the quantification of Pten relative to Hprt amplification.
Figure 3
Conditional Knockout of the Pten Gene in Mouse Prostate
(A) Map of wild-type Pten locus (top), targeting construct (second from top), predicted targeted locus (third from top), Pten locus after Cre-mediated excision of the Neo resistance cassette by crossing with an EIIA-Cre mouse (fourth from top), and Pten locus after Cre-mediated excision of the floxed exons 4 and 5 by crossing with a PB-Cre or PB-Cre4 transgenic mouse (bottom). The genomic sequence is depicted as a black line, with black boxes representing exons 4 and 5. Pink and yellow boxes represent the Neo resistance and the HSV thymidine kinase cassette (TK) and light blue triangles represent the loxP site, respectively. The Pten genomic fragments used as a probe for Southern blot analysis are shown (3′ probe and probe 6.1). Solid arrows represent expected fragments following hybridization with the 3′ probe upon digestion with EcoRI, and dashed arrows represent expected fragments following hybridization with the probe 6.1 upon digestion with XbaI. Locations of PCR primers to detect wild-type, floxed allele (PtenloxP), and deleted allele are shown. XbaI (X), KpnI (K), BamHI (B), and EcoRI (E) sites are shown.
(B) Genotyping of mice. Lanes 1 and 2 show Southern blot analysis of ES cell clones with 3′ probe of control (lanes 2) and recombined clones and #176 (lane 1) after digestion with EcoRI, showing a 6 kb wild-type band (WT) and a 2.5 kb targeted band (R). Lanes 3 and 4 (control) represent Southern blot analysis of mouse tail DNA of offspring by crossing F1 mouse generated from #176 with an EIIA-Cre mouse with probe 6.1 after digestion with XbaI, showing wild-type, targeted (PtenloxP-neo), and floxed (PtenloxP) locus bands. Lanes 5, 6 (control), and 7 represent Southern blot analysis of mouse tail DNA of offspring by crossing the mouse of lane 3 with a wild-type mouse with probe 6.1 after digestion with XbaI. Lanes 8–14 show PCR analysis of tail DNA of offspring by crossing a PB-Cre4 (+), PtenloxP/+ male with a PtenloxP/+ female with primers 1 and 2. Lanes 8, 9, and 14 indicate PtenloxP/+ , lanes 10 and 13 indicate Pten+/+,and lanes 11 and 12 indicate PtenloxP/loxP. Lanes 15–17 show PCR analysis of tail DNA of offspring by crossing a Ptenloxp/+ male with a PB-Cre4(+), PtenloxP/+ female with primers 1, 2, and 3. Lane 15 indicates PtenloxP/+, lane 16 indicates Pten+/+, and lane 17 indicates that the deleted allele exists in tail DNA of this offspring.
(C) Representation of crossing scheme used to generate prostate-specific Pten conditional mutants. PtenloxP/loxP mutants were crossed with PB-Cre transgenic mice or PB-Cre4 transgenic mice.
Figure 2
Ptenhy/− Mice Display Massive Hyperplasia and Invasive CaP
(A) IHC on preneoplastic anterior prostate (AP) tissue of 2-mo-old littermate mice shows decreasing Pten protein levels in the hypomorphic series.
(B) WB analysis of the prostate lobes from (A) is shown on the left and their quantification is shown on the right.
(C) MRI of Ptenhy/− mice aged 6–8 mo shows pathologic structures (encircled by dashed lines) adjacent to seminal vesicles (SV) coinciding with displacement of the bladder (Bl), features typically associated with massive prostate tumors (right). These features were never found in Pten+/+ or Pten+/− mice (left).
(D) Macroscopic view of the Ptenhy/− mouse from (C) confirms massive enlargement of the AP lobes, but normal-sized seminal vesicles.
(E) Quantification of WB analysis on AP lobe total lysates from the Ptenhy/− animal shown in (C) compared to preneoplastic AP of same genotype from (A) and (B), labeled 8 mo and 2 mo, respectively. Note that in the enlarged hyperplastic prostate, Pten protein expression is retained (levels are expressed relative to the corresponding wild-type animals).
(F) Histopathology (H&E) of AP lesions at 8 mo reveals transition from low- to high-grade PIN and invasion (infiltration of stromal tissue is indicated by arrows) in Ptenhy/−, while age-matched Pten+/− tissue only shows hyperplastic features and Pten+/+ tissue is unaffected. Bars are 50 μm.
Figure 5
Molecular Effects of Loss of Pten and Biological Comparison of All Generated Mouse Models
(A) Ki-67 staining of AP lobe sections illustrates increasing proliferation with decreasing Pten levels (numbers are Ki-67-positive cells per 300 cells counted in percent; bars are 50 μm).
(B) The phospho-Akt/Akt ratio is sharply increased in the prostates of 10-wk-old Ptenpc2 animals, as shown by densitometric quantification of WB analysis.
(C) AP staining with anti-phospho-Akt antibodies reveals strong plasma membrane localization of phospho-Akt and apparent reduction of p27 protein detection, whereas phospho-mTOR and phospho-threonine-FOXO3 antibodies show increased staining in Ptenpc2 versus wild-type mouse prostates. Bars are 50 μm.
(D) Kaplan–Meier curve showing prostate enlargement as visualized by MRI. Progressive rates of mass increase for Ptenpc2 (median age, 4 mo), Ptenhy/− (median age, 7 mo), and Ptenpc1 (median age, 16 mo) mice are found. In contrast, no prostatic size irregularities were detected in Pten+/+ or Pten+/− mice.
(E) Incidence of invasive CaP. Invasive CaP was defined as tumor cells disrupting the basal membrane of prostatic glands and growing into the surrounding stroma. Full penetrance was observed in both Ptenpc1 mice as well as in Ptenpc2 mice. In contrast, Ptenhy/− mice with a follow-up of more than 6 mo displayed only 25% incidence of invasive CaP.
Figure 4
Ptenpc2 Mice Develop Invasive CaP
(A) Histopathology analysis of wild-type, Ptenpc1, and Ptenpc2 mice prior to tumor onset. H&E-stained AP (top) shows the difference in both the morphology and proliferative rates of the prostatic epithelium in these two models. IHC staining with anti-Pten antibody (bottom) was carried out on wild-type, Ptenpc1, and Ptenpc2 mice. In wild-type mice, strong cytoplasmic staining was observed in epithelial cells (arrowheads). In Ptenpc1 mice, staining was generally weak, whereas in Ptenpc2 mice, staining was completely absent in the prostatic epithelium. Original magnification, 400×.
(B) MRI (top) shows massive prostate tumor (surrounded by dashed line) in Ptenpc2 mice (at 6 mo) and no detectable difference between the prostates of Ptenpc1 or Pten+/+ mice (at 12 mo; arrowheads). Bladder (Bl) and seminal vesicles (SV) are indicated. Macroscopic view (second from top) of the same animals confirms massive enlargement of both APs in Ptenpc2 and reveals the slightly enlarged AP of Ptenpc1 mice (encircled). H&E staining (bottom) of the prostate from Ptenpc1 mice was characterized by multiple foci of PIN lesions and by the presence of prostatic adenocarcinoma. These lesions contained well-differentiated neoplastic cells and showed focal areas of invasion (arrowheads). H&E stainings of prostates from Ptenpc2 mice showed diffuse, invasive CaP with large, undifferentiated tumor cells growing into stromal areas.
Figure 6
Pten Dose Affects Prostate Tumor Progression
Pten+/− mice develop hyperplasia, dysplasia, and low-grade PIN. Ptenhy/− mice develop at complete penetrance high-grade PIN at a young age (8–10 wk) and roughly 25% present invasive CaP around 8 mo. Ptenpc mice develop invasive CaP at complete penetrance. (See Discussion for a detailed description.) p27Kip1−/− mice, on the contrary, develop only BPH. Possibilities for human therapeutic intervention derived from our findings: in addition to inactivation of PI3K/AKT and mTOR enzymatic activities (in PTEN loss of heterozygosity condition), monitoring and elevation of PTEN expression levels of the remaining allele could not only prevent formation of PIN lesions (in PTEN+/− individuals), but could importantly also be used to counteract the progression to invasive phenotypes (as observed in Ptenhy/−mouse mutants).
Footnotes
Conflicts of Interest. The authors have declared that no conflicts of interest exist.
Author Contributions. LCT, MN, ZAD, JAK, AD, AX, TVD, CC-C, and PPP conceived and designed the experiments. LCT, MN, ZAD, JAK, AD, and AX performed the experiments. LCT, MN, ZAD, JAK, AD, AX, ASBK, TVD, CC-C, and PPP analyzed the data. PR-B, NMG, TVD, and PPP contributed reagents/materials/analysis tools. LCT, MN, ZAD, and PPP wrote the paper.
Academic Editor: Nicholas Hastie, Medical Research Council Human Genetics Unit, Western General Hospital | [
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15018652 | Dppa3 / Pgc7 / stella is a maternal factor and is not required for germ cell specification in mice
Abstract
Background
In mice, germ cells are specified through signalling between layers of cells comprising the primitive embryo. The function of Dppa3 (also known as Pgc7 or stella), a gene expressed in primordial germ cells at the time of their emergence in gastrulating embryos, is unknown, but a recent study has claimed that it plays a central role in germ cell specification.
Results
To test Dppa3's role in germ cell development, we disrupted the gene in mouse embryonic stem cells and generated mutant animals. We were able to obtain viable and fertile Dppa3-deficient animals of both sexes. Examination of embryonic and adult germ cells and gonads in Dppa3-deficient animals did not reveal any defects. However, most embryos derived from Dppa3-deficient oocytes failed to develop normally beyond the four-cell stage.
Conclusion
We found that Dppa3 is an important maternal factor in the cleavage stages of mouse embryogenesis. However, it is not required for germ cell specification.
Background
Among the many specialized cell types present in adult mammals, the first to be programmed or specified during embryogenesis are germ cells, which give rise to eggs and sperm. Which molecules direct this programming of germ cells? In many other animals, including flies and worms, material known as "germ plasm" is laid down in the egg before fertilization, and its subsequent passage to a subset of embryonic cells dictates their fate as germ cells [1,2]. In mammalian embryos, germ cells are specified in a very different manner, through signalling between layers of cells comprising the primitive embryo [3,4].
Recently, Saitou, Barton and Surani proposed a molecular pathway by which these intercellular signals are translated into germ cell fate in mice [5,6]. Central to this proposed program of germ cell specification is stella / PGC7 / Dppa3, a gene expressed in primordial germ cells and their descendants, including oocytes [5,7,8]. Here we will use the name Dppa3, as approved by the Mouse Genome Informatics Database, when referring to this gene. Saitou and colleagues' model of Dppa3's role in germ cell specification was based on the timing and site of the gene's expression, not on functional analysis. Nonetheless, the model makes clear predictions as to the phenotype of mice lacking Dppa3 function: such embryos should not form germ cells. We tested this prediction and sought to clarify the gene's importance by generating Dppa3-deficient mice and examining their germline development.
Results and Discussion
We disrupted the Dppa3 gene in cultured embryonic stem (ES) cells and thereby generated Dppa3-deficient mice. Specifically, we replaced the entire open reading frame of Dppa3 in mouse V6.5 ES cells [9] with a hygromycin-thymidine kinase selection cassette flanked by loxP sites (Figure 1A). The selection cassette was subsequently removed via transient expression of Cre recombinase in targeted ES cells. The resulting heterozygous Dppa3tm1WHT/+ ES cells were used to generate chimeric mice, which transmitted the mutation to offspring. Intercrosses between Dppa3tm1WHT/+ heterozygous animals yielded Dppa3tm1WHT/Dppa3tm1WHT homozygotes as well as Dppa3tm1WHT/+ heterozygotes and +/+ offspring, demonstrating that zygotic function of Dppa3 is not essential for viability (Figure 1B). This allowed us to characterize germ cell development in animals lacking Dppa3.
Figure 1
Generation of Dppa3-deficient animals. A, Schematic representation of genomic ablation of Dppa3. The gene's four exons are shown; non-coding regions of the first and last exons are shaded gray. The hygromycin-thymidine kinase (Hygro-TK) cassette replaces the entire open reading frame of the gene. Cre-mediated excision of the selection cassette leaves only the non-coding portions of the gene, together with a single loxP site (white triangle). Also shown are the locations of genotyping primers p1, p2 and p3 in wild-type and mutated Dppa3 alleles. B, PCR genotyping of the offspring of an intercross between Dppa3tm1WHT /+ animals. Inferred genotypes are shown above the gel image. The wild type allele yields a PCR product of 304 bp with primers p1 and p2. The mutant allele (Dppa3tm1WHT) yields a PCR product of 492 bp with primers p1 and p3. M, DNA molecular weight marker.
Dppa3 is not required for germ cell specification
Our findings do not support the proposed centrality of Dppa3 in germ cell programming. First, the gonads of Dppa3tm1WHT/Dppa3tm1WHT embryos contained germ cells, identified by expression of alkaline phosphatase, in numbers comparable to those of Dppa3tm1WHT/ + and +/+ embryos (Figure 2A). Second, the ovaries of Dppa3tm1WHT/Dppa3tm1WHT adult females expressed Oct4, a marker of oocytes [10,11], despite the absence of Dppa3 expression (Figure 2B). Third, histological examination of the gonads of Dppa3tm1WHT/ Dppa3tm1WHT adults revealed no morphological defects; spermatogenesis in males and ovarian follicle development in females appeared to be normal (Figure 2C,2D). Finally, we obtained fertile Dppa3tm1WHT/Dppa3tm1WHT mice of both sexes (though litters from Dppa3tm1WHT/Dppa3tm1WHT females were small, as described below). Each of these findings demonstrates that the Dppa3 gene is not required for germ cell specification.
Figure 2
Normal germ cell development in the absence of Dppa3. A, Gonads from E12.5 embryos (above: wild type; below: Dppa3tm1WHT/Dppa3tm1WHT) stained for alkaline phosphatase to reveal primordial germ cells. B, RT-PCR analysis of gene expression in wild-type and Dppa3tm1WHT/Dppa3tm1WHT adult ovaries. C,D, Dppa3tm1WHT/Dppa3tm1WHT testis (C) and ovary (D) are histologically normal.
Moreover, this function is not readily ascribed to a gene closely related to Dppa3. We electronically searched the sequenced mouse genome for Dppa3 homologues. We identified several processed (intron-less) pseudogenes of Dppa3, but no functional, full-length homologue. As judged by RT-PCR analysis, the Dppa3 pseudogenes are not expressed in embryonic or adult tissues (data not shown).
Dppa3 is a potent maternal factor
We found that Dppa3 plays an important role in early embryonic development as a maternal factor. While Dppa3tm1WHT/Dppa3tm1WHT males were fully fertile, Dppa3tm1WHT/Dppa3tm1WHT females had small litters. This was true regardless of whether such females were crossed with Dppa3tm1WHT/Dppa3tm1WHT, Dppa3tm1WHT/+ or wild type males (3.5 ± 1.5, 3.1 ± 2.1, or 3.0 ± 0.9 viable pups/litter, respectively). By contrast, Dppa3tm1WHT/+ females of the same (mixed) genetic background had large litters when mated to Dppa3tm1WHT/Dppa3tm1WHT or Dppa3tm1WHT/+ males (9.4 ± 3.5 or 10.1 ± 3.2 viable pups/litter, respectively).
We attribute the small litters from Dppa3tm1WHT/Dppa3tm1WHT mothers to abnormalities that manifest early in embryogenesis, during the cleavage stages of pre-implantation development. While nearly all embryos derived from Dppa3-deficient oocytes developed to the 2-cell or 4-cell stage (Figure 3A,3B,3C,3D), subsequent development was severely compromised in most such embryos (Figure 3E,3F). Some embryos derived from Dppa3-deficient oocytes failed to reach the 8-cell stage and instead showed evidence of compaction at the 4-cell stage. Other embryos derived from Dppa3-deficient oocytes cleaved to form 8 to 16 blastomeres, but failed to compact (Figure 3E,3F). These observations suggest that maternally supplied Dppa3 function is important in the cleavage stages of pre-implantation development.
Figure 3
Abnormal pre-implantation development of embryos derived from Dppa3–deficient oocytes. A,C, Cultured 2-cell (A) and 4-cell (C) control embryos derived from wild-type matings. B,D, Cultured 2-cell (B) and 4-cell (D) embryos produced by crossing Dppa3tm1WHT/Dppa3tm1WHT females with wild-type males. E,F, E3.5 control embryos derived from wild-type matings have progressed to the blastocyst stage (E). By contrast, most E3.5 embryos produced by crossing Dppa3tm1WHT/Dppa3tm1WHT females with wild-type males have not progressed to the blastocyst stage and instead cleave abnormally and degenerate (F). G,H, Many embryos produced by crossing Dppa3tm1WHT/Dppa3tm1WHT females with Dppa3 +/+, Tg(Pou5f1 ΔPE-GFP)10WHT/Tg(Pou5f1 ΔPE-GFP)10WHT males fail to develop normally beyond the 4-cell stage (G) but nonetheless express the Oct4-GFP marker (H).
Might maternal Dppa3 induce zygotic expression of Oct4/Pou5f1, which encodes a transcription factor that is crucial to pre-implantation development [10,12]? To test this possibility, we crossed Dppa3tm1WHT/Dppa3tm1WHT females with Dppa3 +/+, Tg(Pou5f1 ΔPE-GFP)10WHT/Tg(Pou5f1 ΔPE-GFP)10WHT males, the latter transmitting an Oct4-GFP transgene, with the Oct4 promoter driving expression of GFP. We retrieved the resulting embryos at the 2-cell stage and cultured them in vitro for 72 hours to monitor expression of the Oct4-GFP transgene. All such embryos were observed to express the fluorescent marker, regardless of the degree to which the embryos developed or failed to develop during the culture period (Figure 3G,3H). Thus, the poor development of many embryos derived from Dppa3-deficient oocytes cannot be attributed to the absence of zygotic expression of Oct4. Further analysis of the maternal-effect phenotype of Dppa3 should illuminate the molecular and biological context and consequences of the gene's activity.
Conclusions
We conclude that Dppa3 is not required for germ cell specification in mice. The identity of the mammalian gene or genes that program germ cells remains an open question. Dppa3 appears to function as a maternal factor, with an important role early in embryogenesis, during cleavage.
Methods
Generation of Dppa3-deficient animals
The Dppa3 targeting construct contained 1.3-kb and 3-kb segments of mouse genomic DNA, the former located 5' of Dppa3's translation initiation site and the latter located 3' of the termination codon (Figure 1). At the center of the construct was a 3-kb hygromycin-thymidine kinase selection cassette (Hygro-TK) flanked by two loxP direct repeats. V6.5 (C57BL/6 × 129/Sv)F1 ES cells [9] were transfected by electroporation, and recombined clones were selected in the presence of hygromycin (Invitrogen). Correctly targeted clones were identified by long-distance genomic PCR. The Hygro-TK cassette was removed via transient transfection of ES cells with a Cre-expressing plasmid in the presence of ganciclovir (Sigma). The final genomic structure of the resulting clones was verified by Southern analysis. Two independently targeted ES cell clones were microinjected into Balb/c blastocysts to generate chimeras. Animals used in this study were of a mixed C57BL/6 × 129/Sv genetic background.
Primers for PCR genotyping were as follows: p1 (5' TAG CCT GGG GGT AGA CTC GGC TGT AT 3'); p2 (5' AAC GAG AAG AGA AGG GAG GGC TTC 3'); and p3 (5' TCA CAT AAA TCT GGA TCG TTG TGC ATC 3'). The wild type allele gives rise to a PCR product of 304 bp with primers p1 and p2. The mutant allele (Dppa3tm1WHT) gives rise to a PCR product of 492 bp with primers p1 and p3.
RNA isolation and RT-PCR
Total RNAs were isolated from mouse tissues, and expression of Dppa3, Oct4, and Gapd was assayed by RT-PCR, all as described previously[8].
Alkaline phosphatase staining of primordial germ cells
Gonads were dissected from wild type and Dppa3tm1WHT /Dppa3tm1WHT embryos on day 12.5 of gestation and stained for alkaline phosphatase as described previously [13].
Histology
Dissected adult testes and ovaries were fixed overnight in Bouin's solution, imbedded in paraffin, sectioned, and stained with hematoxylin and eosin.
Generation of Oct4-GFP transgenic animals
Mice bearing an Oct4-GFP transgene were generated by microinjection of a 14-kb Oct4Δ PE-GFP linear DNA fragment into C57Bl/6 × SJL F2 hybrid mouse eggs. This construct essentially reproduces the previously described GOF18Δ PE-lacZ construct [14] but contains a gene for enhanced green fluorescent protein (EGFP, Clontech) in place of lacZ at the ATG of Oct4. Mice bearing transgene Tg(Pou5f1 ΔPE-GFP)10WHT accurately reproduced the previously reported Oct4 expression pattern [14] and were bred to generate Tg(Pou5f1 ΔPE-GFP)10WHT /Tg(Pou5f1 ΔPE-GFP)10WHT homozygous animals.
Isolation, culture and analysis of cleavage stage embryos
2-cell embryos were flushed from oviducts at E1.5 and cultured for up to 72 hours in microdrops of KSOM (Specialty Media) under light mineral oil (Squibb) with 5% CO2 in air. E3.5 embryos were flushed from uteri.
Authors' contributions
AB conducted molecular biological, ES cell culture and embryological studies, and co-wrote the manuscript. MG carried out blastocyst injections. ML assisted in mouse and embryological studies. DP coordinated the study and co-wrote the manuscript. All authors read and approved the final manuscript.
Acknowledgements
A.B. was a Leukemia & Lymphoma Society Special Fellow. Supported by the Howard Hughes Medical Institute. | [
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15070402 | Mig12, a novel Opitz syndrome gene product partner, is expressed in the embryonic ventral midline and co-operates with Mid1 to bundle and stabilize microtubules
Abstract
Background
Opitz G/BBB syndrome is a genetic disorder characterized by developmental midline abnormalities, such as hypertelorism, cleft palate, and hypospadias. The gene responsible for the X-linked form of this disease, MID1, encodes a TRIM/RBCC protein that is anchored to the microtubules. The association of Mid1 with the cytoskeleton is regulated by dynamic phosphorylation, through the interaction with the α4 subunit of phosphatase 2A (PP2A). Mid1 acts as an E3 ubiquitin ligase, regulating PP2A degradation on microtubules.
Results
In spite of these findings, the biological role exerted by the Opitz syndrome gene product is still unclear and the presence of other potential interacting moieties in the Mid1 structure prompted us to search for additional cellular partners. Through a yeast two-hybrid screening approach, we identified a novel gene, MIG12, whose protein product interacts with Mid1. We confirmed by immunoprecipitation that this interaction occurs in vivo and that it is mediated by the Mid1 coiled-coil domain. We found that Mig12 is mainly expressed in the neuroepithelial midline, urogenital apparatus, and digits during embryonic development. Transiently expressed Mig12 is found diffusely in both nucleus and cytoplasm, although it is enriched in the microtubule-organizing center region. Consistently with this, endogenous Mig12 protein is partially detected in the polymerized tubulin fraction after microtubule stabilization. When co-transfected with Mid1, Mig12 is massively recruited to thick filamentous structures composed of tubulin. These microtubule bundles are resistant to high doses of depolymerizing agents and are composed of acetylated tubulin, thus representing stabilized microtubule arrays.
Conclusions
Our findings suggest that Mig12 co-operates with Mid1 to stabilize microtubules. Mid1-Mig12 complexes might be implicated in cellular processes that require microtubule stabilization, such as cell division and migration. Impairment in Mig12/Mid1-mediated microtubule dynamic regulation, during the development of embryonic midline, may cause the pathological signs observed in Opitz syndrome patients.
Background
Opitz syndrome (OS) is a congenital disorder affecting primarily midline structures (MIM 145410 and 300000). OS patients usually present with facial anomalies, including hypertelorism and cleft lip and palate. OS also includes laryngo-tracheo-esophageal (LTE), cardiac, and genitourinary abnormalities. These symptoms show high variability even within the same family [1-5]. OS is a heterogeneous disease with an X-linked (Xp22.3) and an autosomal locus (22q11.2) [6]. The gene responsible for the X-linked form, MID1, has been identified [7]. In male OS patients, mutations have been found scattered throughout the entire length of the MID1 gene, suggesting a loss of function mechanism at the basis of this developmental phenotype. Females carrying a mutated MID1 allele usually show only hypertelorism, likely as the result of differential X-inactivation [7-11]. Interestingly, during embryonic development the murine and avian orthologs of the MID1 gene show an expression pattern that, although not highly restricted, correlates with the tissues affected in OS. Within these tissues, the mouse and chick Mid1 transcripts are preferentially enriched in areas of active proliferation [12,13]. Recently, the chick Mid1 gene has been shown to be involved in the Sonic Hedgehog pathway during the establishment of the molecular left/right asymmetry in early embryonic avian development [14].
MID1 encodes a protein belonging to the tripartite motif family and is composed of a RING domain, two B-Box domains, a coiled-coil region, together forming the tripartite motif, followed by a fibronectin type III (FNIII) and an RFP-like domain [7,15,16]. The tripartite motif family, also known as TRIM or RBCC, comprises multi-domain-proteins involved in the definition of cellular compartments [17]. Mid1 self-interacts and forms high molecular weight complexes that are anchored to the microtubules throughout the cell cycle [18,19]. The most frequent MID1 alterations found in OS patients affect the C-terminal portion of the protein. Mutants that reproduce these mutations show an altered microtubule association [9,18,19]. The association of the wild-type protein with microtubules is dynamic and is regulated by its phosphorylation status: dephosphorylation of Mid1, upon interaction with the α4 regulatory subunit of phosphatase 2A (PP2A) [20], displaces Mid1 from microtubules [21,22]. It has also been reported that Mid1 functions as an E3 ubiquitin ligase, regulating the microtubular PP2A catalytic subunit degradation upon interaction with α4. PP2A degradation, in turn, controls the phosphorylation status of yet to be identified microtubule-associated-proteins (MAPs) [23].
We have identified a novel Mid1 interacting protein through yeast two-hybrid screening. This novel protein is expressed in the midline during development and co-operates with Mid1 to stabilize the microtubules.
Results
Identification of Mig12 as a novel Mid1 partner
To date, insights on the function of Mid1 in the cell have emerged from its interaction with the α4 subunit of phosphatase 2A (PP2A), however, the role of Mid1 in the pathogenesis of OS is still undetermined [21-24]. To get clues on possible biological function of Mid1, we searched for additional partners by screening a fibroblast two-hybrid library. MidM, a construct encompassing the C-terminal half of MID1, was used as a bait. This region, which comprises the coiled-coil, the FNIII repeats and the RFP-like domain of MID1, appears to be involved in the anchorage to microtubules [9,18,19]. We obtained 6 positive clones, three of which were of different lengths, belonging to a unique transcript. The largest fragment had an ORF of 514 bp, the shortest of 432 bp. We used BLAST against the nr and EST databases and we found perfectly matching clones covering an ORF of 546 bp. We derived the complete sequence from the deposited transcripts and amplified the entire cDNA. We performed an interaction-mating assay to confirm the binding. Both the full-length and the largest original clone obtained from the library specifically interact with the entire Mid1 protein (MidA) (Fig. 1A). We also found positive interaction with portions of the Mid1 protein: MidD (coiled-coil), MidH (RING-B-boxes-Coiled-coil) and with MidM, the construct used to screen the library. No interaction was observed with MID1 constructs that lack the coiled-coil region (MidF and MidC, Fig. 1A). The identified clone does not interact with other members of the TRIM family (TRIM19/PML, TRIM25/RFP, TRIM29/ATDC) that share structural homology with Mid1 [17] (data not shown).
Figure 1
Identification of a novel Mid1 partner. (A) Interaction-mating assay that confirms Mid1-Mig12 interaction in yeast. B42 fl, Mig12 full-length fused to the B42 activation domain; B42 or, the largest original Mig12 clone fused to the B42 activation domain; LexA Mid, constructs encompassing different MID1 domains fused to the LexA DNA binding domain: A, full-length; C, BB; D, CC; F, RFP-like; H, R-BB-CC; M, CC-FNIII-RFP-like. Both the full-length and the original Mig12 clones specifically interact with the entire Mid1 protein and with some of its truncated mutants, MidD, MidH and MidM, as shown by yeast turning blue on X-gal plates and growing on plates lacking leucine (Leu), only when galactose (Gal), and not glucose (Glu), is used as carbon source. Abbreviations: BB, B-box1 and B-box2 domains; CC, coiled-coil domain; FNIII, fibronectin type III repeat; R, RING domain. (B) Amino acid sequence of human (h) and mouse (m) MIG12 and comparison with the zebrafish G12 and the human SPOT14 proteins. Amino acids that are identical at least in the human and murine Mig12 are in bold. Conserved amino acids are indicated in gray. The human and mouse MIG12 share 90% of similarity and 88% of identity. The hMIG12 and the zebrafish protein share 56% of similarity and 46% of identity, whereas the homology with the human SPOT-14 protein is 49% and 31%, respectively. There is a gap of 25 aa that are not present in the zebrafish and SPOT14 proteins. (C) Co-immunoprecipitation experiments showing Mid1-Mig12 interaction. Western blot (WB) analysis using anti-Mid1 and anti-HA antibodies after immunoprecipitation of HEK293 cells transiently transfected with different combination of MycGFP-tagged Mid1 (MGFP-MID1) and an HA-tagged Mig12 (HA-MIG12); + and - indicate the constructs transfected in each lane. The antibodies used for the immunoprecipitations (IP) are indicated. Mid1 indicates the band corresponding to the endogenous protein. Ig, immunoglobulins. In some experiments, we detected a trace amount of MGFP-Mid1 immunoreactivity in cells transformed with only MGFP-Mid1 and immunoprecipitated with the anti-HA antibody. This signal was always much less than that seen when both tagged constructs were transfected together. (D) The same as in (C) using the MGFP-MidM, MGFP-MidH and MGFP-MidD mutant fusions, instead of the full-length protein, in the co-transfections and an anti-Myc antibody for Western blot analysis.
The full-length sequence matches with various anonymous human (hypothetical protein STRAIT11499, NM_021242; FLJ10386, AK001248) and mouse (AL671335, AK090003, and NM_026524 RIKEN) complete cDNA sequences and several ESTs in the databases. The human gene is located in Xp11.4 and is composed of two exons, one of which encompasses the entire coding region. The mouse gene is located in the A1.1 region of the X chromosome. The human (GenBank accession no. BK001260) and mouse (GenBank accession no. AY263385) cDNAs encode a 182- and a 181-residue-protein, respectively, displaying no known domains with the exception of a low score coiled-coil region at the C-terminus of the protein. This Mid1 interactor records the highest homology with the zebrafish 'Gastrulation specific protein G12' (NP_571410), a protein with unknown function [25], and with the mammalian SPOT-14 (NM_003251), a protein involved in the metabolism of fatty acids [26,27]. The novel transcript was dubbed MIG12 for Mid1 interacting G12-like protein, after the similarity with the Danio rerio protein. Figure 1B shows the alignment of the human and mouse Mig12, the zebrafish G12, and the human SPOT14 proteins.
To confirm that the two proteins also interact in vivo, we transiently transfected a MycGFP-tagged version of MID1 (MGFP-Mid1) and an HA-tagged version of MIG12 (HA-Mig12) in HEK293 cells and immunoprecipitated using either anti-Mid1 or anti-HA antibodies. Immunoprecipitation of Mid1 in the co-transfected sample pulls down the HA-Mig12 protein (right panel) and, vice versa, the immunoprecipitation of Mig12 using the anti-HA antibody pulls down the MGFP-Mid1 protein (left panel) (Fig. 1C). An unrelated polyclonal antibody and a different anti-tag monoclonal antibody (anti-FLAG) did not pull down the two proteins (data not shown), confirming the specificity of Mid1-Mig12 interaction. Moreover, Mig12 transfected alone is also pulled down by immunoprecipitation of the endogenous Mid1 protein (Fig. 1C). The interaction mating experiments suggest that the coiled-coil region of Mid1 is necessary and sufficient for the binding to Mig12. MGFP tagged versions of MidM, MidH, and MidD were co-transfected with HA-MIG12 in HEK293 cells and immunoprecipitated with either anti-Myc or anti-HA antibodies. The three constructs, all encompassing the coiled-coil region, are able to bind Mig12 further confirming that, also in vivo, this region is sufficient for Mid1-Mig12 interaction (Fig. 1D).
Mig12 is mainly expressed in the developing CNS midline
Since Mid1 is implicated in a developmental disorder, to support a physiologically relevant interaction between Mig12 and Mid1 we analyzed the mRNA expression of Mig12 during embryonic development. The Mig12 clone originally obtained from the two-hybrid screening was used as a probe to perform mRNA in situ hybridization on mouse embryos at several embryonic stages. A ubiquitous expression pattern was found both on section and in whole mount experiments from embryonic day 9.5 (E9.5) up to E11.5. At E11.5, we detected a diffuse staining in the central nervous system (CNS) and a more restricted signal in the developing limbs by whole-mount in situ hybridization (Fig. 2A, a). An even more restricted expression pattern is observed at E14.5 when high transcript levels are detected in specific compartments (Fig. 2A, b). The strongest expression is observed in the developing central nervous system and is particularly evident in the coronal sections through the hindbrain region (Fig. 2B, a–c). The signal is observed in the neuroepithelium of the cerebellar primordia (Fig. 2B, a,b), of the pons (Fig. 2B, a, b, e), and of the medulla oblongata (Fig. 2B, c). The ventricular hindbrain signal is mainly confined to the ventral midline (Fig. 2B, a, b, c). This medial expression is maintained throughout the central canal of the spinal cord extending through the floor and roof plates (Fig. 2B, d). In the telencephalon, Mig12 signal is present in the ventricular zone of the telencephalic vesicles (Fig. 2B, f). Within the nervous system, Mig12 transcript is also detected in the dorsal roots and in the trigeminal ganglia (Fig. 2A, b; 2B, d). At this stage, expression of Mig12 is also observed in several additional organs. The transcript is observed in the interdigital web in both the developing hind- and forelimbs at E11.5 (Fig. 2A, a). At E14.5, as the development of the limbs proceeds, Mig12 transcript is detected in the perichondrium of the digits (Fig. 2B, g). The other organs expressing Mig12 include the left and right thyroid lobes and the parathyroid glands (Fig. 2B, h); the phallic part of the urogenital sinus (Fig. 2B, i); the anal canal (rectum) and the epithelium lining the lumen of the bladder (data not shown). Interestingly, many of the sites that show high Mig12 levels also express the Mid1 transcript [12,13] and are affected in OS patients [5,11].
Figure 2
Mig12 expression analysis during embryonic development. (A) Whole mount in situ hybridization on E11.5 mouse embryo showing expression in the central nervous system and in the developing limbs (blue signal, a). Coronal and sagittal sections of E14.5 entire mouse embryos (white signal) (b). (B) Details of coronal (a, b, c, d, h) and sagittal (e, f, g, i) sections of E14.5 mouse embryos. Strong Mig12 expression (red signal) is observed in isthmal (a), pontine (a, b, e) and medulla oblongata (c) neuroepithelia, and it is maintained throughout the entire region of the spinal cord central canal (d). Expression is also observed in dorsal root ganglia (d). Mig12 transcript is detected in the telencephalon at the level of the ventricular zone (f). Signal is also present in other organs: in the perichondrium of the digits (g); in the thyroid (th) and parathyroid (pth) glands (h), and in the phallic part of the urogenital sinus (i). Abbreviations: CB, cerebellum; ccn, central canal neuroepithelium; drg, dorsal root ganglia; IS, isthmus; isn, isthmal neuroepithelium; M, medulla oblongata; mn, medulla oblongata neuroepithelium; P, pons; pc, perichondrium; pnn, pontine neuroepithelium; pth, parathyroid glands; SC, spinal cord; T, telencephalon; th, thyroid gland; us, urogenital sinus; vz, ventricular zone.
Mid1 recruits Mig12 on the microtubules
Transient expression of either MGFP- or HA-tagged Mig12 reveals a diffuse distribution of the protein in Cos7 as well as in other cell lines (U2OS, HeLa, NIH3T3). To exclude a tag-driven mislocalization, we also transfected a non-tagged version of Mig12: the specific anti-Mig12 antibody reveals a distribution comparable to that of the tagged versions. Mig12 is present in both the nucleus and the cytoplasm and the relative abundance in the two compartments is variable (Fig. 3A).
Figure 3
Immunofluorescence analyses reveal co-localization of Mid1 and Mig12 within the cell. (A) Immunofluorescence analysis after transient expression of MGFP-Mig12 (upper panel), HA-Mig12 (middle panel) and untagged Mig12 (lower panel) in Cos7 cells, revealing a diffuse distribution of the protein, in both the nucleus and the cytoplasm. (B) Co-expression of both Mid1 and Mig12 leads to co-localization of the two proteins in cytoplasmic bundles. Standard fluorescence microscopy shows formation of bundles only in Mid1 (left panels) and Mig12 (right panel) co-expressing cells. The arrow indicates a single transfected cell where Mid1 shows the classical distribution along normal interphase microtubules. (C) The co-localization is confirmed by confocal microscopy analysis in which HA-Mid protein is visible as a red signal and MGFP-Mig12 protein as a green signal; co-localization is indicated as a yellow signal in merged images. (D) Co-localization is also observed using the HA-Mig12 construct (middle panels) together with either a Mid1 OS truncated mutant (GFP-Mid1 1331insA) or a Mid1 mutant (GFP-MidD) retaining the coiled-coil domain, both localized in cytoplasmic bodies. No co-localization is observed when HA-TRIM19/PML protein is co-expressed with GFP-Mig12. The right panels represent the merged images.
Mid1 is associated with microtubules during the entire cell cycle [18,19]. An example of its distribution is shown in figure 3B (arrow, upper panel), where Mid1 co-localizes with the normal radial interphase microtubules. Interestingly, when co-expressed in the same cell, Mid1 and Mig12 form bundles within the cytoplasm (Fig. 3B). Mig12 usually also maintains a diffused distribution whose extent depends on its expression level. As shown in the lower panels, the observed bundles show variable thickness and shape that depend on the expression levels of the two proteins. Nevertheless, these bundles are only present when the two proteins are co-expressed. In our experimental conditions we do not observe the formation of bundles in cells transfected with only Mid1 (Fig. 3B, arrow). The co-localization of Mid1 and Mig12 within the bundles has been confirmed by confocal microscopy analysis (Fig. 3C).
We investigated the distribution of Mig12 in cells co-transfected with mutant Mid1 proteins that are not anchored to the microtubules. Mid1 C-terminal OS mutants localize to cytoplasmic bodies [9,18,19]. These mutant forms, that retain the coiled-coil region, are able to recruit Mig12 within these structures (Fig. 3D, upper panels). The same is observed using a construct that drives the expression of only the coiled-coil domain of Mid1 (Fig. 3D, middle panels). This recruitment is not observed when other TRIM proteins, that share the same domain composition of Mid1, are expressed with Mig12. This is demonstrated by co-transfections of Mig12 with TRIM19/PML (Fig. 3D, lower panels), TRIM5 or TRIM27 (data not shown). These results confirm that Mid1, through its coiled-coil domain, is able to specifically recruit Mig12 to different structures within the cell.
Since Mid1 is a microtubular protein, we asked whether the bundles observed in cells co-expressing Mig12 and Mid1 are structures of microtubular nature. Co-localization of tubulin with the bundles, in immunofluorescence experiments, demonstrates that these structures are microtubule arrays rearranged by overexpression of the two proteins and that are often present as continuous or fragmented perinuclear rings (Fig. 4A).
Figure 4
Mid1 and Mig12 co-sediment with microtubules. (A) Immunofluorescence analysis in Cos7 cells co-transfected with HA-Mid1 (left panels) and MGFPMig12 (middle panel) proteins. Coincidence of the bundles with microtubules is revealed using monoclonal antibodies against β-tubulin (right panel). These images show the different thickness and distribution of the bundles. (B) Cos7 cells were transfected with either MGFPMid and HA-Mig12 (left panel) or HA-Mig12 alone (right panel). Lysates (L) from cells were supplemented with 40 μM taxol to stabilize polymerized microtubules. After sedimentation on sucrose cushion, supernatant (S) and pellet (P) fractions were assayed for the presence of Mid1, Mig12, and tubulin using appropriate antibodies. In the co-transfection (left panel) both Mid1 and Mig12 were detected in the pellet together with the polymerized microtubules. As expected Mig12 is also present in the soluble fraction where neither Mid1 nor the tubulin are found. Mig12 is found partially associated with the polymerized tubulin fraction also in the single HA-Mig12 transfected cells (right panel). (C) Western blot analysis using the anti-Mig12 antibody reveals a 24 KDa protein in two different cell lines lysates (1, Cos7; 2, HeLa cells). To confirm specificity, incubation with the primary antibody was also performed in the presence of either the fusion protein used to immunize rabbits (GST-Mig12) or an unrelated fusion protein (GST-ur). (D) Detection of endogenous Mig12 in the polymerized microtubule fraction (+ taxol) in HeLa cells and as control in the non-treated sample (-taxol); legend as in (A). (E) Single Mig12 transfected Cos7 cells show partial localization with microtubules, particularly in the MTOC region (upper panels) and at the mitotic spindle poles (lower panels).
To confirm these data, we performed microtubule sedimentation after taxol treatment in cells co-transfected with both Mid1 and Mig12. After fractionation on a sucrose cushion, the supernatant and the pellet containing the polymerized tubulin were assayed by immunoblot for the presence of both proteins. Mig12 and Mid1 are recovered in the pellet, where tubulin is also found. Mig12, as expected, is also present in the supernatant. This result further indicates that the bundles observed in immunofluorescence experiments are of microtubular nature (Fig. 4B, left panel). A control protein that does not associate with the microtubules, spastin Δ N [28], is not present in the microtubule fraction, confirming that the presence of Mig12 in the pellet is not due to contamination during the sedimentation process (data not shown). Moreover, the presence of Mig12 in the pellet, as well as that of tubulin, is lost when the cells are not treated with the microtubule stabilization agent, taxol (data not shown). Thus, when overexpressed, Mid1 and Mig12 have the ability to rearrange interphase radial microtubules into these structures.
Interestingly, singly transfected Mig12 also partially sediments with the microtubular pellet, as expected to a lesser extent than the double transfectant (Fig. 4B, right panel). Since the affinity purified anti-Mig12 antibody we produced allows the specific detection of the endogenous protein in immunoblot experiments in cell line lysates, as shown in figure 4C, we carried out sedimentation of polymerized microtubules in HeLa cells to test the presence of endogenous Mig12 in the microtubule pellet. These results indicate that the protein, likely by interacting with endogenous Mid1 protein, is at least partially associated with microtubules (Fig. 4D). A closer look at some single transfected cells reveals indeed a partial co-localization of Mig12 with the microtubules, also in the absence of exogenous Mid1 (Fig. 4E). Some filaments are observed over the diffuse staining and in many cells enrichment of Mig12 protein in the MTOC region is evident (Fig. 4E, upper panels) as well as partial co-localization with the mitotic spindle (Fig. 4E, lower panels).
Mid1 and Mig12 induce stable microtubule bundles
To better understand the nature of these microtubule arrays, we asked what happens to the Mid1-Mig12 bundles upon disruption of the microtubular architecture. Cells were co-transfected and exposed to nocodazole, a microtubule-depolymerizing agent, for 1 hour before fixation and then analyzed by immunofluorescence. The filaments observed after overexpression of the two proteins were more resistant to the drug compared to control microtubules (Fig. 5A). In contrast, cells overexpressing only Mid1 show complete disruption of the microtubular apparatus, which is consistent with the absence of bundles (Fig. 5A, arrow). Partial disruption of the Mid1-Mig12 bundles was observed only after longer exposure to nocodazole (4 h, data not shown).
Figure 5
Mid1 and Mig12 together stabilize the microtubules. (A) Nocodazole treatment does not disrupt the Mid1/Mig12 generated bundles of tubulin, whereas it disrupts the microtubules in Mid1 single transfected cells (arrow). (B) The bundles represent stable microtubules as demonstrated by perfect coincidence with the anti-acetylated tubulin antibody signal (blue).
Modification of tubulin subunits by acetylation marks older microtubules and therefore indicates those that are more stable [29]. Specific antibodies to acetylated tubulin decorate the Mid1-Mig12 induced nocodazole-resistant bundles, thus indicating stable microtubules (Fig. 5B). The ability to stabilize the microtubules is not a characteristic of cells overexpressing Mig12 alone: in fact, treatment with nocodazole does not reveal any residual microtubular structures in these cells (data not shown).
These data suggest that Mig12 co-operates with Mid1 to stabilize microtubules. The Mid1-Mig12 microtubule-stabilizing effect might be implicated in specific processes during the development of the midline systems that are affected in Opitz syndrome patients.
Discussion
The role of the Opitz syndrome gene product, Mid1, in the pathogenesis of this human disorder is still unclear [14,24]. We now present data that support a role of Mid1 in the regulation of microtubule dynamics. We report the identification of a novel gene, MIG12, that encodes a Mid1 interacting protein. MIG12 shares high sequence homology with a zebrafish gene product, the 'gastrulation protein G12', which is expressed in a narrow window of time during D. rerio gastrulation [25]. A Mig12 paralog in mammals, SPOT14, is a nuclear protein that responds to the thyroid hormone and regulates lipid synthesis [26,27]. However, the mechanism of action for both G12 and SPOT14 is still unknown. Further, the absence of recognizable domains in its peptide sequence does not allow any a priori hypothesis on MIG12 function to be drawn.
The expression pattern of Mig12 during embryonic development is consistent with that of Mid1 [12,13]. Furthermore, this pattern overlaps with tissues whose development is defective in OS [5,9,11]. The strong expression in the midline of the developing central nervous system might be related to the neurological signs found in a high number of patients that manifest agenesis or hypoplasia of the corpus callosum and of the cerebellar vermis, and mental retardation. Moreover, expression of Mig12 in the rostral medial CNS could also be involved in the determination of proper craniofacial formation. It is well known that factors expressed in the CNS midline are implicated in resolving a single eye field into two lateral fields, an event that determines the head midline width and the face traits as reviewed in [30,31]. One of these, Sonic hedgehog (Shh), plays a crucial role in the ventral midline neural tube patterning and regulates the morphogenesis of a variety of midline and lateral organs. It is interesting to note the recent association of the Mid1 gene and the Shh pathway in the early midline and laterality specification in the chicken [14]. Interference with the correct Mig12-Mid1 pathway might be responsible for the craniofacial defects observed in OS. Expression in the embryonic urogenital and anal apparatus is also reminiscent of defects observed in OS, hypospadias and imperforate or ectopic anus. In addition, we can parallel the inter-digit Mig12 expression observed in the mouse embryos with OS manifestations, as we observed syndactyly in a MID1-mutated patient [11]. The low frequency of mutations in MID1 and the high variability of the phenotype in OS patients suggest the involvement of other genes in the OS phenotype. It is plausible that other proteins involved in the Mid1 pathway are implicated in the heterogeneity of OS (or in other syndromes showing clinical overlap with OS) and Mig12 might well be a candidate.
When Mig12 is over-expressed, it barely decorates microtubules with a signal almost imperceptible due to its diffused distribution in the cytoplasm. Accordingly, endogenous Mig12 is partially found associated with the polymerized tubulin fraction in cell lysates. Interestingly, when co-expressed with Mid1 it induces the formation of microtubule bundles. This effect is not observed when Mid1 is expressed alone. Mid1 specifically recruits Mig12 to the microtubules and the consequent induction of bundles could be explained by the propensity of both proteins, Mid1 [18] and Mig12 (CB, GM, unpublished results), to homo-interact. The formation of multimers might tether a high number of microtubule interacting moieties that, in turn, mediate and favor the association of parallel microtubule arrays. The shape and location of these microtubule bundles is variable within the cell: perinuclear rings, sub-cortical bundles and a roundish mass in the MTOC region. In some cases, we also observed fragmentation of these thick microtubular structures (CB, GM, unpublished results) that might suggest the involvement of a putative microtubule severing activity [32]. These microtubule bundles are resistant to depolymerizing agents, such as nocodazole, and are composed of acetylated tubulin and therefore represent stable microtubules. This bundling and stabilizing effect has been observed for other microtubule binding proteins, in particular microtubule-associated-proteins (MAPs) and other proteins involved in mitotic spindle organization, cytokinesis and the control of cell motility such as, PRC1, NuMA, CLASPs, and many others [33-36]. It is worth noting that recently two proteins sharing homology with the C-terminal half of Mid1, Mir1 and GLFND that have a coiled-coil-FNIII-RFP-like structure, have been shown to bundle and stabilize microtubules [37,38]. So far, we have no indications on the behavior of Mid1-Mig12 complexes during mitosis. Mid1 decorates the mitotic spindle [18] and Mig12, when transfected alone, appears to be both associated with the spindle poles and diffused within the cell. We have never observed mitotic cells overexpressing both proteins. Whether this is due to interference with the division process is still to be clarified.
The bundling effect observed in our over-expression system probably reflects a weaker and finely tuned-regulated process in physiological conditions. The shuttling of Mig12 between nucleus and cytoplasm might also be dynamically regulated and, in certain conditions, segregation in the nucleus might be necessary to prevent interference with the interphase microtubule network. Mid1 might recruit Mig12 to microtubules only when needed. It is possible that phosphorylation of Mid1 [21,22] and/or putative post-translational modifications of Mig12 might regulate their physiological association and the subsequent stabilization of the microtubule network. The ultimate aim of the regulation of microtubule stability and dynamics involving the Mid1-Mig12 pathway is still to be elucidated and may be connected to cell cycle progression or cell migration, events known to require microtubule stabilization [39]. Alteration of either process can be seen as possible causes of pathological signs in OS. Mig12, as well as Mid1, appears to be preferentially expressed in highly proliferating embryonic fields (e.g., the ventricular zone of the developing brain). Nevertheless, these are also cells that, after mitosis has been completed, are committed to migrate. The zebrafish gastrulation protein G12 is expressed in a restricted lineage characterized by extensive cell migration [25]; it is tempting to speculate that this process could be the one implicated in the pathogenesis of the Opitz syndrome.
Conclusions
We have reported the identification of a novel Opitz syndrome gene product interacting protein, Mig12, that co-operates with Mid1 to stabilize microtubules. These data are consistent with the role of Mid1 in microtubule dynamics. Mid1, in fact, controls MAP phosphorylation through the regulation of PP2A microtubular levels [23] and Mig12 may participate in this pathway. During embryonic development of midline structures, impairment in Mid1-Mig12-mediated microtubule dynamics regulation might be detrimental and lead to Opitz syndrome.
Methods
Plasmid constructs
The MID1 expression vectors MycGFP-MID1 and HA-MID1 have already been reported [18]. The MID1 deletion mutants, MidC, MidD, MidF, MidH, and MidM have been excised from HA-pCDNA3 vectors [18] and cloned EcoRI/XhoI in the two-hybrid vectors pJG4-5 and pEG202 [40]. Full-length MIG12 cDNA was generated by PCR amplification, using specific primers designed on ESTs sequences, from NIH3T3 total RNA as template. The PCR product was then cloned into EcoRI and XhoI sites in the eukaryotic expression vectors pcDNA3, pcDNA3-MGFP and pcDNA3-HA. Both Myc-GFP and HA tags are positioned at N-terminus region of MIG12 coding region. Full-length MIG12 was also cloned in the pJG4-5 two-hybrid vector fused to the B42 activation domain [40].
Yeast two-hybrid screening
The two-hybrid screening was performed using MIDM (CC-FNIII-RFP-like) cloned in pEG202 vector that contains the LexA DNA-binding domain. The bait was transformed into the yeast strain EGY48 that was subsequently transformed with an NIH3T3 cDNA library cloned into pJG4-5, containing the B42 activation domain. Transformants (5 × 106 independent clones) were seeded on plates containing either X-gal or lacking Leucine to select positive clones that have activated both LexA driven reporter genes (lacZ and LEU2). Interaction mating assay to confirm the positivity was performed using the same system and two different yeast mating types (EGY48 MAT α and EGY42 MAT a) as described [40].
Cell culture and transfection
Monkey Kidney Cos-7 cells and HEK 293T cells were cultured in Dulbecco's modified Eagle's medium, supplemented with 10% fetal bovine serum, at 37°C in a 5% CO2 atmosphere. All transfections were carried out by calcium phosphate precipitation [41]. In a typical transfection experiment 20 μg of expression vector were used per 15-cm dish. For immunofluorescence experiments, using chamber-slides (8 wells, Nunc), 0.5 μg DNA/well were transfected.
Immunoprecipitation, Immunoblot, and Antibodies
In co-immunoprecipitation experiments 4.5 × 106 HEK 293T cells per 15-cm dish were seeded. 60 h after transfection cells were collected, washed and extracted with RIPA buffer (150 mM NaCl, 1% Igepal, 0.5% DOC, 0.1% SDS, 50 mM Tris-HCl pH 8) supplemented with protease inhibitors (Roche). Extracts were sonicated and centrifuged at 10000 g for 10 min at 4°C to remove cell debris. The supernatants were immunoprecipitated with either 6 μg of anti-HA antibody, 500 μl anti-Myc (9E10) hybridoma supernatant or 8 μg anti-Mid1 polyclonal antibody (H35) [18], for 3 h at 4°C and the immuno-complexes collected with protein A-Sepharose beads for 30 min. The beads were washed six times with RIPA buffer and proteins eluted from the beads by boiling in SDS loading buffer. Proteins were separated on either 10% or 12% SDS PAGE and blotted onto PVDF membranes (Amersham). The membranes were rinsed in methanol and blocked in TTBS (20 mM Tris-HCl pH 7, 50 mM NaCl and 0.1% Tween-20), 5% dry milk. Incubation with the primary antibodies was performed using anti-c-Myc monoclonal antibody (1:5 dilution), anti-HA monoclonal antibody (Roche) (1:500 dilution) and anti-Mid1 polyclonal antibody (1:250 dilution) in TTBS, 5% dry milk. Antibody binding was detected with a secondary anti-mouse or anti-rabbit IgG coupled with horseradish peroxidase, followed by visualization with the Enhanced Chemiluminescence Kit (Amersham). A specific anti-Mig12 antiserum has been raised against a full-length Mig12 protein fused to GST and produced in bacteria. Affinity purification of the antibody was performed with the GST-Mig12 covalently attached to a CNBr-activated sepharose column using standard procedures. To perform competition experiments, 20 μg of the same protein were used to compete the binding in immunoblot analysis. As non-specific competitor, the same amount of an unrelated GST fusion protein (Mid1 RING domain) was used.
Immunofluorescence
Cos7 cells were grown on chamber-slides (8 wells, Nunc) in DMEM, 10% FBS, and transfected as described. After 36 h, cells were fixed in 4% paraformaldehyde/PBS for 10 min at room temperature, permeabilized with 0.2% Triton X-100/PBS for 30 min, blocked with normal serum for 1 h and incubated for 3 h with the primary antibodies and 1 h with the appropriate secondary antibodies. The following primary antibodies were used: protein A-purified polyclonal anti-Mid1 (1:200 dilution), monoclonal anti-β-tubulin (1:250 dilution) (Molecular Probes), monoclonal anti-HA (CA25) antibody (1:250 dilution) (Roche), monoclonal anti-acetylated tubulin (1:200 dilution) (Sigma). The following secondary antibodies were used: fluorescein isothiocyanate (FITC)-conjugated anti-rabbit antibodies alone or both tetramethylrhodamine isothiocyanate (TRITC) conjugated anti-rabbit and FITC conjugated anti-mouse-antibodies (1:100 dilution) (Dako). For confocal microscopy, Cy3-conjugated anti-mouse antibody was used (1:200 dilution) (Amersham). When indicated, nocodazole in DMSO was added at the final concentration of 40 μM for 1 h at 37°C before fixation.
Microtubule binding assay
Cells were harvested either 48 hours post-transfection (Cos7 cells) or when at 80% confluence (non-transfected HeLa cells) and lysed in PEM-DNNA buffer (80 mM PIPES pH 6.8, 1 mM EGTA, 1 mM MgCl2, 0.5 mM DTT, 150 mM NaCl, 1% Igepal) supplemented with protease inhibitors, at 4°C for 1 hr. The lysate was centrifuged at 610 g for 10 min at 4°C. Cytosol was then purified by successive centrifugations at 10,000 g for 10 min, at 21,000 g for 20 min and at 100,000 g for 1 hr at 4°C. Each supernatant was then supplemented with 2 mM GTP (Roche) and 40 μM taxol (Molecular Probes) and incubated at 37°C for 30 min. Corresponding samples without taxol were also prepared. Each sample was layered over a 15% sucrose cushion and centrifuged at 30,000 g for 30 min at 30°C to sediment polymerized microtubules. The resulting supernatants were saved and the pellets were suspended in an equal volume of sample buffer for electrophoresis and immunoblot analysis.
RNA in situ hybridization
One of the original clones obtained from the screening (540 bp fragment whose 5' corresponds to nt 113 of the MIG12 coding region) was linearized with the appropriate restriction enzymes to transcribe either sense or antisense 35S-labeled riboprobe. Mouse embryo tissue sections were prepared and RNA in situ hybridization experiments performed as previously described [42]. Autoradiographs were exposed for 2 days. Slides were then dipped in Kodak NTB2 emulsion and exposed for 14–21 days. In the micrographs red represents the hybridization signal and blue shows the nuclei stained with Hoechst 33258 dye. Whole-mount in situ hybridization was performed using the same probe and following the protocol described in [43].
Authors' contributions
CB carried out the two-hybrid screening, the RNA in situ hybridization analysis, the immunoprecipitation and immunofluorescence studies. BF produced the anti-Mig12 specific antibody and performed the microtubule sedimentation experiments. RF provided assistance in the cloning and preparation of the vectors. GM coordinated the study and wrote the paper. All authors read and approved the final manuscript.
Acknowledgements
We thank Salvatore Arbucci (IGB-ABT, Naples) and Francesca De Falco for assistance with the confocal microscopy and Alexandre Reymond and Alessia Errico for helpful suggestions. We are grateful to Graciana Diez-Roux, Elena Rugarli and Graziella Persico for a critical reading of the manuscript. This work was supported by the Italian Telethon Foundation and by Research Grant No. 1-FY00-700 from the March of Dimes Birth Defects Foundation. | [
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15238161 | Hedgehog can drive terminal differentiation of amniote slow skeletal muscle
Abstract
Background
Secreted Hedgehog (Hh) signalling molecules have profound influences on many developing and regenerating tissues. Yet in most vertebrate tissues it is unclear which Hh-responses are the direct result of Hh action on a particular cell type because Hhs frequently elicit secondary signals. In developing skeletal muscle, Hhs promote slow myogenesis in zebrafish and are involved in specification of medial muscle cells in amniote somites. However, the extent to which non-myogenic cells, myoblasts or differentiating myocytes are direct or indirect targets of Hh signalling is not known.
Results
We show that Sonic hedgehog (Shh) can act directly on cultured C2 myoblasts, driving Gli1 expression, myogenin up-regulation and terminal differentiation, even in the presence of growth factors that normally prevent differentiation. Distinct myoblasts respond differently to Shh: in some slow myosin expression is increased, whereas in others Shh simply enhances terminal differentiation. Exposure of chick wing bud cells to Shh in culture increases numbers of both muscle and non-muscle cells, yet simultaneously enhances differentiation of myoblasts. The small proportion of differentiated muscle cells expressing definitive slow myosin can be doubled by Shh. Shh over-expression in chick limb bud reduces muscle mass at early developmental stages while inducing ectopic slow muscle fibre formation. Abundant later-differentiating fibres, however, do not express extra slow myosin. Conversely, Hh loss of function in the limb bud, caused by implanting hybridoma cells expressing a functionally blocking anti-Hh antibody, reduces early slow muscle formation and differentiation, but does not prevent later slow myogenesis. Analysis of Hh knockout mice indicates that Shh promotes early somitic slow myogenesis.
Conclusions
Taken together, the data show that Hh can have direct pro-differentiative effects on myoblasts and that early-developing muscle requires Hh for normal differentiation and slow myosin expression. We propose a simple model of how direct and indirect effects of Hh regulate early limb myogenesis.
Background
Each muscle in a developing chick limb acquires a unique character from its inception [1]. Fibres form by the terminal differentiation of dividing myoblasts that elongate in particular orientations to form specific attachments to the skeleton. Simultaneously, the fibres of each muscle take on gene expression patterns characteristic of their future function. For example, those muscles destined to maintain body posture express certain isoforms of slow myosin from their inception, whereas future fast muscle regions fail to express this slow myosin [2]. It has been suggested that distinct cell lineages underlie the formation of slow and fast muscle fibres, and much evidence for myoblast heterogeneity has been obtained from studies both in vitro and in vivo [[3-7], reviewed in [8]]. Nevertheless, it is clear that for fibres to undergo differentiation at the appropriate time and place extrinsic cues must regulate muscle patterning.
Work on muscle patterning in somites over the past decade has shown that various protein factors secreted by adjacent tissues act as extrinsic signals regulating the formation and fate of myogenic cells [[9], reviewed in [10-12]]. One such factor is Sonic hedgehog (Shh), derived from the ventral midline, which is required for expression of markers of the earliest population of myogenic cells in the medial somite of both birds and mice [13-15]. These medial somitic cells contribute to the early-born muscle fibres of the myotome, but their subsequent fate is not known in amniotes [16,17]. Ventral midline Hedgehog (Hh) signals are also required for formation of the earliest muscle cells in the zebrafish embryo, the adaxial slow cells [[18,19], reviewed in [20]]. The fate of these cells is known, they generate a population of slow muscle fibres that migrate to form a layer of slow muscle that covers the lateral surface of the somite [21,22]. In all vertebrates examined, a second myogenic cell population arises in the lateral somite by a distinct Hh-independent genetic pathway in response to signals from more lateral and dorsal tissues. Signals such as FGFs, BMPs and WNTs and their antagonists are prime candidates for patterning of lateral somitic cells, at least in amniotes [reviewed in [8,9,23,24]]. Wnt proteins from dorsal tissues are also implicated in medial myogenesis [25-30].
In the somite, induction of precursor myoblast populations is occurring close in space and time to terminal differentiation of myoblasts into contractile fibres. This makes analysis of the precise effects of extrinsic signals hard to determine. For example, Shh can promote both primary myogenesis and subsequent cell survival in somitic explants and in vivo, but the precise target cell populations are unclear [13,15,31-33]. In contrast, in the limb bud myogenic induction and terminal differentiation are temporally and spatially separated. Myogenic cells of the limb derive from a population of precursors that migrates into the limb bud from the lateral somite [34-36]. These cells already express genes required for myogenesis prior to their migration [37,38]. Evidence suggests that several distinct populations of myogenic cells enter the limb bud [5,39,40]. Thus, muscle formation within the limb bud omits some of the early steps that occur in the somites. Consequently, we chose the somewhat simpler and more accessible limb bud to analyse the effects of Hh on the differentiation and patterning of muscle fibre types.
Previous work has shown that early manipulations that alter limb anteroposterior axis formation and skeletal pattern, including Shh mis-expression, change muscle and fibre type pattern in parallel [41,42]. Moreover, myoblast clones appear uncommitted to a particular character either early or late in myogenesis [43,44] indicating that local signals control fibre pattern. Nevertheless, implantation of cloned myoblasts into limbs can alter fibre pattern, although in such experiments it is difficult to rule out implantation of cells that are already undergoing terminal differentiation [45,46]. Myoblast implantation at later stages shows that local limb signals can re-programme myoblast differentiation [47,48]. It is clear that Hh expression within the limb at these later stages does not have a spatial pattern that is well correlated with formation of individual muscles or fibre types. Nevertheless, augmenting Hh signalling in a way that does not affect anteroposterior axis formation severely disrupts muscle patterning and differentiation leading to enlarged but disorganised muscles [49-51]. Conversely, a dramatic loss of muscle is observed in Shh-deficient mouse limbs [52]. Interpretation of these Hh results has differed, possibly because muscle is not the only tissue affected. Moreover, these studies do not distinguish direct effects of Hh on myogenic cells from indirect effects acting via non-myogenic cell populations in the limb. In the current paper, we use both in vitro and in vivo approaches to analyse the effect of Hh on muscle differentiation and slow fibre formation. We establish definitively using in vitro cultures that myoblasts can be directly induced to terminal differentiation by Shh. Moreover, Shh enhances slow myosin accumulation. With these in vitro results in mind, in vivo analysis of limbs with increased or decreased Hh signalling indicates that Hh is a muscle differentiation factor that promotes early slow myogenesis.
Results
Shh promotes myogenesis in limb monolayer culture
We examined Shh action on wing myogenic cells by exposing limb bud cells to Shh in monolayer cultures, in which we hoped the effects of secondary signals elicited by Shh action on non-myogenic cell populations would be minimized. Wing buds were dissociated at HH22 and grown in growth medium either in the presence or absence of Shh-containing conditioned medium. After two days (the equivalent of HH28), Shh-treated cultures are noticeably more dense (Fig. 1A). Immunohistochemical detection of desmin, a marker of myogenic cells, shows that myogenic and non-myogenic cells increase in parallel on Shh exposure, so that the proportion of myogenic cells remains unaltered (Fig. 1B). However, control cultures seeded at higher density show a reduced overall proportion of myogenic cells compared to low density control cultures, suggesting that interactions between cells in high density cultures repress myogenic cell accumulation. Nevertheless, even at high density, the proportion of myogenic cells is maintained on exposure to Shh, demonstrating an increase in desmin+ cell number (compare Fig. 1A,1B). We next examined pan-MyHC immunoreactivity, a marker of differentiated myocytes and myotubes. In all control cultures, ~50% of myogenic cells express MyHC. With Shh, this proportion increases to ~70% in low and ~85% in high density cultures, equivalent to a threefold increase in absolute numbers of differentiated cells (Fig. 1C). Thus, Shh enhances both number and terminal differentiation of chick limb bud myogenic cells, confirming results on explant cultures of limb and somite [32,49,53].
Figure 1
Shh induces terminal differentiation and slow MyHC in wing primary cultures. Dissociated cells HH22 wing bud cells were plated at low or high density and grown for two days in the absence (white bars) or presence of Shh-conditioned medium (ShhQT6 CM, black bars) or control CM (QT6 CM, grey bars) and the number of total (A), desmin-reactive (B), pan-MyHC-reactive (C) and slow MyHC-reactive (D) cells determined. Data from two low and one high density experiment are presented (mean ± SEM, derived by addition of fractional errors, numbers above columns are dishes for the numerator). A. Irrespective of plating density, the number of cells in dishes exposed to Shh was ~70% greater than that of control dishes. B. The proportion of desmin-expressing myogenic cells was unchanged by Shh, at either low or high cell density. Note that the proportion of myogenic cells declined at high density, consistent with faster proliferation of non-myogenic cells. C. The proportion of differentiated myogenic cells (defined as the fraction of nuclei within desmin-containing cytoplasm that were in MyHC-containing cytoplasm) increased with Shh, irrespective of density. D. With Shh more differentiated cells express slow MyHC (defined as the fraction of nuclei within MyHC-containing cytoplasm that were in slow MyHC-containing cytoplasm). Note that even the high density culture has more total slow MyHC expressing cells because the same proportion of an increased number of differentiated cells express slow MyHC. ** P ≤ 0.002 compared to control(s), t-test on cell numbers after correcting for change in numerator.
Shh can directly induce muscle terminal differentiation
Experiments using limb bud cells do not exclude an indirect effect of Shh, as non-myogenic cell types are present in the culture. Nor do such experiments resolve whether Shh directly promotes myoblast terminal differentiation, proliferation or both, or acts by preventing apoptosis. We next tried to isolate clones of myoblasts from early chick wing buds, but despite repeated attempts this proved impossible, probably due to the known difficulty of cloning embryonic chick myoblasts combined with the low abundance of myogenic cells in early wing buds. So the effect of Shh exposure was tested on the adult mouse-derived C2C12 myoblast cell line (Fig. 2). All three C2C12-derived myoblast lines tested [54,55] respond to Shh treatment by increased terminal differentiation (Fig. 2A,2B). No change in cell density is apparent (Fig. 2A), nor is BrdU incorporation altered (Fig. 2C). No change in TUNEL, acridine orange or Annexin V staining is ever observed (data not shown). Therefore, Shh can act directly on these myoblasts to promote terminal differentiation.
Figure 2
Shh increases C2 myoblast differentiation. C2 cells were exposed to ShhQT6 conditioned or control QT6 conditioned medium during both growth and differentiation. A. C2 myoblast lines C2/4 and C2C12 show enhanced differentiation in response to Shh. B. Quantitative comparison of enhanced differentiation of C2 myoblast lines in response to Shh calculated as the number of nuclei within MyHC-containing cytoplasm (i.e. number of differentiated myocytes). Note that the effect of Shh appears more dramatic if the proportion of nuclei in MyHC-containing cytoplasm is measured, because Shh-treated wells have the same or fewer total cells. For example, in one experiment C2/4 cells showed around 44% differentiation compared to ~6% in controls although across all experiments cell numbers were not significantly affected by Shh treatment. C. BrdU staining of C2/4 cells treated with ShhQT6 or control QT6 conditioned growth medium prior to a 2 hour BrdU pulse. D. In situ mRNA hybridisation on C2/4 myoblasts treated for one day with ShhQT6 conditioned growth medium revealed increased reactivity with a Gli1 antisense, but not sense probe, compared to myoblasts exposed to conditioned medium from control QT6 cells. E. Immunocytochemical analysis of C2/4 myoblasts similarly exposed to ShhQT6 conditioned or control QT6-conditioned growth medium using antibodies to MyoD, Myogenin and pan-MyHC.
Members of the Gli family of transcription factors are both targets and mediators of Shh signalling [56]. Thus, if C2 myoblasts respond directly to Shh they would be expected to show elevated levels of gli gene transcription. The addition of Shh up-regulates gli1 transcripts in over 50% of C2 myoblasts (Fig. 2D), but does not induce gli2 expression (data not shown). Thus, C2 cells appear to be responsive to Shh through the normal Shh response pathway.
To examine the mechanisms involved in the Shh response we determined the effect of Shh on C2 cells in growth factor-rich medium, which normally maintains C2 myoblasts in the cell cycle and inhibits terminal differentiation. Control cultures have low levels of nuclear MyoD protein, very little Myogenin and only rare MyHC-containing cells. However, after two days of Shh treatment, C2 cells show enhanced MyoD protein accumulation in many nuclei and a markedly higher frequency of groups of Myogenin immunoreactive cells, some of which also contain detectable MyHC (Fig. 2E). Shh, therefore, can drive nuclear MyoD accumulation and terminal differentiation of myoblasts even in the presence of growth factors found in serum.
Shh can directly promote slow fibre formation
When wing bud cultures are analysed for expression of slow MyHC, the vast majority of differentiated muscle cells fail to express detectable slow MyHC, just as nascent fibres do not express slow MyHC at the initiation of myogenesis in vivo (see below). In control low density cultures, a small proportion of differentiated cells (1–2%, defined as the proportion of nuclei within MyHC-containing cytoplasm that are in slow MyHC-containing cytoplasm) contain slow MyHC, detected by A4.840 immunoreactivity (Fig. 1D). Shh exposure doubles the proportion of myocyte/myotubes expressing slow MyHC to ~4%, which corresponds to a six-fold increase in absolute numbers of slow MyHC-reactive cells (Fig. 1D). However, in high density culture, slow MyHC is suppressed in controls and Shh fails to induce an increase in the proportion of cells expressing slow MyHC.
The small rise in cultured wing cells expressing slow MyHC from ~1–2% to ~4% of total myocytes suggested that a sub-population of myogenic cells might be induced to express slow MyHC by Shh. However, as with overall terminal differentiation, selective cell survival and/or indirect effects of Shh could not be ruled out. To avoid these criticisms, we again turned to C2 cells. Two lines of C2 cells, C2C12 and C2/4, express very little slow MyHC after three days differentiation. Shh exposure did not induce slow MyHC in these cells, despite their Shh responsiveness (Fig. 2 and data not shown). However, another line of C2C12 cells, designated C2X, expresses a low level of slow MyHC in 5–15% of myocytes in the absence of Shh (Fig. 3). Exposure to Shh during two days of proliferation in growth medium and a subsequent three day period in differentiation medium enhances differentiation to a similar extent to that in other C2C12 lines (Figs 2B and 3A,3B,3D). In addition, the frequency and intensity of slow MyHC-reactivity in myocytes increases dramatically to above 30% of differentiated cells (Fig. 3A,3C,3D). Both effects of Shh conditioned medium on C2X cells are blocked by addition of the anti-Hh antibody 5E1, confirming that Shh is the inducing component of the medium (Fig. 3B,3C). Purified Shh also up-regulates slow MyHC (Fig. 3C). Despite the ability of 5E1 to block the activity of exogenously applied Shh, it does not reduce baseline slow MyHC expression, suggesting that the low level of slow MyHC expressed in these cells is not due to autocrine exposure to a Hh signal (Fig. 3C). After Shh exposure, a very similar proportion of mono-, di-, tri- and tetranucleate myocytes contain slow MyHC (Fig. 3D), indicating that increased fusion is not responsible for the extra slow MyHC. We conclude that Shh can act on this murine myoblast cell line to induce both terminal differentiation and slow MyHC accumulation.
Figure 3
Shh promotes slow differentiation. A. Mouse C2X myoblasts were treated with control QT6 or ShhQT6 conditioned medium for two days growth and three days differentiation. Dual immunofluorescent analysis with pan MyHC (A4.1025, upper panels) and slow MyHC (A4.840, lower panels). Insets: anti-slow MyHC monoclonal antibodies A4.840 and BA-D5 confirm the induction of slow MyHC [61]. B, C. The number of differentiated myocytes, i.e. nuclei in MyHC-positive cytoplasm (B) and the proportion of differentiated myocytes expressing slow MyHC (C) was determined under various treatment regimes. Shh conditioned medium significantly increased (p < 0.001, t-test, n = 18 replicate wells in 3 experiments) both differentiation and the proportion of myocytes expressing slow MyHC compared to either untreated cells or control conditioned medium. This effect was blocked by addition of the 5E1 (1:300 diluted from 1.9 mg/ml) functionally-blocking anti-Shh monoclonal antibody, although basal differentiation and slow MyHC expression in control myocytes was unaffected. Purified preparations of mouse or zebrafish Shh N-terminal fragment also significantly induce differentiation and slow MyHC (p < 0.01, t test, n = 11 replicate wells in 2 experiments). D. Shh enhances differentiation and fusion of C2X myoblasts (left panel). Shh enhances slow MyHC accumulation in all classes of myotubes (right panel). Note that although the number of mononucleate myocytes is unaltered by Shh exposure, mononucleate myocytes are as efficiently induced to express slow MyHC as more mature multinucleate myotubes.
Initial slow myogenesis correlates with Hh signalling in vivo
The ability of Shh to enhance slow myogenesis in a murine cell line and a subset of chick primary myoblasts, just as it does in zebrafish, prompted us to re-examine the effects of Hh in vivo. First, we characterised developing chick wing buds with respect to slow MyHC expression using several monoclonal antibodies (Fig. 4). As we have described previously [47], myogenesis in the zeugopod (forearm region) commences at HH25 in both ventral and dorsal muscle mass (DMM) (Fig. 4B). At HH27/28, slow MyHC is detectable in both muscle masses. Two anti-slow MyHC antibodies, Na8 and A4.840, detect a subset of muscle fibres within the DMM running from the posterior of proximal DMM to the middle of the distal DMM (Fig. 4A,4B,4C,4D,4E,4F,4G,4H,4I,4J,4K). This pattern is similar to that previously reported for the product of the SMHC2 gene, a definitive marker of slow fibres in adult chickens [42,57]. By contrast, anti-slow MyHC antibody BA-D5 is expressed in most, if not all, early fibres, consistent with the reported expression pattern of SMHC1 gene product in all early primary fibres of the chick embryo (Fig. 4E,4I) [58]. At HH31, all three anti-slow MyHC antibodies react in both DMM and VMM, with a pattern approximately reciprocal with a neonatal/fast MyHC antibody N3.36, consistent with previous results using other antibodies [4]. Slow MyHC is most abundant in the posterior proximal (Fig. 4) and medial distal (data not shown) regions of the former DMM, which have split into individual muscles (Fig. 4L,4M,4N). Thus, BA-D5 immunoreactivity is lost within some muscles of the dorsal compartment, but is retained in the region that initially expressed the Na8 and A4.840-reactive MyHCs. Consequently, antibodies Na8 and/or A4.840 were used in all subsequent studies to mark definitive slow muscle.
Figure 4
Early formation of a restricted group of slow MyHC-expressing fibres in the chick forewing. A. Schematic of muscle fibre types in the wing DMM of the HH28 chick zeugopod showing the location of sections in C-H. Definitive slow (red) and non-slow (green) fibres are indicated. B. Immunohistochemistry of transverse cryosection of HH25 chick wings showing earliest differentiation of embryonic MyHC-reactive cells. No fibre type differences were detected at this stage (data not shown). C-K. Serial cryosections at proximal (C-G) and distal (H-K) positions (as indicated in panel A) of HH28 transverse cryosections stained for pan-MyHC (C,D,E), and slow MyHC-reactive BA-D5 (E,I), A4.840 (F,J) and Na8 (G,K). Proximally the definitive slow fibres are located in posterior DMM (cf D with F, C with G). Distally, slow fibres are localised to the central region of the DMM (cf H with I-K). L-N. Serial transverse cryosections of HH32 proximal zeugopod with pan-MyHC (L) revealing muscle splitting, N3.36-reactive neonatal/fast MyHC (M) and definitive A4.840-reactive slow MyHC (N). Some muscles, such as the EMU, contained both fast and slow fibres (M,N arrowheads). Other muscles, such as the superficial region of the FCU, had many slow fibres but fewer fast fibres (M,N; right-pointing arrows). Other muscles, such as the PP/PS, had few slow but numerous fast fibres (M,N; left-pointing arrows). EDC extensor digitorum communis; Anc anconeus; EMU extensor metacarpi ulnaris; EIL extensor indicis longus; EMR extensor metacarpi radialis; PS pronator superficialis; PP pronator profundus; FDP flexor digitorum profundus; UMV ulni metacarpalis ventralis; FCU flexor carpi ulnaris. O-V. In situ mRNA hybridisation for ptc1 (O-Q), gli1 (R-T) or gli2 (U,V) and subsequent MyHC staining with MF20 (P,Q,S-V) in control (O,P,R,S,U) and RCAS/Shh (Q,T,V) limbs. Wholemount in situ at HH22-26 shows ptc1 and gli1 mRNA in distal/posterior limb (O,R; dorsal views of right wing bud, anterior is up). By HH28, an additional zone of ptc1 mRNA is present proximally in the posterior DMM (O). Gli1 mRNA is also accumulating in distinct zones away from the ZPA (R). Sections of the zeugopod show ptc1 mRNA in both muscle and non-muscle regions, with restriction of ptc1 signal to regions without MyHC stain. RCAS/Shh implant up-regulates ptc1 throughout the dorsal region around the implant (Q, inset shows shh mRNA in/near DMM). Normal gli1 expression (S) is partially reciprocal to gli2 mRNA (U). Gli1 mRNA is up-regulated and gli2 reduced around a RCAS/Shh implant (T,V; inset shows shh mRNA in/near DMM). Dorsal is up and posterior to the left in panels B-N,P,Q,S-V. (u) ulna, (r) radius. Scale bar = 600 μm (A-K), 250 μm (L-N).
To examine the role of endogenous Hh signalling in wing myogenesis we analysed expression of gli1 and ptc1, a gene encoding a Hh receptor. Both genes are themselves downstream targets of Hh signalling [56]. Ptc1 and gli1 are highly expressed in posterior and distal limb regions from HH21-28, due to Shh deriving from the zone of polarizing activity (ZPA; Fig. 4O,4R). Ptc1 expression declines in wing cells as they leave the progress zone and commence histogenesis (Fig. 4O). Distal gli1 expression is more extensive but declines similarly (Fig. 4R). However, by HH27 new regions of Hh signalling arise around the posterior region of the DMM, perhaps because Indian hedgehog (Ihh) expression commences in cartilage anlage [59]. Ptc1 and gli1 are expressed in both non-myogenic tissues such as perichondrium and limb mesenchyme, and in the myogenic zone surrounding muscle fibres (Fig. 4P,4S). It is striking that the muscle region with highest ptc1 and gli1 mRNA roughly corresponds to the early slow zone, although it is also clear that Hh signalling in this region is not restricted to myogenic cells and that there are many fibres not expressing slow MyHC in the region of strong Hh signalling (Fig. 4F,4G,4O,4P,4R,4S).
Shh over-expression delays myogenesis and induces ectopic slow muscle 48 hours after grafting
To investigate the influence of Hh on wing muscle formation, we implanted pellets of chick embryo fibroblasts expressing a Shh/RCAS replication competent retroviral vector into the dorsal surface of HH22 chick limb buds and allowed the embryos to develop for two days until HH27-28 (Figs 4P,4Q,4R,4S,4T,4U,4V,5). Previous work had shown that this implantation regime does not disrupt digit pattern as do earlier and more anterior/distal implants [42,49]. After shh over-expression, ptc1 and gli1 mRNAs are up-regulated near Shh-expressing cells in a broader and more anterior region in and around the DMM than in contralateral controls (Fig. 4Q,4T). Analysis of gli2, another Hh-responsive gene implicated in Hh signalling, shows reciprocal expression to gli1 in un-manipulated limbs (Fig. 4U). Shh over-expression does not induce gli2 (Fig. 4V). Thus, ptc1, gli1 and gli2 expression suggest that the posterior DMM receives Hh signals around the time of slow muscle initiation at HH27, and that RCAS/Shh implantation augments this signal and expands it into the anterior DMM.
Figure 5
Shh/RCAS-infection of chick wings blocks early muscle differentiation. A. Pellet of chick embryo fibroblasts expressing theShh/RCAS replication-competent retroviral vector is grafted into the dorsal mesenchyme of chick right wing buds at HH22 and embryos were maintained to HH27-28. B-G. Contralateral (B-D) and Shh/RCAS infected (E-G) wings were serially sectioned and stained for pan-MyHC (B,E), anti-slow MyHC (C,F), and anti-Hh (D,G) antibodies. Control left wings have a normal distribution of muscle fibres and lack Shh staining in muscle-forming mesenchyme (B-D) whereas some Shh/RCAS right wings show a loss of pan-MyHC-reactive cells (E). Ectopic slow MyHC is present in the anterior dorsal muscle mass (F, arrow). H-J. Fibre numbers counted in adjacent sections at comparable levels of control contralateral and operated HH27-28 limbs. The values presented must be regarded as an imprecise reflection of absolute fibre numbers because resolution of small fibres from adjacent larger fibres was difficult and varied depending on the orientation of fibres within the section. Total fibre numbers (H), slow fibre numbers (I) and the proportion of fibres that contained slow MyHC (J) are presented from four limbs showing normal DMM extent (Increased slow limbs), five limbs showing reduced DMM (Reduced muscle limbs) and the pooled data (All limbs). Dorsal is up and posterior to the left in panels B-G. (u) ulna, (r) radius. Scale bar = 500 μm.
Similarly implanted limbs were serially-sectioned and analysed for expression of Shh protein, slow MyHC and pan MyHC. Control contralateral limbs show a broad DMM, with slow MyHC in the posterior portion above the ulna condensation (Fig. 5B,5C). Among treated wings, three classes of outcome are observed. In ~50% (8/15) wings, there was a complete loss of muscle tissue in the posterior region of the dorsal zone and reduced muscle in the anterior region (Fig. 5E,5H). In another ~30% (4/15) wings, the DMM had altered shape and a variable reduction in the total number of differentiated fibres in the region of Shh accumulation (Fig. 5E,5H). The three remaining Shh/RCAS wings revealed no phenotype, correlated with low ectopic Shh protein and young age (data not shown). In all affected wings, the anterior DMM that would not normally express any slow MyHC contained significant levels of slow MyHC, such that the proportion of all fibres that contained slow MyHC was doubled (Fig. 5F,5I,5J). In most operated wings showing a muscle phenotype, Shh was detectable in or close to the DMM (Fig. 5G, arrowhead). The location of highest ectopic Shh correlated with loss of muscle fibres in severely-affected wings (Fig. 5E,5F,5G; arrowheads). Thus, exposure of developing chick wing buds to Shh leads to a reduction in total muscle differentiation 48 hours after grafting, as already reported [50,51]. Despite this reduced myogenesis, we found an increase of slow muscle.
Prolonged Shh over-expression enhances limb and muscle size without increasing slow
To examine the longer term consequence of Shh over-expression on muscle formation we permitted implanted embryos to develop for three or four days to HH30 or 32 when significantly more fibres have formed in controls. At these stages, Shh-treated limbs are obviously bigger than control limbs (Fig. 6 compare A to D and G to K for HH30 and 32, respectively). At HH30, operated limbs (Fig. 6D,6E) show increased DMM area and altered muscle splitting, compared to control wings (Fig. 6A,6B). By HH32, analysis of MyHC expression in the autopod shows a large increase in muscle fibres compared to contralateral control limb (data not shown). In contrast, in zeugopod, expansion of the DMM is variable: some limbs have enhanced muscle mass whereas others simply appear disorganized, with muscle splitting less clear cut than in the VMM (data not shown). Thus, as reported previously [60], Hh over-expression ultimately perturbs muscle splitting and enhances terminal differentiation.
Figure 6
Later-forming fibres do not accumulate slow MyHC in response to Shh. Dorsal muscle mass from contralateral (A-C) and operated (D-F) wing implants grown to HH30 and stained for Pan MyHC (MF20, A,D) and slow MyHC (Na8, B,E) after in situ hybridisation for Shh mRNA (C,F). Shh mRNA is widespread in the dorsal muscle mass, which is expanded and poorly split (A,D). In contralateral muscle most slow fibres are in the medial region of the dorsal/posterior muscle block (B). In the operated limb, slow MyHC is more abundant in the dorsal region, with numerous cells in ectopic lateral locations (arrows, E). Note that the ventral region of the dorsal muscle mass has few if any ectopic slow fibres.
Concerning slow differentiation, Shh over-expression increases the number of slow fibres at HH30 (Fig. 6B,6E). This change may relate to the increase in total fibres as many fibres do not express slow MyHC despite proximity to the Shh source (Fig. 6A,6D). By HH32, no increase in slow fibres, as a proportion of total fibres, is detected and slow fibre pattern appears normal (data not shown). Thus, the induction of an increased proportion of slow fibres was not a continuing process within the chick wing bud, despite the continued presence of Shh.
Blockade of Hh reduces slow muscle differentiation
To address the role of endogenous Hh in muscle patterning, we implanted hybridoma cells secreting a functionally-blocking anti-Hh antibody into the proximal dorsal limb at HH 21-24 and examined subsequent muscle differentiation (Fig. 7). Anti-Hh hybridoma cells cause a reduction in limb cross sectional area by 15% (P < 0.001, n = 14, 0.64 to 0.55 mm2), whereas control hybridoma cells have no effect. Overall DMM area is also reduced at HH27/28 but without obvious change in location or shape. However, the initial appearance of slow MyHC is more severely reduced or blocked entirely compared to contralateral control limbs (7/9 limbs at HH27/29, Figs 7A,7B,7C,7D,8). The effect of the implant is generally more severe in younger limbs, but in all affected cases extended over at least 480 μm of the zeugopod (Fig. 8). We next quantified slow MyHC-expressing fibres in at least four sections spaced by 120 μm within each limb in comparison to contralateral control limbs at the same proximodistal level. At HH27/28 or 28 anti-Hh treated limbs showed a 79% (± 12% SEM, n = 4) reduction in slow fibres. More mature treated limbs at HH28/29 showed a lesser reduction of 27% (± 19% SEM, n = 5). In even older limbs (HH30), there is a reduction in DMM extent in the zeugopod region of anti-Hh treated limbs (4/4 limbs, Fig. 7E,7F,7G,7H). However, slow MyHC is not noticably reduced (4/4 limbs, Fig. 7E,7F,7G,7H). Limb outgrowth and digit formation are unaffected at any stage (data not shown), suggesting that the anti-Hh antibody does not reach sufficient titre to prevent ZPA activity. Although implants were generally found within the elbow or stylopod region, changes in stylopod muscle are less marked than in zeugopod (data not shown), consistent with a diminished role for Hh in the later stages of stylopod muscle formation. Control hybridoma cells (encoding a high-titre IgG1 anti-MyHC antibody N2.261) have no significant effect on any parameter examined (6 limbs, Fig. 7I,7J). Thus, Hh is required for normal initiation of slow myogenesis and early muscle differentiation in zeugopod.
Figure 7
Anti-Hedgehog antibody delays slow MyHC and reduces muscle differentiation in chick wing bud. Implants of anti-Hh (5E1; A,B,E,F) or control anti-MyHC (N2.261; I,J) hybridoma cells were placed into HH21-24 chick wing buds and analysed two days later at HH27-30 for pan MyHC (A4.1025; A,C,E,G,I) or slow MyHC (Na8; B,D,F,H,J), in parallel with contralateral limb controls (C,D,G,H). A-D. A less mature wing shows failure of slow MyHC expression (arrows) with little effect on overall muscle differentiation. E-H. A more mature wing showing reduced muscle differentiation, but slow MyHC is present in the residual muscle mass. I,J. Control implants have no effect on timing, extent or type of muscle differentiation. Slow fibres are less abundant in some regions, as in unmanipulated wings (red arrows, G-J). In this example, the Cellagen block containing hybridoma cells detected by the secondary reagents is located within the forewing region (arrowheads) but does not disrupt muscle pattern. Dorsal up, posterior to left. Contralateral images have been reversed to aid comparison.
Figure 8
Preferential inhibition of slow myogenesis by anti-Hh antibody. Schematic drawings of 120 μm spaced sections from each of the seven affected anti-Hh implanted limbs and aligned contralateral controls. Black outline shows the DMM, red stipple indicates slow MyHC. Drawings were made from identical low magnification images of adjacent sections reacted immunohistochemically for A4.1025 or Na8. Two young limbs (HH27/28) show marked reduction in slow MyHC, but little effect on DMM area. Slightly older limbs show a reduction in slow MyHC accompanying a diminished muscle mass. In two limbs, the Cellagen implant is present within the elbow region. Distal limb at top of stack, elbow at base. Within each section dorsal is up and anterior to left. Contralateral limbs are flipped horizontally to aid comparison.
Murine Hh knockouts contain slow muscle fibres
The early loss, but later recovery, of slow MyHC expression in anti-Hh treated limbs raised the possibility that the implant might lose effectiveness with time in vivo. However, later implants had lesser effects on muscle growth (4/4, data not shown), arguing that Hh has less role in later myogenesis. To examine the issue more definitively, we turned to mice lacking Hh genes. In the mouse, many slow fibres arise in deep regions of the limb near to the source of Ihh from developing long bones [61,62]. As in other mammals, only a single slow MyHC gene is known in mice, but primary slow fibres do fall into two distinct populations with different innervation and fate: deep fibres remain slow, superficial ones turn fast [61,63]. We, therefore, examined myogenesis in mice lacking Ihh at a stage when deep slow fibres display their unique character. Hindlimb elongation is severely reduced in these animals, and this is accompanied by a decrease in muscle tissue (Fig. 9A). However, forelimb growth is relatively normal, and so is limb muscle content and pattern (Fig. 9B). No obvious lack of slow MyHC is observed in either fore- or hindlimbs of Ihh-/- mice (Fig. 9A,9B). Thus, ablation of Ihh alone permits fairly good limb muscle patterning, similar to the situation in older chick limbs in which Hh signalling is reduced.
Figure 9
Murine Hh knockouts have inefficient differentiation and delayed slow myogenesis. A, B. Hindlimbs (A) or forelimbs (B) from E18.5 Ihh-/- or sibling mice were cryo-sectioned and stained for slow (A4.840) and fast (N3.36) MyHC. C. Whole E9.5 Shh-/- or sibling embryos were cryo-sectioned and stained serially for pan (A4.1025) or slow (A4.840) MyHC. Comparable anteroposterior levels are shown, based on the orientation of the heart elsewhere in the sections. s soleus, t tibialis anterior, e extensor digitorum longus, g gastrocnemius, T tibia, F fibula, u ulnar, r radius, A anterior, P posterior, D dorsal, V ventral, L lateral, M medial.
In shh-/- mice limb outgrowth and muscle growth is severely curtailed preventing meaningful analysis of muscle pattern [52]. So we examined slow MyHC expression in the reduced somitic muscle. In wild-type E9.5 mice, around 12 rostral somites contain differentiated muscle fibres expressing MyHC (Fig. 9C). Slow fibres were observed in rostral somites, but not in the two-three caudalmost MyHC-expressing somites (n = 6 mice). In rostral somites, many fibres lacked slow MyHC, but expressed embryonic MyHC (Fig. 9C and data not shown). This confirms that embryonic MyHC is acquired before slow MyHC in early muscle fibres [64]. In sections from shh-/- mice, fewer differentiated fibres were present, but around eight somites contained differentiated muscle which was often mis-oriented (6/6 individuals; Fig. 9C). Among residual fibres slow MyHC was undetectable in most animals (5/6; Fig. 9C). The single animal containing slow MyHC was developmentally more advanced, based on the presence of MyHC in more somites. Thus lack of Shh in somites leads to reduced early differentiation and delayed slow MyHC accummulation, as observed in chick limbs treated with anti-Hh antibodies.
Discussion
Hitherto, all studies of the actions of Hh on amniote muscle have failed to rule out indirect effects deriving, for example, from Hh eliciting secondary signals from adjacent non-muscle cells. Here we show that Hh can directly promote terminal differentiation and slow MyHC accumulation by at least some myoblasts in cell culture. We find that Hh is required for the earliest definitive slow myogenesis in chick limb buds and use this new understanding to develop a simple model of a role of Hh signalling in limb myogenesis.
Hh directly induces muscle differentiation
The results presented show that Shh can promote the terminal differentiation of muscle fibres both in vivo and in vitro. Our observation that Shh promotes terminal differentiation of C2 cells (Fig. 2) definitively demonstrates that Shh can be a myoblast differentiation factor, at least on this adult muscle-derived cell line. This effect occurs even in the presence of growth factors. Early limb myogenic cells in culture also respond to Shh exposure by increased differentiation (Fig. 1). We also observed increase of muscles in Shh-treated limb (Fig. 6). This is consistent with the numerous reports showing that Shh increases muscle differentiation in explant cultures or in vivo [14,25,33,52,53,65]. Conversely, and crucially, we find that local reduction of Hh function in the chick wing reduces muscle differentiation (Figs 7,8). This suggests that lack of Shh-driven myoblast differentiation may contribute to the severe reduction of muscle in early limb and somites in shh deficient mice [14,52]. It is, therefore, highly likely that one action of Hh is the direct promotion of myoblast differentiation in developing chick wing bud.
A direct action of Hh on myoblasts is also supported by the rapid accumulation of gli1 mRNA, a downstream target of Hh signalling, in C2 cells and in limbs. In unmanipulated chicks, significant levels of ptc1 and gli1 mRNA accumulate in muscle masses, being highest in the posterior DMM (Fig. 4). This suggests that myogenic cells in both wing DMM and somite are exposed to Hh signals around the time of their first differentiation [13,49,59,66,67].
Which Hh could promote muscle differentiation in limbs? As the anti-Hh antibody blocks the function of both Shh and Ihh [68], our in vivo manipulations do not address this issue. Ihh is an obvious candidate, as ptc1 and gli1 expression are up-regulated in the posterior DMM at the time and location of commencement of Ihh expression in cartilage anlage around HH27 [59]. However, forelimb myogenesis in Ihh-/- mice appears relatively normal, although we can not rule out undetected transient defects. Even in Ihh-/- hindlimb, where muscle differentiation is greatly reduced, it is impossible to ascribe this reduction to a direct action of Ihh because failure of long bone elongation could prevent muscle growth through lack of stretch-induced hypertrophy signals. Similarly, Shh is not absolutely required for the initiation of some murine limb muscle differentiation [52]. However, gli1 expression suggests that Shh signalling extends quite far into what we have called the pre-myogenic zone [47] in the distal limb until at least HH27/28 (Fig. 4). So differentiating myogenic cells may also be exposed to low levels of Shh. In zebrafish somites, distinct levels of Hh signalling elicited by the combined action of at least three Hh genes lead to different myogenic outcomes [18,69-72]. So the additive effects of Ihh and Shh, perhaps having different effects at particular overall concentrations, likely contribute to the sculpting of muscle differentiation.
Hh and slow myogenesis
We found that Shh has a consistent positive effect on slow myoblast differentiation. Differentiating chick wing myocytes express slow MyHC more frequently after Shh exposure in vitro (Fig. 1) or Shh over-expression in vivo (Fig. 5). Conversely, an early effect of blocking Hh in the wing is failure of slow MyHC accumulation (Figs 7,8). And Shh-/- mice have delayed slow MyHC accumulation in somites, perhaps due to loss of an early fibre population (Fig. 9). Lastly, one line of C2 cells accumulates more slow MyHC after Shh exposure (Fig. 3). Other C2 lines that do not express significant levels of slow MyHC in control conditions fail to up-regulate slow MyHC in response to Hh, perhaps indicating that Hh is unable to open the slow MyHC genomic locus. Both zebrafish and Xenopus embryos require Hh signalling to make some early populations of slow fibres, but not others [73,74]. This argues strongly that Hh-driven slow myogenesis is an ancestral character of amniotes. Nevertheless, as in lower vertebrates, slow fibre formation does eventually occur in amniotes with defective Hh signalling. Indeed, considering the complex pattern of slow and fast fibres in older muscle, it is clear that many factors in addition to Hh must be involved in establishing the pattern. Our evidence suggests that Hh acts primarily during the earliest stages of limb myogenesis.
What is the relationship between the differentiation-promoting and slow MyHC-promoting actions of Shh on C2 cells? Whereas intracellular signalling and terminal differentiation was triggered rapidly in all C2 cells, slow MyHC up-regulation required longer Shh exposure. Therefore, one can argue that differential cell survival could account for the Shh-dependent increase in slow MyHC, particularly as Shh, or tissues that secrete it, have been shown to promote survival of some myogenic cells [31,32,52,75]. However, we think a purely survival effect is unlikely, for several reasons. First, slow MyHC accumulation in single cells is greater with Shh, suggesting induction rather than simply enhanced survival. Second, assays for apoptosis in our C2 cultures revealed very little cell death, and this was unaffected by Shh exposure (unpublished result). Third, blockade of Hh in the wing bud reduces slow MyHC without a proportional reduction in differentiated muscle (Figs 7,8). Fourth, Hh over-expression in vivo induces ectopic slow while simultaneously reducing total differentiation (Fig. 5). Fifth, in cultured zebrafish blastomeric cells, some of which spontaneously form muscle, Shh induces conversion to a slow fate without affecting cell survival [76]. Conversely, reduction of Hh signalling in zebrafish prevents slow myogenesis without inducing cell death [69]. As altered cell survival does not explain the differentiation promoting activity of Hh, it seems unnecessary to invoke it in regard to slow myogenesis. In C2 cells, blockade of apoptosis appears unlikely to explain the slow promoting activity of Hh, leaving promotion of slow differentiation as the prime explanation. In vivo, the potential combination of direct and indirect effects of Hh, possibly on several myoblast subsets, make attribution of direct effects to Hh action on myoblasts impossible (see below). Nevertheless, our in vitro findings highlight direct induction of slow differentiation by Hh as a mechanism requiring serious consideration.
Could simply enhancing terminal differentiation account for the increase in slow? In vivo manipulations fail to reveal a correlation between increased slow expression and enhanced differentiation (Fig. 5). Nor do the number of nuclei in a cultured myotube (a rough assay of maturity) predict whether slow MyHC is induced (Fig. 3). Indeed, C2/4 cells differentiate well in response to Shh but fail to show the up-regulation of slow MyHC elicited by Shh in C2X cells, which differentiate less extensively with or without Shh. In addition, not all myoblasts in any line respond similarly to Shh exposure. It seems probable, therefore, that both intrinsic and micro-environmental differences between myoblasts regulate their response to Shh and could influence whether the response is simply terminal differentiation, or includes other events, such as slow MyHC accumulation.
Myoblast hetereogeneity of response to Hh
Intrinsic myoblast heterogeneity, possibly based on cell lineage, may also influence Hh response. As with C2 cells, not all cultured chick limb bud myoblasts respond similarly to Shh exposure. Shh efficiently enhanced terminal differentiation from 50% to ~80% of myogenic cells, showing that at least 30% of myogenic cells are likely to be Shh-responsive. However, only a few percent of chick myoblasts acquired slow MyHC in response to Shh (Fig. 1). Early limb myogenic cells have distinct clonally-heritable tendencies to either express slow MyHC or not do so [3,45,48,77]. We suggest that, while most myoblasts may be Shh sensitive, sub-populations may respond differently based on their intrinsic capacity. This view parallels that of Stockdale and colleagues based on experiments showing differences in the myoblast populations forming distinct limb muscles [6,7]. Fibres in distinct muscles differ in slow MyHC from their inception [1]. So it is possible, by analogy with the situation in Drosophila [8] that the increase in slow fibres reflects a change in muscle identity of founder myoblasts, rather than a direct induction of slow MyHC. Altered myoblast identity could contribute to the failure of muscle splitting after Shh over-expression in vivo. Thus, by showing differential effects of Shh on distinct clonal myoblast lines that parallel those in primary cultures and in vivo (see below), our findings indicate that cell intrinsic differences determine the response of myogenic cells to Hh.
In the zeugopod, the earliest muscle differentiation is reduced, but not ablated, by introduction of anti-Hh antibody or in Shh-/- mice (Figs 7,8; [52]). Hh signalling is required for some but not all early somitic myogenesis [15]. This shows that some myoblast populations do not require Hh for differentiation. In later limbs, blocking Hh reduces fibre formation and extra Shh augments differentiation (Figs 5,6,7,8). Similarly, Hh blockade reduces slow MyHC and Shh over-expression augments slow fibres early, but has little or no effect on slow MyHC expression in later muscle. Taken together, these observations indicate that myoblasts generating the earliest fibres in the zeugopod (before about HH28) may respond differently to Hh from those contributing to DMM growth after this stage. Many limb signals other than Hhs undoubtedly influence muscle pattern and likely affect the response to Hh [78,79]. Resolution of whether myoblast lineage or environmental effects underlie this difference will be important.
Indirect proliferative effects of Hh on myogenic cells
Direct pro-differentiative effects of Hh on some myoblasts do not rule out other direct or indirect effects of Hh. Our results confirm and extend previous reports showing that Shh can induce proliferation of myoblasts [49,50], both in limb buds and in primary cultures. However, other cell types are affected in limbs because Shh-treated limbs are bigger (Fig. 6). There is good evidence for effects of Hh on non-myogenic limb tissue. Shh over-expression causes limb hypertrophy with up-regulation of ptc1 and gli1 outside muscle masses and increase in non-myogenic tissue area (Figs 5,6). Similarly, Shh causes growth of non-myogenic as well as myogenic cells in our chick primary cultures (Fig. 1) and in somite explants [32]. Conversely, inhibition of Hh in wing with anti-Hh antibody reduces limb growth in addition to reducing muscle mass size: this effect is first noticeable in non-muscle tissue (Figs 7,8). Nobody has reported a proliferative effect of Shh on myoblasts independently of a proliferative effect on other cells. On the contrary, in the C2 cell line we clearly found no mitogenic effect of Shh (Fig. 2). Moreover, in zebrafish, Shh is not a mitogen for slow muscle precursors: muscle differentiation is delayed in the sonic-you mutants that lack Shh [69] and induced in embryos over-expressing Shh [18,19]. Therefore, either chick wing cells respond differently to Shh compared with other myoblasts or mitogenic effects of Shh on myoblasts are indirect. It is highly likely that Hh action on non-myogenic cells leads to release of myoblast mitogens. One hypothesis has already proposed that Shh acts through BMPs to amplify the number of myogenic cells [50] and BMP induction by Ihh causes cartilage proliferation indirectly [68]. So the temporary inhibition of terminal differentiation by Shh over-expression in limbs (Fig. 5; [50]) may be a consequence of an indirect effect of Hh signalling on myoblast proliferation.
Combining our results with published data, we propose a model in which Hh promotes muscle differentiation directly in myoblasts within the muscle masses. But Hh also promotes myoblast proliferation indirectly by eliciting muscle growth factors from non-myogenic limb cells. This hypothesis explains how Hh may contribute to growth of muscle masses by increasing myoblasts at the edge, where proliferative signals from non-myogenic cells would predominate over the direct differentiative signal. Deeper within the muscle mass, pro-differentiative signals including Hh would be in the ascendant, adding new primary fibres at the periphery of the existing differentiated zone. At early stages distinct levels of Hh signalling may trigger slow myogenesis, possibly in sub-populations of myoblasts. Once muscle splitting commences Hh signalling declines, as indicated by reduced gli1 expression, and other influences probably determine the decision of later myoblasts to divide or differentiate. Muscle-specific ablation of Hh responsiveness will be required to test this hypothesis definitively.
Conclusions
We show that Hh can directly promote myoblast differentiation, at least in vitro. In vivo in chick limb bud, Hh signalling is occurring at the right time and place to affect early slow myogenesis. We introduce a new methodology, Cellagen implants of hybridoma cells secreting functionally-blocking antibody, and show that Hh is required for proper early slow (but not fast) muscle differentiation. Conversely, Hh over-expression induces ectopic early slow muscle in chick limb bud. Neither gain nor loss of later Hh function affects differentiation of later-formed slow muscle. Thus early limb Hh levels promote slow myogenesis, but are unlikely to be solely responsible for the details of slow fibre pattern. The data suggest a simple model of how direct and indirect effects of Hh sculpt early limb myogenesis.
Methods
Manipulated chick forelimbs
Chick embryo fibroblasts (CEF) were transfected with Shh/RCAS, a replication-competent retrovirus containing the entire cShh coding sequence. Anti-Hh 5E1 hybridoma cells (~2 × 104) were embedded in a 10 μl Cellagen block. Pellets (~100 μm diameter) or block fragments (2 μl) were implanted into Rhode Island Red chick embryo right wing buds at Hamburger and Hamilton stages (HH) 21-24, avoiding complications due to skeletal pattern alteration by grafting on the dorsal side in the future forewing region, as previously described [49]. Embryos were maintained in a humidifier at 37°C, for 2–4 days and analysed at HH27-29 (E6) pre-splitting, and HH30-32 (E7/8) mid/post-splitting of DMM into its component muscles.
Embryos were fixed at -20°C in methanol, rehydrated in graded PBS, soaked for 2 hrs in 20% sucrose, transferred to a 2:1 mixture of 20% sucrose and Tissue-Tek cryoprotectant (Bright), experimental and contralateral wings aligned and frozen in a single block and cryosectioned. Primary monoclonal antibody supernatants of A4.1025 [80], BA-D5, A4.840 and N3.36 [81] were diluted 1:10 [61,80]. EB165 and Na8 ascites, gifts of E. Bandman (University of California, Davis) were used at 1:5000 [57,82]. To detect ectopic Shh protein, 5E1 supernatant [83] was dilutied 1:10. MF20 and most other antibodies used in this study are available from Developmental Studies Hybridoma Bank. First antibodies were detected with biotin-conjugated horse-derived anti-mouse IgG, or a biotin-conjugated goat-derived anti-mouse IgM (Vector) and ABC Vectastain kit as described [18]. In situ mRNA hybridisation was after [49]. Identification of chick forewing muscles was according to [84] and staging based on limb and muscle mass morphology according to [41].
Primary cultures
Following the methods of Stockdale [3,85], both forelimbs were removed from embryos around HH22 in Dulbecco's modified Eagle's medium (DMEM, Sigma). Limbs were washed with sterile PBS, incubated with trypsin:EDTA (Gibco) for 10 mins, dissociated by trituration and the cells washed, filtered through two 80 μm pore filters (Gibco) and pre-plated on a 90 mm collagen-coated dish for 10 min at 37°C. After this incubation period, 30–40% of cells stuck to the dish but fewer than 1:1000 were myogenic. Non-adherent cells were collected and plated in triplicate at either 2 × 105 (low density) or 4 × 105 (high density) cells per Nunc 35 mm plate in either unconditioned DMEM with 10% horse serum (HS) and 2% chick embryo extract (CEE) or medium that had been conditioned for 24 hours on 90% confluent RCAS/Shh infected QT6 quail fibroblasts (ShhQT6) or the parent QT6 fibroblasts [49,86]. Fresh medium (conditioned or not) was added after 24 hours, and the cells fixed 50–55 hrs after plating. Prior to conditioning QT6 and ShhQT6 cells were maintained in DMEM 10% foetal bovine serum (FBS -Gibco) and 2% chick serum (Gibco). Cultures were washed in PBS, fixed for 5 mins in -20°C methanol, rehydrated in PBS and stained as for the cryosections. Replicate dishes were singly stained with antibodies A4.840 to detect slow MyHC, A4.1025 to detect pan-MyHC and anti-desmin (Sigma, 1:500 dilution) to detect both myoblasts and myotubes. Dual immunofluorescence showed that all MyHC-reactive cells are strongly desmin-reactive and that immunohistochemistry is more sensitive (data not shown). Total nuclear (cell) numbers were counted on a Zeiss Axioplan 2 microscope in ten separate 10x fields on each dish. All nuclei within immunohistochemically-stained cells were counted on each dish.
Growth and addition of Shh conditioned medium to mouse myoblasts
C2 cells were obtained from three sources i) C2C12 from ATCC, ii) C2/4 from Y. Nabeshima and iii) C2X which arose in late passage cultures of C2C12 from the lab of H. Blau. All were maintained on plastic by standard procedures prior to plating on collagen-coated glass chamber slides (16-well, LAB-TEK, Nalge Nunc International, USA) in DMEM 10% FBS, 2% chick serum with antibiotics and differentiated by switching to DMEM 2% HS. Conditioned medium was created by incubating QT6 or ShhQT6 cells for 36 hours with either growth or differentiation medium. Each QT6 cell culture was used to condition only a single batch of medium. Purified Shh was synthesised in vitro and the biologically active proportion of the protein in each preparation is unknown, low and varies between batches (P. Ingham and T. Jessell personal communication) so no meaningful concentration can be given. C2 cells were fixed with methanol, stained by dual immunofluorescence with A4.840, A4.1025 and/or BA-D5 using class-specific secondary reagents (Jackson) and viewed under epifluorescence on a Zeiss Axiophot. Unless otherwise stated quantitation of differentiation was by scoring the number of nuclei in MyHC-containing cytoplasm (i.e. the number of C2 cells that differentiated into myocytes, whether or not these subsequently fused). Bromodeoxyuridine was added for the last two hours of a 24 hour culture in QT6 or QT6Shh conditioned growth medium. In situ hybridisation was performed on chamber slide cultures fixed with 4% paraformaldehyde followed by methanol and employed digoxigenin-labelled riboprobes essentially as described [87].
Authors' contributions
XL performed the C2 experiments. CSB analysed chick cultures, Shh over-expression in limb buds and mouse mutants. HB repeated the Shh over-expression, performed wholemount in situs and did the Hh knockdown. MAB and DD implanted RCASShh cells and did section in situs. SMH conceived and coordinated the study and wrote the manuscript with help from DD. All authors approved the final manuscript.
Acknowledgment
We thank Everett Bandman for antibodies, Tom Jessell for antibodies, 5E1 hybridoma cells and mouse Shh protein, Peter Currie and Philip Ingham for zebrafish Shh protein, Alex Joyner, Andrew Lumsden and Susanne Dietrich for cDNAs, Andy McMahon for Ihh-/- mice, Chin Chiang for Shh-/- mice and Abi Jensen, Graham Dunn, Pete Currie and Phil Ingham for advice. SMH was supported by MRC and EU QLK6-2000-530, XL by an EC Biomed 2 Ageing grant, and DD by the CNRS. CSB held a MRC PhD studentship and part of this work was reported in his PhD thesis (London University, 1999). | [
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15328538 | Loss of Skeletal Muscle HIF-1α Results in Altered Exercise Endurance
Abstract
The physiological flux of oxygen is extreme in exercising skeletal muscle. Hypoxia is thus a critical parameter in muscle function, influencing production of ATP, utilization of energy-producing substrates, and manufacture of exhaustion-inducing metabolites. Glycolysis is the central source of anaerobic energy in animals, and this metabolic pathway is regulated under low-oxygen conditions by the transcription factor hypoxia-inducible factor 1α (HIF-1α). To determine the role of HIF-1α in regulating skeletal muscle function, we tissue-specifically deleted the gene encoding the factor in skeletal muscle. Significant exercise-induced changes in expression of genes are decreased or absent in the skeletal-muscle HIF-1α knockout mice (HIF-1α KOs); changes in activities of glycolytic enzymes are seen as well. There is an increase in activity of rate-limiting enzymes of the mitochondria in the muscles of HIF-1α KOs, indicating that the citric acid cycle and increased fatty acid oxidation may be compensating for decreased flow through the glycolytic pathway. This is corroborated by a finding of no significant decreases in muscle ATP, but significantly decreased amounts of lactate in the serum of exercising HIF-1α KOs. This metabolic shift away from glycolysis and toward oxidation has the consequence of increasing exercise times in the HIF-1α KOs. However, repeated exercise trials give rise to extensive muscle damage in HIF-1α KOs, ultimately resulting in greatly reduced exercise times relative to wild-type animals. The muscle damage seen is similar to that detected in humans in diseases caused by deficiencies in skeletal muscle glycogenolysis and glycolysis. Thus, these results demonstrate an important role for the HIF-1 pathway in the metabolic control of muscle function.
Introduction
During exercise in normoxia, the partial pressure of oxygen in muscle tissue has been shown to dip to as low as 3.1 mm Hg, whereas in the capillary, it remains at 38 mm Hg (Hoppeler et al. 2003). In order to maintain effort, skeletal muscle exertion must be able to rely on pathways designed to help the tissue cope with oxygen stress after oxygen delivery capacity is exceeded. A switch between aerobic and nonaerobic metabolism during strenuous exertion requires mechanisms to adjust metabolic function, and this need is acute in extended exertion in skeletal muscle. It is clear that the transcription factor hypoxia-inducible factor 1α (HIF-1α) is an essential factor in maintenance of ATP levels in cells (Seagroves et al. 2001). In fact, although HIF-1α is typically thought of as acting only during hypoxia, its loss has an effect on both normoxic and hypoxic ATP levels in a number of tissue types (Seagroves et al. 2001; Cramer et al. 2003), and this implicates the factor in regulation of metabolic function even during conditions of normal physiologic oxygenation.
In skeletal muscle, signaling of fatigue has been studied extensively, and signaling of exhaustion involves, to some degree, elevated systemic lactic acid, a by-product of the glycolytic pathway of metabolism (Myers and Ashley 1997). Thus, the glycolytic pathway is intrinsically involved in muscle function and fatigue, and this in turn is linked to the response to hypoxia. To understand how the primary hypoxia-responsive transcription factor controls skeletal muscle function, we targeted mouse skeletal muscle for tissue-specific deletion of HIF-1α via the use of a conditionally targeted allele of the gene (Ryan et al. 2000; Schipani et al. 2001). This mouse strain was crossed into a strain transgenic for the skeletal-muscle-specific muscle creatine kinase (MCK) promoter, which drives expression of the cre recombinase gene (Bruning et al. 1998; Sauer 1998). We found that loss of the regulation of hypoxic response in muscle has a profound effect on the function of the muscle during exertion, with effects that mimic human metabolic myopathies.
Results/Discussion
In 4-mo–old mice with the skeletal-muscle HIF-1α gene knocked out (HIF-1α KOs), the frequency of excision was evaluated through real-time PCR techniques. We saw deletion frequencies consistent with those described previously for this cre recombinase transgene (Bruning et al. 1998) with some variation in penetration; mean frequency of deletion was 54.9%, with the highest frequency of muscle-specific deletion of HIF-1α being 72% in the gastrocnemius of 4-mo–old mice homozygous for the loxP-flanked allele (Table 1). This transgene is expressed at a lower level in cardiac tissue, and cardiac deletion was detected (Table 1); however, none of the phenotypes described below were seen in cardiac myocyte-specific deletions of HIF-1α (Figure 1A). Gross muscle sections were evaluated histologically to evaluate both vascularization and fiber type (Tables 2 and 3), and ultrastructurally to determine number of mitochondria (Figure 1B). No changes were detected in any of these features in HIF-1α KOs, except for a slight but statistically significant decrease in type IIA fibers in the soleus muscles (Table 3). Similar hematocrit and blood hemoglobin levels were seen in HIF-1α KOs and wild-type (WT) mice (Figure 2).
As can be seen in Table 4, significant changes in HIF-1α–dependent gene expression occur in muscle during exercise, including changes in genes involved in glucose transport and metabolism. Vascular endothelial growth factor (VEGF), which increases vascular permeability, and glucose transporter 4 (GLUT4), the muscle-specific glucose transporter, show increased levels in exercise and likely increase the availability of glucose to the muscle. The muscle-specific form of phosphofructokinase (PFK-M), phosphoglycerate kinase (PGK), and lactate dehydrogenase-A (LDH-A) are also up-regulated at the mRNA level by exercise, and this up-regulation is inhibited by the loss of HIF-1α, further demonstrating that HIF-1α is important for transcriptional response during skeletal muscle activity.
In Table 5, we show the changes in enzymatic activity in a number of key glycolytic enzymes affected by deletion of HIF-1α. As can be seen from the data, several of the enzymes assayed showed a decrease in activity in response to exercise. In particular, the activity of one of the key rate-limiting enzymes, PFK, was significantly lower following exercise in HIF-1α KOs compared to WT mice, indicating that HIF-1α KOs may have difficulty maintaining optimal PFK activity. The responses of other glycolytic enzymes to exercise were fairly similar between WT mice and HIF-1α KOs. These include no significant changes in phosphoglucose isomerase activity and significant, yet similar, decreases in aldolase, glyceraldehyde 3-phosphate dehydrogenase, and PGK activities. An exception to this is that WT muscles were able to significantly increase pyruvate kinase (PK) activity (see Table 4; p < 0.05). LDH activity was also increased in the WT mice, although the level did not reach statistical significance. Activities of both PK and LDH were not significantly changed in HIF-1α KO muscles following exercise. Increased activities of PK, and subsequently LDH, could be expected to lead to increased levels of lactate in the WT mice relative to HIF-1α KOs.In Figure 3A, it can be seen that the decrease in PFK activity in the HIF-1α KOs is correlated with a trend approaching significance (p = 0.10) toward an increased amount of hexose monophosphates (HMPs), which are pre-PFK glycolytic metabolites, following stimulation of the HIF-1α KO muscle. This increase was not due to differences in glucose uptake, since animals of both genotypes were able to significantly increase intramuscular glucose to a similar degree (Figure 3B). Consistent with decreased flow through the glycolytic pathway, however, the increased amount of HMPs was correlated with increased muscle glycogenolysis (Figure 3C) and increased depletion of phosphocreatine (PCr) (Figure 3D), with a resultant decrease in the PCr/ATP ratio in HIF-1α KO muscle (Figure 3E), although there was only a nonsignificant drop in overall muscle ATP concentrations (Figure 3F). Intramuscular levels of lactate did increase in both HIF-1α KOs and WT mice during stimulation, although lactate accumulation did not differ significantly between them (Figure 3G). In order to evaluate whether these changes had any effect on overall muscle force, we measured force and calcium release in isolated single fibers; as can be seen in Figure 4A and 4B, there were no significant changes in these parameters, indicating that the muscle can compensate at this level for the metabolic changes induced by loss of HIF-1α.
Given altered levels of glycolytic throughput without significant changes in intramuscular ATP levels, it is likely that there is increased activity of oxidative pathways in the HIF-1α KO muscle. Increased muscle oxidative activity is typical in patients with myopathies involving muscle glycolysis or glycogenolysis, including phosphofructokinase disease (PFKD) and McArdle's disease (Vissing et al. 1996). We analyzed the activity of citrate synthase (CS), a key allosteric enzyme of the citric acid cycle, in WT and HIF-1α KO muscle (Figure 5A), and found that it was up-regulated in HIF-1α KOs. CS is a mitochondrial enzyme that responds to decreases in ATP concentration allosterically, allowing for increased oxidative activity in the mitochondria. In addition, significant up-regulation of the mitochondrial enzyme beta-hydroxyacyl CoA dehydrogenase (B-HAD) was seen in HIF-1α KO muscle (Figure 5B). B-HAD is also affected by energy levels in the cell, and decreases in NADH/NAD+ concentration ratios cause the enzyme to increase mitochondrial oxidation of fatty acids (Nelson and Cox 2000). Increased activity of oxidative pathways in the muscle should result in more rapid lactate clearance, as in fact occurs in PFKD patients during exercise; this phenomenon gives rise to a “second wind” in these patients, and under some circumstances allows for an increase in exercise endurance (Vissing et al. 1996; Haller and Vissing 2002), although this was disputed in one recent study (Haller and Vissing 2004). This decreased lactate accumulation postexercise clearly occurs in the HIF-1α KOs, as can be seen in Figure 5C. This systemically lower level of lactate postexercise indicates that there may be a shift toward a more oxidative metabolism in skeletal muscle.
As mentioned above, patients with muscle glycolytic deficiencies demonstrate both increased exercise-induced muscle damage and a “second wind”; the latter phenomenon allows them to exercise for extended periods of time at submaximal levels. This is thought to be due to an increase in rates of oxidative ATP production, and a decreased utilization of and need for muscle glycogen (Vissing et al. 1996; Haller and Vissing 2002). To assess whether this is also the case in the HIF-1α KOs, both WT mice and HIF-1α KOs were subjected to endurance tests to assess muscle function. To first determine whether HIF-1α KOs were capable of extended activity during exercise, the animals were given a swimming endurance test. As can be seen in Figure 6A, HIF-1α KOs were capable of significantly longer-duration swimming activity when compared to matched WT controls (p < 0.05).
Further testing was done to determine the parameters of this increased endurance. HIF-1α KOs were run on an enclosed treadmill, with a 5° incline and an initial velocity of 10 m/min, with an increase in velocity every 5 min. In their first runs, HIF-1α KOs again had significantly greater endurance, as shown by their consistently longer run times compared to WT controls (p < 0.01, Figure 6B).
As it has been shown that muscle groups and fibers respond differently to eccentric exercise (i.e., downhill running) than to concentric exercise (i.e., uphill running) (Nardone and Schieppati 1988), mice from both genotypes were run on a 10° decline with the same velocity and time parameters as in the uphill runs. Eccentric exercises have been shown to recruit primarily fast-twitch glycolytic fibers for contraction, as opposed to the traditional recruitment of slower, smaller, oxidative motor units in concentric contraction, where animals with an increased capacity for muscle oxidation would be at an advantage (Nardone and Schieppati 1988). Now, the trend from swimming and uphill running tests was reversed, with WT mice able to run for a significantly longer time than HIF-1α KOs (p < 0.05, Figure 6C). Within genotypes, WT mice ran for significantly longer times downhill than uphill (p < 0.01); HIF-1α KOs did the reverse, and ran for significantly shorter times downhill than uphill (p < 0.05). Substrate utilization confirms the shift toward glycolytic fibers in downhill running; both genotypes had higher average respiratory exchange ratio (RER) values when running downhill compared with running uphill (Figure 6D and 6E).
PFKD and McArdle's disease demonstrate significant myopathic effects in muscle, including soreness and cramping induced by bouts of exercise. After 1 d of recovery from endurance testing, HIF-1α KOs had increased levels of the MM isoform of creatine kinase in their serum (unpublished data), indicative of skeletal muscle damage. To further investigate this finding, mice were run on a treadmill daily for 4 d. By the second day, the trend for increased endurance in the HIF-1α KOs was absent, and by the final day, HIF-1α KOs were running for significantly shorter times than they had on the first day (p < 0.01, Figure 7A). In addition, a repeated measures ANOVA performed on run times showed that the response of the HIF-1α KOs to the protocol was significantly different than that of the WT mice (p < 0.05). Histological examination of gastrocnemius tissue following 1 d of recovery revealed significantly greater amounts of muscle damage in HIF-1α KO tissue than WT tissue (Figure 7B). Staining of the tissue for proliferating cellular nuclear antigen (PCNA) and counts of positive nuclei (Olive et al. 1995) also revealed more cell division in HIF-1α KOs than in WTs, another indication that HIF-1α KOs had been subject to greater tissue damage (Figure 7C and 7D).
As noted above, both PFKD and McArdle's disease are marked by increased resting intramuscular levels of glycogen, a failure of serum lactate to rise during exertion, an exercise-induced “second wind,” and signs of muscle damage following exertion, including elevated levels of creatine kinase in the serum (Tarui et al. 1965; Layzer et al. 1967). In addition, PFKD is characterized by elevated levels of HMPs (Tarui et al. 1965; Layzer et al. 1967; Argov et al. 1987; Grehl et al. 1998) and greater PCr utilization during contraction (Argov et al. 1987; Grehl et al. 1998). We see many of these hallmarks of muscle deficiencies in glycolytic processing in HIF-1α KOs. The effects are not likely due to glucose uptake, as WT and HIF-1α KO intramuscular glucose levels were not different at rest or following stimulation (see Figure 3B), and both types of mice responded similarly to a glucose tolerance test (Figure 8A). Periodic acid–Schiff (PAS) staining of tissue from mice of both genotypes gave further demonstration of increased glycogen levels in resting muscles from HIF-1α KOs (Figure 8B).
Given the differences in performance observed in the HIF-1α KOs in eccentric and concentric exercise, it is clear that the HIF-1 pathway and hypoxic response have a central role in determining the capacity for work and endurance through regulation of glycolysis. It is also clear that these mice will provide an important model system to investigate the physiology of muscle response during work and oxygen depletion, and may be useful as a model for a group of very debilitating myopathic syndromes in humans.
Materials and Methods
Mouse strains and crosses.
Mice were generated from HIF-1α loxP-flanked allele mouse stocks backcrossed into a C57Bl6/J background. These were crossed into a C57Bl6/J strain containing the MCK/cre transgene. Controls were in all cases littermates that were genotyped as containing only the loxP-flanked HIF-1α allele or only the MCK/cre transgene. No phenotypic differences were seen in the two controls, so they were considered interchangeably as WT control animals.
Genotyping and real-time PCR for HIF-1α deletion
Mice from the above crosses were genotyped using DNA extracted from tail sections. DNA was then extracted from the gastrocnemius, heart, liver, and uterus of eight 4-mo–old, loxP-flanked HIF-1α–positive and MCK/cre–positive mice. HIF-1α levels were measured by real-time PCR analysis using the Universal PCR Master Mix Kit (Applied Biosystems, Foster City, California, United States) and the ABI Prism 7700 Sequence Detector (Applied Biosystems). Conditions for the PCR were one 10-min incubation at 95 °C (polymerase activation), followed by 40 cycles of 15 s at 95 °C (denaturation) and 1 min at 60 °C (anneal/extend). The degree of excision was calculated by comparing HIF-1α DNA levels to c-Jun DNA levels. HIF-1α real-time PCR primers and probe were as follows: forward primer, HIFLOX501/F 5′-CTATGGAGGCCAGAAGAGGGTAT-3′; reverse primer, HIFLOX574/R 5′-CCCACATCAGGTGGCTCATAA-3′; probe, HIFLOX/P 5′-(6FAM)AGATCCCTTGAAGCTAG(MGBNFQ)-3′.
Muscle histology and electron microscopy.
Paraffined gastrocnemius sections were deparaffinized and stained with Gill II hematoxylin. Sections were then washed successively in water, a bluing agent, water again, and 95% ethanol, and restained with eosin. Hematoxylin and eosin staining was performed by the University of California at San Diego (UCSD) Cancer Center Histology Resource (La Jolla, California, United States). Imaging was performed on sections mounted on slides using Cytoseal 60 (VWR, West Chester, Pennsylvania, United States). Electron microscopy was performed by standard methods on gastrocnemius muscle. Briefly, fixation was by 25.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4). Postfix was in 1% osmium tetroxide. The section was stained in 2% uranyl acetate in sodium maleate buffer (pH 5.2), then placed in Epon resin (VWR, West Chester, Pennsylvania, United States), and cured overnight at 60 °C. Fiber typing was performed using the metachromatic dye ATPase method (Ogilvie and Feeback 1990). PAS staining was performed as has been described (Bancroft and Stevens 1996).
Assessment of exhaustion.
Untrained, age-matched WT mice and HIF-1α KOs (WT, n = 10; KO, n = 14) were run either on an Omnipacer treadmill (Columbus Instruments, Columbus, Ohio, United States) or on an enclosed-chamber modular treadmill (Columbus Instruments) with a 5° incline at an initial velocity of 10 m/min. Velocity was increased by 2 m/min every 5 min during the assessment. Exhaustion was determined to be the point at which the animal would not resume running when provoked through a low-voltage power grid. Gas flow (O2 and CO2) into and out of the enclosed chamber treadmill was monitored using the Paramax O2 sensor and a CO2 sensor (Columbus Instruments) and analyzed using Oxymax software (Columbus Instruments) to determine metabolic parameters. The downhill running assessment (WT, n = 8; KO, n = 6) was carried out in the enclosed-chamber modular treadmill at a 10° decline using the same protocol as above.
In the swimming exhaustion assessment, a second group of WT and HIF-1α KOs (n = 8 for each class) was placed in a 30 °C water bath with mild turbulence. Exhaustion was determined to be the point at which the animal experienced three successive periods below the surface of more than 3 s.
Isolated stimulation and metabolic analysis.
The Achilles tendon was surgically freed from live, anesthetized mice (WT, n = 8; KO, n = 6) and attached to a force transducer to record contractile force. Muscles were electrically stimulated through excitation of the sciatic nerve. Stimulation was in the form of 8–10-V direct titanic contractions using 200-ms trains at 70 Hz with 0.2 ms duration. Initial frequency of tetanic contraction was one every 8 s and was increased every 2 minutes to one every 4 s and one every 3 s, up to the end point of 6 min. Isolated muscles were then immediately harvested and snap-frozen for ATP, lactate, phosphocreatine, and glycogen analyses. Samples were freeze-dried and analyzed by enzymatic assay as has been previously described (Bergmeyer 1974). The unstimulated gastrocnemius muscle from each mouse was used as a resting control.
Real-time PCR measurement of gene expression.
For basal gene expression levels, total RNA was isolated from gastrocnemius tissue from seven WT and five HIF-1α KOs using RNA-Bee (Tel-Test, Friendswood, Texas, United States). Reverse transcription was performed using the Superscript First Stand Synthesis System for RT-PCR (Invitrogen, Carlsbad, California, United States). Amplification was performed using the ABIPrism 7700 as described above. Reverse transcription real-time PCR primers and probes were as follows. For PGK-1: reverse primer, PGK/R 5′-CAGGACCATTCCAAACAATCTG-3′; forward primer, PGK/F 5′-CTGTGGTACTGAGAGCAGCAAGA-3′; probe, PGK/P 5′-(6∼FAM)TAGCTCGACCCA-CAGCCTCGGCATAT(TAMRA)-(phosphate)-3′. For VEGF-A: reverse primer, VEGF/R 5′-ATCCGCATGATCTGCATGG-3′; forward primer, VEGF/F 5′-AGTCCCATGAAGTGATCAAGTTCA-3; probe, VEGF/P (6∼FAM)TGCCCACGTCAGAGAGCAACATCAC(BHQ∼6∼FAM). For GLUT4: reverse primer, GLUT-4/R 5′-CCCATGCCGACAATGAAGTT-3′; forward primer, GLUT-4/F 5′-TGTGGCCTTCTTTGAGATTGG-3′; probe, GLUT-4/P 5′(6-FAM)TGGCCCCATTCCCTGGTTCATT(BHQ1-Q)-3′. For PFK-M: reverse primer, PFK-M/R 5′-AAGTCGTGCAGATGGTGTTCAG-3′; forward primer, PFK-M/F 5′-GCCACGGTTTCCAATAACGT-3′; probe, PFK-M/P 5′-(6-FAM)CCTGGGTCAGACTTCAGCATCGGG(BHQ1-Q)-3′. For LDH-A: reverse primer, LDH-A/R 5′-ATGCACCCGCCTAAGGTTCTT-3′; forward primer, LDH-A/F 5′-TGCCTACGAGGTGATCAAGCT-3′; probe, LDH-A/P 5′-(6- FAM)TGGCAGACTTGGCTGAGAGCAT(BHQ1-Q)-3′.
For changes in gene expression due to exercise, age-matched male mice (WT, n = 5; KO, n = 6) were run on a treadmill at 25 m/min for 30 min. Following the run, mice were euthanized and RNA was isolated and analyzed as described above.
Analysis of enzyme activity levels
For changes in enzyme activity levels with exercise, mice (WT, n = 5; KO, n = 12) were run on a treadmill using the same protocol as for the gene expression analysis. Tissue was harvested after the run and from resting mice (WT, n = 6; KO, n = 10), and enzymes were extracted and analyzed spectrophotometrically as has been described (Reichmann et al. 1983), with the exception that fructose 1,6-bisphosphate was replaced with fructose 2,6-bisphosphate for stabilization of PFK. Units of activity were normalized to milligrams of total protein using a BCA protein quantification kit (Pierce Biotechnology, Rockford, Illinois, United States).
Creatine kinase, serum lactate, hematocrit, and hemoglobin levels.
Creatine kinase levels were analyzed from serum from WT mice and HIF-1α KOs 24 h after running-induced exhaustion using a kit from Sigma (St. Louis, Missouri, United States). Creatine kinase isoforms were analyzed enzymatically and then fractionated by gel electrophoresis. Serum lactate levels were analyzed by the UCSD Comparative Neuromuscular Laboratory from blood obtained by cardiac puncture from six WT mice and six HIF-1α KOs following 25 min running time on the treadmill ramp at 25 m/min. Hematocrit and hemoglobin levels were measured from resting mice (WT, n = 6; KO, n = 4) by the UCSD Animal Care Program Diagnostic Laboratory.
Glucose tolerance curve.
Animals were assigned into either a sham (WT, n = 5; KO, n = 4) or glucose tolerance group (WT, n = 8; KO, n = 8). Experimental animals were injected with 0.3 g/ml glucose in PBS to achieve a dosage of 2 g/kg. Sham animals were injected with an equivalent amount of PBS. Blood was drawn from the tail at time intervals of 0, 15, 30, 60, and 120 min. Samples were then centrifuged to isolate plasma. Plasma blood glucose was quantified using the Infinity Glucose Kit (Sigma).
Calcium uptake measurements.
Intact individual muscle fibers (WT, n = 6; KO, n = 4) were mechanically dissected from the flexor brevis muscle and loaded with fura-2. Fibers were then stimulated while force generation and Ca2+ release were monitored.
Four-day endurance test
Endurance was tested by running 24 animals (WT, n = 10; KO, n = 14) on the Omnipacer Treadmill or the enclosed-chamber modular treadmill using the same exhaustion protocol described above. Mice ran according to this protocol every day for 4 d with a minimum of 22 h of rest between trials. Following the fourth trial, mice were given 24 h of rest and then euthanized. Tissue was harvested and stained using hematoxylin-eosin (as described above) and α-PCNA (Pharmingen, San Diego, California, United States) combined with a DAB Kit (Vector Labs, Burlingame, California, United States).
Statistical analysis
Statistical analyses (unpaired Student's t-test, Mann-Whitney test, ANOVA) were carried out using StatView software (SAS Institute, Cary, North Carolina, United States) or Prism software (GraphPad Software, San Diego, California, United States).
Abbreviations
B-HAD - beta-hydroxyacyl CoA dehydrogenase
CS - citrate synthase
GLUT4 - glucose transporter 4
HIF-1α - hypoxia-inducible factor 1α
HIF-1α KO - skeletal-muscle HIF-1α knockout mouse
HMP - hexose monophosphate
LDH-A - lactate dehydrogenase-A
MCK - muscle creatine kinase
PAS - periodic acid–Schiff
PCNA - proliferating cellular nuclear antigen
PCr - phosphocreatine
PFKD - phosphofructokinase disease
PFK-M - muscle-specific form of phosphofructokinase
PGK - phosphoglycerate kinase
PK - pyruvate kinase
RER - respiratory exchange ratio
VEGF - vascular endothelial growth factor
WT - wild-type
Figures and Tables
Figure 1
Exercise Capacity of Cardiac HIF-1α KOs and HIF-1α/MCK/cre Mitochondrial Density
(A) Mice lacking cardiac HIF-1α perform no differently in endurance running trials than WT mice, showing that the increase in exercise capacity seen in MCK/Cre mice is due to deletion of HIF-1α in skeletal muscle, not cardiac tissue.
(B) Mice lacking skeletal muscle HIF-1α have a slight but nonsignificant increase in mitochondrial density as measured by the number of mitochondria per electron microscope field of view.
Figure 2
Hematocrit and Hemoglobin Levels in HIF-1α KOs and WT Mice
(A) Hematocrit levels are virtually identical in both HIF-1α KOs (n = 3) and WT (n = 4) mice, indicating that loss of HIF-1α in skeletal muscle does not affect oxygen carrying capacity of the blood.
(B) In addition to similar hematocrit levels, WT mice and HIF-1α KOs have very close blood hemoglobin levels.
Figure 3
Intramuscular Metabolite Levels at Rest and Following Stimulation
(A) Glycolytic intermediates were measured from gastrocnemius muscles following the isolated stimulation protocol. Resting values represent levels in the unstimulated gastrocnemius from the same animals. HIF-1α KOs had a trend toward greater accumulated levels of HMPs during the stimulation protocol, although the difference did not reach statistical significance (p = 0.10). This difference could be indicative of a blockage in the glycolytic pathway at PFK.
(B) No significant differences were seen between HIF-1α KOs and WT intramuscular glucose levels at rest or following stimulation. Both HIF-1α KO and WT muscles were able to significantly increase glucose uptake, leading to greater levels of intramuscular glucose in response to stimulation (WT, p < 0.001; KO, p < 0.05).
(C) HIF-1α KOs have more stored glycogen than do WT mice. Glycogen levels were measured following the same stimulation protocol as in (B). The change in glycogen from rest to poststimulation was also greater in the HIF-1α KOs, indicating that they metabolized more glycogen in response to stimulation (p < 0.01; *p < 0.05, WT at rest vs. KO at rest).
(D) HIF-1α KOs utilize more PCr in response to stimulation than do WT mice. Similar levels of PCr were seen at rest, but HIF-1α KOs metabolized significantly more during stimulation (p < 0.05) and had much lower levels following the protocol (**p < 0.01, WT poststimulation vs. KO poststimulation).
(E) A trend toward lower PCr/ATP concentration ratios was seen in HIF-1α KOs relative to WT mice following stimulation, although the difference did not quite reach statistical significance (p < 0.10). A trend toward a greater drop from rest to poststimulation in the PCr/ATP ratio was also seen in HIF-1α KOs following stimulation (p < 0.10), indicating that they had to rely more heavily on PCr for ATP generation.
(F) Slight but nonsignificant differences were seen in whole-muscle ATP levels at rest or following stimulation. Although HIF-1α KOs exhibited altered substrate utilization, they were able to meet their ATP demands during the protocol.
(G) Both HIF-1α KOs and WT animals produced significant intramuscular lactate during the stimulation protocol; however, there was no significant difference in the amount produced by either genotype. Resting intramuscular lactate levels were also similar for WTs and HIF-1α KOs.
Figure 4
Force Generation and Ca2+ Release in Isolated Muscle Fibers during Stimulation
(A) No differences were seen in total force generation in isolated muscle fibers. Mechanically dissected fibers from the flexor brevis muscle were subjected to a fatiguing protocol. Neither initial nor final forces differed between HIF-1α KO and WT fibers.
(B) Ca2+ release and reuptake in HIF-1α KO and WT fibers was not different during the stimulation protocol. Ca2+ levels were measured in individual fibers through use of fura-2 Ca2+ indicator. The altered substrate utilization did not affect the ability of the fibers to maintain proper Ca2+ flux.
Figure 5
Oxidative Metabolism and Serum Lactate Production in HIF-1α KOs and WT Mice
(A) HIF-1α KOs have higher resting levels of CS activity. CS is an enzyme in the Krebs cycle that can be regulated allosterically by ATP levels. Increased CS activity is indicative of increased muscle oxidative capacity, which is common in patients with glycogenolytic or glycolytic myopathies (#p < 0.10, KO vs. WT).
(B) HIF-1α KOs have higher resting levels of B-HAD activity, which is indicative of a greater ability to oxidize fatty acids (**p < 0.01, WT vs. KO).
(C) Lower serum lactate levels were seen in HIF-1α KOs following a timed 25-minute run (*p < 0.05, WT vs. KO).
Figure 6
Endurance Capabilities of Untrained Mice
(A) HIF-1α KOs have greater endurance in swimming tests as shown by their ability to swim on average more than 45 min longer than WT (*p < 0.05, WT vs. KO).
(B) HIF-1α KOs have greater endurance than WT mice in uphill running tests. Although only a 10-min difference is seen between run times, it is to be noted that because of the protocol, this 10 min included two velocity increases (**p < 0.01, WT vs. KO).
(C) HIF-1α KOs have less endurance than WT mice in downhill running tests. The same protocol was used as in Figure 4A, except the mice were run on a 10° decline (*p < 0.05, WT vs. KO).
(D) RER uphill vs. downhill in WT mice. As would be expected from eccentric exercises relying more heavily on glycolytic fibers, the RER values are higher in mice running downhill than in those running uphill.
(E) RER uphill vs. downhill in HIF-1α KOs. Once again, higher RER values are observed for mice running downhill than those running uphill.
Figure 7
Increased Muscle Damage in HIF-1α KOs Following Repeated Exercise
(A) WT mice and HIF-1α KOs underwent a 4-d endurance test, in which animals were run to exhaustion on each of four successive days with a minimum of 22 h rest between trials. HIF-1α KOs demonstrated initially greater endurance under the protocol; however, by the second day, their endurance advantage was eliminated, and by the fourth day, HIF-1α KOs were running for a significantly shorter time (**p < 0.01) than on the first day, while WT animals were running for approximately similar times as on the first day. Repeated measures ANOVA revealed that the decrease in performance on each successive day was unique to HIF-1α KOs (p < 0.05).
(B) Example of hematoxylin and eosin staining of gastrocnemius muscles after 1 d of recovery by mice after the 4-d endurance test. Evidence of greater damage can be seen in HIF-1α KO muscles compared to WT muscles.
(C) Example of PCNA staining of gastrocnemius muscles from exercised mice, demonstrating increased levels of muscle regeneration in HIF-1α KOs.
(D) Number of PCNA-positive nuclei per square millimeter in gastrocnemius muscles of WT mice (n = 5) and HIF-1α KOs (n = 7) that ran repeatedly for 4 d. Although HIF-1α KOs have almost twice as many PCNA-positive nuclei per square millimeter, the difference is not significant, because of wild variations in that population. F-test analysis of the data reveals that the variance is much greater in the HIF-1α KO population than the WT population (p < 0.05).
Figure 8
Glucose Tolerance and Glycogen Storage
(A) No significant differences were seen in resting blood glucose levels in HIF-1α KOs or WT mice. Following injection of glucose at a dosage of 2 g/kg, no differences were seen in the maximum levels of blood glucose or the rate of glucose disappearance in either genotype.
(B) Representative PAS staining of gastrocnemius muscle from WT mice and HIF-1α KOs. HIF-1α KOs demonstrate darker staining, indicating more stored glycogen.
Table 1
Excision of HIF-1α in Various Tissues
Deletion levels are the average percent of HIF-1α deleted ± SE
Table 2
Fiber Typing of Gastrocnemius Muscle
Values are percent ± SE
Table 3
Fiber Typing of Soleus Muscle
Values are percent ± SE
* p < 0.05, WT vs. KO
Table 4
Relative Gene Expression Levels
Expression levels are means relative to resting WT for each gene ± SE
Percent increase indicates percent increase of postexercise average gene expression over resting average
*p < 0.05, rest vs. postexercise; **p < 0.01, rest vs. postexercise
Table 5
Glycolytic Enzyme Activity Levels from Gastrocnemius Muscles
Activities are in U/mg protein ± SE
GAPDH, glyceraldehyde 3-phosphate dehydrogenase; PGI, phosphoglucose isomerase
*p < 0.05, WT vs. HIF-1α KO for given exercised or resting state; **p < 0.05, rest vs. postexercise within given genotype
Footnotes
Conflicts of interest. The authors have declared that no conflicts of interest exist.
Author contributions. RSJ conceived and designed the experiments. SDM, RAH, MJK, IMO, WM, RPH, and FJG performed the experiments. SDM, RAH, MJK, IMO, MCH, PDW, FJG, and RSJ analyzed the data. CRK and RSJ contributed reagents/materials/analysis tools. SDM and RSJ wrote the paper.
Academic Editor: Michael Bate, University of Cambridge
Citation: Mason SD, Howlett RA, Kim MJ, Olfert IM, Hogan MC, et al. (2004) Loss of skeletal muscle HIF-1α results in altered exercise endurance. PLoS Biol 2(10): e288.
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15560850 | BAG-1 haplo-insufficiency impairs lung tumorigenesis
Abstract
Background
BAG-1 is a multifunctional co-chaperone of heat shock proteins (Hsc70/Hsp70) that is expressed in most cells. It interacts with Bcl-2 and Raf indicating that it might connect protein folding with other signaling pathways. Evidence that BAG-1 expression is frequently altered in human cancers, in particular in breast cancer, relative to normal cells has been put forward but the notion that overexpression of BAG-1 contributes to poor prognosis in tumorigenesis remains controversial.
Methods
We have evaluated the effect of BAG-1 heterozygosity in mice in a model of non-small-cell lung tumorigenesis with histological and molecular methods. We have generated mice heterozygous for BAG-1, carrying a BAG-1 null allele, that in addition express oncogenic, constitutively active C-Raf kinase (SP-C C-Raf BxB) in type II pneumocytes. SP-C C-Raf BxB mice develop multifocal adenomas early in adulthood.
Results
We show that BAG-1 heterozygosity in mice impairs C-Raf oncogene-induced lung adenoma growth. Lung tumor initiation was reduced by half in BAG-1 heterozygous SP-C C-Raf BxB mice compared to their littermates. Tumor area was reduced by 75% in 4 month lungs of BAG-1 haploinsufficient mice compared to mice with two BAG-1 copies. Whereas BAG-1 heterozygosity did not affect the rate of cell proliferation or signaling through the mitogenic cascade in adenoma cells, it increased the rate of apoptosis.
Conclusion
Reduced BAG-1 expression specifically targets tumor cells to apoptosis and impairs tumorigenesis. Our data implicate BAG-1 as a key player in oncogenic transformation by Raf and identify it as a potential molecular target for cancer treatment.
Background
BAG-1 is a multifunctional protein that is expressed in most cells. Originally identified as a Bcl-2 binding protein [1], other interaction partners of BAG-1 were described, including the serine threonine kinase C-Raf [2]. The C-terminal "BAG domain" of BAG-1 mediates the interaction with the Hsc70 and Hsp70 heat shock proteins [3], molecular chaperones that bind proteins in non-native states assisting them to reach a functional active conformation [4]. BAG-1 acts as a nucleotide exchange factor in this activation cycle [3]. The above findings indicated that BAG-1 might connect protein folding with other signaling pathways. Signaling networks promoting cell growth and proliferation are frequently deregulated in cancer [5]. The classical mitogenic cascade transmits stimuli from growth factor receptors via Ras, Raf, MEK and ERK to the cell nucleus [6]. C-Raf, like A- and B-Raf kinases also act at the outer membrane of mitochondria to augment cell survival [7,8]. Previously we had observed the stimulation of C-Raf kinase activity by BAG-1 in vitro [2]. Ras and B-Raf mutations have been found in various human cancers [9,10]. Evidence that BAG-1 expression is frequently altered in human cancers, in particular in breast cancer, relative to normal cells has been put forward but the notion that overexpression of BAG-1 contributes to poor prognosis in tumorigenesis remains controversial [11].
Methods
Animals
Mice used in these studies were generated and maintained according to protocols approved by the animal care and use committee at University of Würzburg. To inactivate the BAG-1 gene, we constructed a vector where exons 1 and 2 are replaced with a neomycin resistance gene. A phage clone with a 15-kb genomic insert from mouse strain 129/Sv spanning all seven exons of BAG-1 was identified and characterised using standard methods. The targeting construct contained 1,1-kb from the BAG-1 locus upstream of the neomycin resistance gene of plasmid pPNT [12] and 6-kb downstream. The upstream arm of 1,1 kb is located 5' to the start codon in the first exon of BAG-1 and the 3' arm of 6 kb is located downstream of exon 2. The mutation was introduced into embryonic stem cells by homologous recombination. Positive clones were identified by Southern blot analysis. Germline transmitting chimeras were obtained and bred to C57BL/6 mice. Further details will be described elsewhere. Heterozygous BAG-1 mice were genotyped by a PCR assay. The targeted BAG-1 allele was detected with primers P1 (5'-GAG TCT CCC GAT CCC TTT TCC), located upstream of exon 1, and P2 (5'-GAT TCG CAG CGC ATC GCC TT), located in the neomycin resistance gene, yielding a product of 600 base pairs. BAG-1 heterozygous mice were backcrossed at least three times onto C57BL/6 background before crossing with SP-C C-Raf BxB mice. Lung tumour mice expressing oncogenic C-Raf BxB were backcrossed at least six times onto C57BL/6 background.
Western blot
For the analysis of BAG-1 expression, lung lysates of the indicated genotypes were separated on 12,5% polyacrylamide-SDS (sodium dodecyl sulphate) gel, transferred to nitrocellulose Protran BA83 membrane (Schleicher&Schüll) and probed with rabbit anti-BAG-1 (FL-274) antibody (1:250, Santa Cruz Biotechnology). Amounts of protein were determined by Bradford protein assay to ensure equal protein loading for the analysis. Blots were developed using the appropriate horseradish peroxidase coupled secondary antibody and the ECL system (Amersham Pharmacia Biotech). Subsequently, the membrane was stripped and reprobed with rabbit antibody to glyceraldehyde 3 phosphate dehydrogense (1:2000, ab9485, Abcam Ltd.).
Histopathology and immunohistochemistry
Animals were sacrificed and lungs were fixed under 25 cm water pressure with 4% paraformaldehyde and embedded in paraffin. 5 μm sections were stained with hematoxylin and eosin and analysed. Pictures were taken using a Leica DMLA microscope and a Hitachi HV-C20A colour camera. Immunohistochemical staining to detect activated caspase-3, phospho-ERK (extracellular signal-regulated kinase), PCNA (proliferating cell nuclear antigen) have been described elsewhere [13]. Apoptotic, PCNA and p-ERK indices were determined by evaluating randomly chosen adenomas or fields of normal lung in 3–4 sections and determining the percentage of positive cells per 2000 cells at ×400.
Results and discussion
BAG-1 heterozygosity impairs C-Raf driven tumorigenesis
In order to assess the functional role of BAG-1 on tumorigenesis, we have generated a null allele of BAG-1. To inactivate the BAG-1 gene, exons 1 and 2 were replaced with a neomycin resistance gene. This strategy was chosen to disrupt the expression of all known isoforms of BAG-1 which are generated by alternate translation initiation of a single mRNA; the start codons are present in exons 1 and 2. Western blot analysis of liver protein extracts of BAG-1 deficient embryos showed the complete loss of all BAG-1 protein isoforms. Embryos homozygous for this allele died at midgestation at around E13,5, but the heterozygous animals (BAG-1+/-) are normal. A comprehensive description of the BAG-1-/- phenotype is subject of another manuscript.
Previously, we had generated a lung cancer mouse model by targeting constitutively active C-Raf kinase (SP-C C-Raf BxB) to the lung [14]. These mice develop multifocal adenomas early in adulthood. Based on the observation, that BAG-1 can activate C-Raf [2], we asked whether heterozygosity for BAG-1 would affect C-Raf BxB driven adenoma growth. We observed that lung tumour initiation was reduced by half in 1, 2 and 4 months old BAG-1+/- mice transgenic for SP-C C-Raf BxB compared to their BAG-1+/+ littermates. Tumour area was reduced by 75% in 4 month lungs of BAG-1 haploinsufficient mice compared to mice with two BAG-1 copies, see Figure 1. The histological picture emphasises the difference in adenoma formation between a representative SP-C C-Raf BxB/BAG-1+/+ and SP-C C-Raf BxB/BAG-1+/- lung. The difference in the staining intensity of the two lung sections derives mainly from the observation that the adenoma cells have a tendency to bind more intensively hematoxylin and eosin compared to normal lung cells. Thus, reduction of the BAG-1 gene dosage impairs the oncogenic activity of C-Raf in vivo.
Reduced BAG-1 expression in BAG-1 heterozygous lungs
Quantitative immunoblots demonstrated that the specific BAG-1 protein concentration in the lungs of BAG-1+/- mice was half the amount of BAG-1+/+ littermates, see Figure 2a. Moreover, immunohistochemical staining showed that BAG-1 was expressed in adenoma cells, see Figure 2b. There was no obvious difference in the BAG-1 immunohistochemistry of SP-C C-RafBxB/BAG-1+/+ and SP-C C-RafBxB/BAG-1+/- lungs.
Tumour cells of BAG-1 heterozygous mice show increased apoptosis
Concerning the molecular mechanism how a reduction of the BAG-1 protein expression in the heterozygous mice would impair tumorigenesis, we determined the fraction of apoptotic cells. Staining for activated caspase-3 revealed indistinguishable apoptosis in healthy regions of the lung of 1 month old SP-C C-Raf BxB mice with either one or two BAG-1 alleles, in line with the unaltered, normal lung structure of BAG-1+/- mice. In the adenomas, however, we observed a significant increase of apoptotic cells in BAG-1+/- SP-C C-Raf BxB mice compared with their SP-C C-Raf BxB/BAG-1+/+ littermates, see Figure 3a. This mechanism of action of BAG-1 on the regulation of cell survival is compatible with the phenotype of embryonic day 12,5 BAG-1 null embryos. Immunohistochemical staining for activated caspase-3 and trypan blue staining of dissociated cells showed hypocellularity and elevated levels of apoptosis in the livers of BAG-1-/- embryos (unpublished observations).
Proliferation and p-ERK signalling are unaffected in BAG-1 heterozygous mice
To exclude the alternative mechanism that the decreased level of BAG-1 expression in heterozygous animals would cause reduced cell proliferation in the adenomas, we performed proliferating cellular antigen (PCNA) staining. No significant differences were observed in the fraction of proliferating adenoma cells between SP-C C-Raf BxB animals heterozygous or wild type for BAG-1, see Figure 3b. Also, the percentages of adenoma cells positive for Ki-67, another proliferation marker and Bmi-1, a chromatin-associated protein expressed in stem cells, were not affected by the BAG-1 heterozygosity (not shown). Furthermore, staining of lung sections for phosphorylated ERK revealed no quantitative differences in the adenomas of SP-C C-Raf BxB animals heterozygous or wild type for BAG-1, see Figure 3c. Thus, signalling through the mitogenic cascade was not affected by the BAG-1 heterozygosity in the adenoma cells.
Conclusions
Tumours often are highly dependent on signalling pathways promoting cell growth or survival and may become hypersensitive to downregulation of key components within these signalling cascades. This study identifies BAG-1 as a protein specifically required at wild type expression levels for the survival of tumour cells and reveals it as potential anticancer target. Since many key components of survival pathways are regulated by interaction with (co-)chaperones [15], our finding is not without precedent but novel insofar as we have uncovered that reduced BAG-1 expression specifically targets tumour cells to apoptosis and impairs tumorigenesis. Whether this effect on adenoma cell survival requires that BAG-1 interacts with C-Raf or Hsc70/Hsp70 or with both partners requires additional studies. Questions concerning specific roles of the different BAG-1 isoforms were not addressed with this BAG-1 deficient mouse as both isoforms of BAG-1, p50 and p32 are absent in protein extracts of knock-out embryos. Another setting where BAG-1 has a physiological role is the heart, where up-regulation of BAG-1 after ischemia rescues cells from apoptosis [16].
A possible model combining the findings of this report and other data indicates that BAG-1 functions as an activator of C-Raf at the outer mitochondrial membrane where enzymatically activated C-Raf finds apoptosis-related targets such as BAD [17], see Figure 4. We can purify overexpressed C-Raf either in an enzymatically inactive form in a complex with Hsp70 or in an enzymatically active form in a complex with Hsp90/50 (unpublished observations), and BAG-1 is proposed to regulate this activation with ATP generated in the mitochondria. Experiments dealing with this questions are currently ongoing.
Therefore, the therapeutic efficacy of a standard chemotherapeutic agent [13] should be increased dramatically by co-application with a BAG-1 inhibitor, since it would target the adaptability of cancer cells to environmental stress and overcome their genetic plasticity. One way to reduce BAG-1 expression is through use of RNA interference-based gene silencing, in particular as BAG-1 overexpression has been observed in human tumours [11]. Drugs that bind to the ATP binding site of Hsc70/Hsp70 might also be expected to be effective as they would inhibit the interaction of BAG-1 with the ATPase domain of heat shock proteins. Such new specific BAG-1 inhibitors may be identified, aided by the known three-dimensional structure of the BAG domain [18,19].
Competing interests
The author(s) declare that they have no competing interests.
Authors' contributions
RG caried out the molecular and histological studies and participated in the design and co-ordination of the study. BWK carried out the histological and immunohisto-chemical studies. GC participated in the histological and immunohistochemical experiments. URR participated in the design and co-ordination of the study. All authors read and approved the final manuscript.
Pre-publication history
The pre-publication history for this paper can be accessed here:
Acknowledgements
We thank L. Fedorov, N. Gribanow, D. Heim, S. Hilz and T. Potapenko and Y. Yang for support. This work was supported by Deutsche Krebshilfe – Mildred Scheel Foundation (grants 10-1793-Ra7; 10-1935-Ra8) and by the DFG (grants TR17-TPB7; -TPZ2). G. Camarero was a postdoctoral fellow supported by Spanish Government (Ministerio de Educación y Cultura).
Figures and Tables
Figure 1
BAG-1 haplo-insufficiency delays C-Raf driven adenoma growth. (a) Adenoma initiation in SP-C C-Raf BxB mice (BAG-1+/+) and their Bag-1 haplo-insufficient littermates (BAG-1+/-) at 1, 2 and 4 months of age. Adenoma foci values represent mean ± s.e. from at least 4 mice of each genotype analyzed in a blinded fashion by two independent readers. (b) Adenoma area in the lungs of 4 months old mice. Each value represents mean ± s.e. from at least 4 mice of each genotype analyzed in a blinded fashion. (c-d) Examples of hematoxylin-eosin stained sections of lungs from SP-C C-Raf BxB transgenic mice wildtype for BAG-1 in comparison to a BAG-1 heterozygous littermate. Scale bar, 200 μm.
Figure 2
BAG-1 expression in the lung of SP-C C-Raf BxB transgenic mice (a) Lanes 1–8 show immunoblotting data for expression of BAG-1 in the lungs of 8 month old SP-C C-Raf BxB transgenic mice heterozygous (+/-) or homozygous (+/+) for BAG-1 as indicated below the lanes. Lane 9 shows absence of BAG-1 expression in a BAG-1 null (-/-) embryonic day 12,5 liver extract; lane 10 control liver. The markers along the left indicate relative molecular mass. The same blots were subsequently reacted with an antibody against GAPDH to demonstrate protein equal loading and are shown below. (b) BAG-1 immunostaining in SP-C C-Raf BxB transgenic mouse lung cancer tissue.
Figure 3
Increased apoptosis but no change in PCNA and p-ERK in tumor cells of SP-C C-RafBxB transgenic BAG-1 heterozygous mice (a-c) Quantification of immunohistochemical staining for apoptosis using an antibody that detects activated caspase-3 (a), PCNA (b) and phosphorylated ERK (p-ERK, c) of adenoma cells from 1-month-old SP-C C-Raf BxB transgenic mice of the indicated BAG-1 genotype. Each value represents mean ± s.e. from at least 4 mice of each genotype analyzed in three different experiments.
Figure 4
Model for cooperative action of BAG-1 and C-Raf in tumorigenesis A possible model combining the findings of this report and other data is shown. It indicates that BAG-1 functions as an activator of C-Raf at the outer mitochondrial membrane where enzymatically activated C-Raf finds apoptosis-related targets (for details see text). | [
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15615595 | Identification of cardiac malformations in mice lacking Ptdsr using a novel high-throughput magnetic resonance imaging technique
Abstract
Background
Congenital heart defects are the leading non-infectious cause of death in children. Genetic studies in the mouse have been crucial to uncover new genes and signaling pathways associated with heart development and congenital heart disease. The identification of murine models of congenital cardiac malformations in high-throughput mutagenesis screens and in gene-targeted models is hindered by the opacity of the mouse embryo.
Results
We developed and optimized a novel method for high-throughput multi-embryo magnetic resonance imaging (MRI). Using this approach we identified cardiac malformations in phosphatidylserine receptor (Ptdsr) deficient embryos. These included ventricular septal defects, double-outlet right ventricle, and hypoplasia of the pulmonary artery and thymus. These results indicate that Ptdsr plays a key role in cardiac development.
Conclusions
Our novel multi-embryo MRI technique enables high-throughput identification of murine models for human congenital cardiopulmonary malformations at high spatial resolution. The technique can be easily adapted for mouse mutagenesis screens and, thus provides an important new tool for identifying new mouse models for human congenital heart diseases.
Background
Congenital malformations are a major cause of death in childhood, and are typically characterized by lesions that do not compromise fetal survival. For instance, congenital heart disease (CHD) typically consists of lesions such as ventricular and atrial septal defects, which are compatible with fetal hemodynamics [1]. Although human genetic studies have identified some genes that cause congenital cardiac malformations, the molecular and developmental mechanisms underlying most of these defects remain largely unknown. Despite the high incidence of CHD (~1% of live births), only a handful of genes have been identified that when mutated, result in congenital heart disease [2-4].
The mouse is a particularly good model for studying mechanisms of cardiac diseases as its anatomy and development resembles that of the human more closely than any other genetically tractable organism. Importantly, the mouse is amenable to genotype-driven approaches such as transgenic knockouts of defined candidate genes [5], genome wide mutagenesis approaches using gene-trap [6] or transposon insertion screens [7], and high-throughput phenotype-driven screens that rely, for instance, on N-ethyl-N-nitrosourea (ENU) mutagenesis [8,9]. However, high-throughput cardiovascular genomic approaches in the mouse have been hampered by the paucity of phenotyping tools that allow efficient identification of complex cardiac malformations. As mouse embryos are opaque, late developmental defects are particularly difficult to identify. For instance, cardiac septal defects or outflow tract abnormalities can only be confidently identified after 14.5 days post coitum (dpc) when cardiac and outflow tract septation are completed in normal embryos. The identification of malformations in late gestation embryos typically relies on serial histological sectioning, which is extremely labor intensive. Furthermore, this often results in the irretrievable loss of 3D information, which is essential for the interpretation of complex cardiac malformations. In addition, standard pathological analysis is not amenable to high-throughput phenotype screening protocols that are required for any mutagenesis screen aiming at the functional dissection of the developmental biology of cardiac diseases. Therefore, new technological approaches must be harnessed that allow an efficient phenotyping of heart defects and also of subtle cardiac abnormalities that are at danger of being overseen in traditional histopathology screens. This is even more important in the light of upcoming new endeavors in functional mouse genome annotation [10]. Currently, new large-scale mouse mutagenesis screens are being set up in the US and in Europe that aim to produce heritable mutations in every gene in the mouse genome [11-13]. To make full use of these new mouse mutant resources more precise and efficient phenotyping methods are urgently needed [12-14]. The success of genome wide saturation mutagenesis screens depends therefore on improved phenotyping, and new high-resolution imaging approaches for mouse mutants are one of the most important which need to be established.
We previously reported the development of fast gradient-echo MRI of single mouse embryos [15-17]. This resulted in the acquisition of a 3D dataset in under 9 hours, with an experimental image resolution of 25 × 25 × 26 μm/voxel. We showed that MRI is capable of accurately identifying normal embryonal structures, and cardiac and adrenal malformations in knockout mouse embryos, and we have validated this technique by performing in depth histological examinations of imaged embryos [15-17]. These experiments showed that single-embryo MRI could correctly identify all cardiac lesions (atrial septal defects, ventricular septal defects, outflow tract defects such as double-outlet right ventricle, and aortic arch defects) except those under 20 μm – which is below the resolution of the MRI technique. As this method images single embryos in overnight runs, it still lacks the throughput required for phenotype-driven mutagenesis screens. For instance, in a typical recessive ENU mutagenesis screen, to screen 50 ENU mutant lines using a 3-generation breeding scheme would require the analysis of ~1200 embryos [18]. We now report the development of a method of imaging up to 32 embryos simultaneously in a single unattended overnight run, at high spatial resolution. Allowing ~30 minutes per embryo, the analysis of 1200 embryos would take 75 working days for a single trained individual. We show that this high-throughput multi-embryo MRI technique can be used to rapidly identify unsuspected embryonal cardiac and visceral malformations. Using this technique we could identify a novel, and hitherto unsuspected role for the phosphatidylserine receptor (Ptdsr) in controlling ventricular septal, outflow tract and pulmonary artery development. In addition, we found thymus hypoplasia in Ptdsr-deficient embryos. These findings suggest that a novel Ptdsr-mediated pathway is required for cardiac and thymus development.
Results
Multi-embryo imaging
We modified our previously described fast gradient echo magnetic resonance imaging technique [15-17] to image embryos embedded in four to eight layers (16 – 32 embryos total) in 28 mm nuclear magnetic resonance tubes using a single quadrature driven birdcage coil (Figure 1a). Preparation and embedding of embryos typically took less than an hour. In initial experiments we imaged up to 16 embryos simultaneously in overnight runs of <9 hours, with an experimental resolution of 51 × 51 × 39 μm. Subsequently, we used a custom made optimized probe with an increased sensitivity range to enhance imaging throughput. This allowed us to image 32 embryos simultaneously (Figure 1a,b), but with a larger matrix size and an increased field-of-view in the long axis of the tube. For these experiments we imaged embryos for ~12 hours, and achieved an improved experimental resolution of 43 × 43 × 36 μm. The optimized coil used for 32 embryos MRI (Figure 1) has a sensitivity range in z-direction of close to 50 mm. The artefacts seen at both ends in the longitudinal image (Figure 1b) are caused by B1-inhomogeneities at the end-rings of the coil. However, accurate image analysis for the entire data set remains possible if the height of the embryo stack does not exceed approximately 47 mm as demonstrated in the corresponding axial views of the top and the bottom layer (Figure 1c,d). The resolution achieved with the multi-embryo MRI technique allowed us to visualize the heart, cardiac septa, central nervous system, and visceral organs in fine detail (Figure 1d–f), in embryos taken from each layer. The data are permanently archived on DVDs, for subsequent analysis. Construction of 3D reconstructions of the heart typically takes ~4 hours, but was not necessary for the identification of cardiovascular defects.
Figure 1
High-throughput high-resolution magnetic resonance microscopy. (a) Stack of 32 embryos embedded in a NMR tube. (b) Section through the long axis of the NMR tube showing embryos in eight layers. (c) Sagittal section through layer 8 showing the four embryos in this layer. (d–f) Transverse, sagittal, and coronal sections through individual embryos in layers 5, 1 and 4 respectively. The voxel size is 25.4 × 25.4 × 24.4 μm. Structures indicated are the spinal cord (sc), the right and left lungs, atria and ventricles (rl, ll, ra, la, rv, lv), primary atrial and interventricular septa (pas, ivs), mitral valve (mv), midbrain roof (mbr), midbrain (mb), mesencephalic vesicle (mes), thalamus (tha), hypothalamus (hy), pons (po), cerebellum (c), medulla oblongata (mo), pituitary (pit), tongue (t), thymus (th), left superior vena cava and main bronchus (lsvc, lmb), aorta (ao), liver (li), stomach (s), left adrenal and kidney (lad, lk), pancreas (pa), intestines (i), umbilical hernia (uh), aqueduct of Sylvius (aq), fourth ventricle (fv), inner ear (ie), larynx (lar), right ventricular outflow tract (rvot), spleen (sp), and testes (te). Scale bars = 500 μm; axes: d – dorsal; v – ventral; r – right; l – left; a – anterior, p – posterior.
Sensitivity and specificity of multi-embryo imaging
To assess the sensitivity and specificity of multi-embryo imaging in comparison to single-embryo imaging, we used a model of Cited2 deficiency [19]. Embryos lacking Cited2 (Cited2-/-) have diverse cardiac malformations, including atrial and ventricular septal defects, outflow tract and aortic arch malformations, and adrenal agenesis [15,17,19]. As the Trp53-repressor gene Cdkn2aP19ARF is a target of Cited2 [20], we also examined embryos lacking both Cited2 and Trp53 to determine if this would rescue the heart and adrenal defects in Cited2-/- mice. We imaged 50 embryos using the multi-embryo technique in the 16-embryo mode. Embryonal genotypes included 12 wild-type, 13 Cited2+/-, 14 Cited2-/-, three Cited2-/-: Trp53-/-, four Cited2-/-: Trp53+/-, two Trp53+/-, and two Trp53-/-. We analyzed the data from each embryo for cardiac malformations without knowledge of the genotype. This typically took a maximum of 30 minutes per embryo. We scored each embryo for atrial and ventricular septal defects (ASD, VSD), outflow tract (e.g. double-outlet right ventricle, common arterial trunk), and aortic arch malformations (e.g. right-sided or bilateral aortic arch, retroesophageal subclavian artery), and for adrenal agenesis. Each embryo was then re-imaged singly at high resolution, and the data re-analyzed as before. In this group we identified 20 embryos with ASD, 19 with VSD, 18 with outflow tract defects, 11 with aortic arch defects, and 21 with bilateral adrenal agenesis, using high-resolution single embryo imaging. In comparison to single embryo imaging, the overall sensitivity and specificity of multi-embryo imaging for cardiac malformations was 88% and 92% respectively. For ASD (18 identified by single embryo imaging) the sensitivity and specificity was 85% and 95%; for VSD 94% and 94%; for outflow tract malformations 94% and 100%; and for aortic arch malformations 91% and 100% respectively (Figure 2). For bilateral adrenal agenesis, the sensitivity was 100% and specificity was 95% (Figure 3). Embryos lacking both Cited2 and Trp53 had cardiovascular defects and adrenal agenesis, indicating that Trp53 does not play a major role in the genesis of these defects in mice lacking Cited2. These results indicate that multi-embryo MRI is a potentially powerful high-throughput tool for efficiently characterizing cardiovascular malformations and identifying other defects in organogenesis.
Figure 2
Identification of septal, outflow tract, and aortic arch malformations using multi-embryo MRI (a – e') Images of transverse sections from 5 Cited2-/- embryos obtained using the multi-embryo technique (a–e) compared with images from the same embryos obtained subsequently using the single embryo technique (a'–e'). (a, a') Section showing left and right atria and ventricles (la, ram, live, rave). The atria are separated by the primary atria septum (pas), which is deficient at its ventral margin creating an osmium premium type of atria septal defect (ASD-P). (b, b') Section showing a ventricular septal defect (VSD) in the interventricular septum (ivs). (c, c') Section showing double outlet right ventricle, wherein the ascending aorta (a-ao) and the pulmonary artery (pa) both arise from the right ventricle (rv). The aortic valve (ao-v) is indicated. (d, d') Section showing a right-sided aortic arch (ao-a) passing to the right of the trachea (tr) and the esophagus (es). (e, e') Section showing bilateral aortic arches (ao-a) forming a vascular ring around the trachea (tr) and the esophagus (es). Also indicated are the thymus (th) and the right superior vena cava (r-svc). (f – j) Serial transverse sections through a wild-type heart obtained using single embryo MRI, demonstrating corresponding normal structures, including the systemic venous sinus (svs), left superior vena cava (l-svc), pulmonary vein (pvn), descending aorta (d-ao), mitral and tricuspid valves (mv, tv), the secondary atrial septum (sas), left and right ventricular outflow tracts (lvot, rvot), pulmonary valve (pv), and arterial duct (ad) of the pulmonary artery. Scale bars = 635 μm for multi-embryo, and 317 μm for single embryo images; axes: d – dorsal; v – ventral; r – right; l – left.
Figure 3
Identification of adrenal agenesis using multi-embryo MRI Images of coronal sections from 2 embryos obtained using the multi-embryo technique (a, b) compared with images from the same embryos obtained subsequently using the single embryo technique (a', b'). (a, a') Normal right adrenal gland (rad) anterior to the right kidney (rk) in a wild-type embryo. The right lung (rl) is indicated. (b, b') Agenesis of right adrenal gland in a Cited2-/- embryo. Scale bars = 635 μm for multi-embryo, and 317 μm for single embryo images; axes: d – dorsal; v – ventral; a – anterior, p – posterior.
Cardiac malformations in mice lacking Ptdsr
We next evaluated the role of multi-embryo MRI in analyzing unexplained lethality in embryos generated in collaborating laboratories. Recently, we have generated mice lacking the phosphatidylserine receptor (Ptdsr-/-) on a C57BL/6J background, by gene targeting in embryonic stem cells [21]. Ptdsr is a nuclear protein of unknown function, which is essential for the development and differentiation of multiple organs during embryogenesis [21-24]. Ablation of Ptdsr function in knockout mice causes perinatal lethality, growth retardation [21,22,24] and a delay in terminal differentiation of the kidney, intestine, liver and lungs during embryogenesis [21]. In addition, Ptdsr-/- embryos develop complex ocular lesions [21] as well as haematopoietic defects [24]. However, as many malformations have been described in Ptdsr mutants, none of those detected could explain the observed perinatal lethality of Ptdsr-/- mice. In the process of phenotypical characterization of our Ptdsr-deficient mouse line, we frequently observed subcutaneous edema of varying sizes in Ptdsr-/- embryos by gross inspection ([21] and Figure 4). As the development of edema in various mouse mutants is frequently associated with cardiovascular defects [25] we started to investigate if this also holds true for Ptdsr-deficient mice. We examined 8 embryos lacking Ptdsr, and 8 littermate wild-type or heterozygous controls using multi-embryo MRI. We found that 5 of 8 Ptdsr-/- embryos had cardiac malformations, which included ventricular septal defects, double outlet right ventricle, and pulmonary artery hypoplasia (Figure 5). None of the wild-type or Ptdsr+/- embryos had cardiac malformations. These findings were confirmed on single embryo imaging (Figure 6). Furthermore, to verify the identified cardiac defects in the Ptdsr-/- mice we performed serial transverse sectioning of all analyzed embryos. In all cases, we could recognize again the same heart defects that were identified before using the multi-embryo MRI technique (Figure 7). In addition, we analyzed by multi-embryo MRI a second Ptdsr-knockout mouse line (Ptdsrtm1.1 Gbf), which is identical to the initially analyzed Ptdsr mutant except that the loxP-flanked neomycin selection cassette was removed by breeding the original Ptdsrtm1 Gbf knockout line [21] to a CMV-Cre deleter mouse line [26]. From this Ptdsrtm1.1 Gbf knockout mouse line we analyzed 8 Ptdsr-/- embryos, and as littermate controls, 3 Ptdsr+/+ and 1 Ptdsr+/- embryos. We found ventricular septal defects in five out of the eight Ptdsr-/- embryos (data not shown). Again we found no evidence for cardiac malformations in wild-type or heterozygous littermate control embryos.
Figure 4
Edema in Ptdsr-/- mice (a) The Ptdsr-/- mutant (15.5 dpc) is growth retarded and the severe edema along the back of the embryo is visible. (b, c) Sagital sections of embryos at 16.5 dpc. The mutant embryo (c) exhibits massive subcutaneous edema compared to a wild-type (b) littermate. Scale bar = 100 μm in (b) and (c).
Figure 5
Identification of cardiac malformations in Ptdsr-/- embryos using multi-embryo MRI (a–e) Transverse thoracic sections showing the heart of heterozygous or wild-type control embryos from each litter. The left and right ventricles (lv, rv) are separated by the interventricular septum (ivs). The left and right atria (la, ra) are also indicated, separated by the primary atrial septum (pas). (f–i) Corresponding sections through littermate Ptdsr-/- embryos, showing ventricular septal defects (VSD). Scale bar = 635 μm; axes: d – dorsal; v – ventral; r – right; l – left; a – anterior, p – posterior. Individual embryos are indicated by number.
Figure 6
Cardiac malformations and thymus hypoplasia in Ptdsr-/- embryos. (a–c) Transverse and oblique (through the plane of the ascending aorta) sections, and 3D reconstruction (left-ventral oblique view) of a heart of a wild-type embryo at 15.5 dpc. The left and right ventricles (lv, rv) are separated by the interventricular septum (ivs). The left and right atria (la, ra), and the trachea (tr) are also indicated. The ascending aorta (a-ao) arises from the left ventricular outflow tract (lvot), via the aortic valve (ao-v), and continues on as the aortic arch (ao-a), which joins the descending aorta (d-ao). The pulmonary artery (pa) arises from the right ventricular outflow tract (rvot), and continues as the arterial duct (ad), which joins the descending aorta. (d–f) Corresponding images of a Ptdsr-/- embryo, showing a smaller heart with a ventricular septal defect (VSD). The aorta arises from the right ventricle. The pulmonary artery is small and its connection to the descending aorta (arterial duct) could not be identified. (g–i) Corresponding images of another Ptdsr-/- embryo, showing a ventricular septal defect (VSD). The aorta overrides the VSD resulting in a double-outlet right ventricle. (j, k) Coronal sections of Ptdsr+/+ and Ptdsr-/- embryos, showing the two lobes of the thymus (th). The arterial duct of the pulmonary artery in the Ptdsr-/- embryo is narrowed. (l) Correlation between embryo weight and volume. Scattergram of embryo weight versus embryo volume measured from multi-embryo MRI datasets for 16 embryos using Amira. The co-efficient of regression (r) is indicated. (m, n,) Absolute embryo and thymus volumes (μl) were measured from the MRI datasets from 5 wild-type (wt), 3 heterozygote (h), and 8 Ptdsr-/- (m) embryos at 15.5 dpc. There was no significant difference in the wild-type and heterozygote data, which were therefore pooled together (wt/h). (o) Relative thymus volumes (% of embryo volume) were calculated as Ptdsr-/- embryos were slightly smaller than littermate wild-type embryos. The data are represented as mean ± S.D. The probability of a type I error (P) is indicated. Scale bars = 317 μm; axes: r – right; l – left; d – dorsal; v – ventral; a – anterior, p – posterior. Individual embryos are indicated by number.
Figure 7
Cardiac malformations in Ptdsr-/- embryos: analysis using histology Embryos analyzed by MRI (Figure 3) were sectioned transversely and stained with hematoxylin and eosin. (a–c) Serial caudal to cranial sections of the wild-type embryo showing normal cardiac and vascular anatomy. The left and right ventricles (lv, rv) are separated by the interventricular septum (ivs). The ascending aorta (a-ao) arises from the left ventricle, continues on as the aortic arch (ao-a), which joins the descending aorta (d-ao). The pulmonary artery (pa) arises from the right ventricle via the pulmonary valve (pv) and continues as the arterial duct (ad), which joins the descending aorta. The left and right atria (la, ra), trachea (tr), right main bronchus (rmb) and esophagus (es) are indicated. (d) Section through embryo 33 indicating the ventricular septal defect (VSD). (e, f) Sections through embryo 55 showing that both aorta and pulmonary artery arise from the right ventricle (double outlet right ventricle), and that the arterial duct of the pulmonary artery is narrow in comparison to the aorta – indicating pulmonary artery hypoplasia. The aortic valve (aov) is indicated. (g–i) Serial caudal to cranial sections through embryo 35 showing a VSD, aorta arising from the right ventricle (double outlet right ventricle), and a severely narrowed arterial duct. Scale bars = 500 μm; axes: r – right; l – left; d – dorsal; v – ventral. Individual embryos are indicated by number.
We also observed a modest degree of thymic hypoplasia in Ptdsr-/- embryos (Figure 6k). To confirm this, thymus and embryo volumes were measured in 8 Ptdsr-/- embryos and 8 wild-type or heterozygous control embryos at 15.5 dpc. Embryo volume measured from the MRI datasets correlated very strongly with embryo weight (Figure 6l), and was modestly reduced in Ptdsr-/- embryos (Figure 6m). The volume of the thymus was significantly reduced in Ptdsr-/- embryos, even after correction for embryo volume, to 58% of the control value (Figure 6n,o). To correlate the identified cardiopulmonary malformations in Ptdsr-/- embryos with expression of the Ptdsr gene during heart development, we made use of our Ptdsr gene-trap reporter mouse line [21]. Using X-Gal staining in heterozygous embryos staged between 9.5 dpc and 12.5 dpc we found specific Ptdsr expression in the heart starting at 10.5 dpc and getting more defined to the compact zone and the trabeculi from 11.5 dpc onwards (Figure 8). Furthermore, when we analyzed Ptdsr-/- hearts by histopathology at 16.5 dpc we observed a severe differentiation defect in the compact zone as well as in the trabeculi (Figure 9), thus demonstrating that Ptdsr is in addition required for heart muscle differentiation at later stages of development. Taken together these results indicate that Ptdsr plays a hitherto unsuspected role in cardiovascular development as well as in cardiac muscle differentiation.
Figure 8
Analysis of Ptdsr expression in the embryonic heart (a, b) Staining of heterozygous Ptdsr-βgeo-embryos [21] using X-Gal at 10.5 dpc (a) and 11.5 dpc (b). (a) At 10.5 dpc Ptdsr expression can be seen throughout the heart. (b) Transverse sections of X-Gal stained embryos at 11.5 dpc showed an increased expression of Ptdsr in the myocardial wall and a beginning decrease of the expression in the trabeculation. Scale bar = 100 μm in (b).
Figure 9
Myocardial wall malformations in Ptdsr-/- embryos (a, b) Sagital sections of wild-type (a) and homozygous mutant (b) embryos at 16.5 dpc revealed a thinning of the myocardial wall (compact zone) and an increased myocardial trabeculation (b) in the mutant heart. Scale bar = 100 μm.
Discussion
Utility of MRI in identifying mouse models of human malformations
Our results show that it is possible to efficiently identify and quantitate relatively subtle cardiac and visceral malformations in late gestation mouse embryos using multi-embryo MRI. We have deliberately optimized our technique at late gestation in order to identify those congenital defects that allow survival through most of gestation. Importantly, these defects resemble human congenital malformations, and would provide mouse models for the study of these diseases. Our method represents a simpler alternative to the multi-coil approach published recently [27,28] in which up to eight fixed mouse embryos were imaged simultaneously, with a resolution of 200 μm. In comparison, the multi-embryo method described here has a higher experimental resolution of 43 μm. Notably, it requires substantial developmental and financial effort to equip an experimental MR-system with multiple coil and receive capability.
Role of MRI in investigating murine embryonal or perinatal lethality
Many mouse gene knockouts display late gestational lethality, but incomplete analysis and loss of 3D information consequent to histological sectioning, results in major developmental malformations being frequently missed. As shown here, Ptdsr-/- embryos on a C57BL/6J background develop heart defects. This was not noted in recent reports [22,24], and emphasizes the need not only for completely examining several mutant embryos, but also repeating these examinations in different genetic backgrounds. A major advantage of MRI is the ability to easily ship fixed embryos from referring laboratories to the laboratory that performs the MRI analysis. This minimizes the expense of animal relocation, re-derivation, and breeding required to generate the embryos, and significantly reduces animal experimentation.
Role of MRI in high-throughput phenotype driven screens
MRI screens performed at late gestation would be expected to identify genes that affect later aspects of development, and identify hypomorphic and haploinsufficient alleles of genes that affect earlier steps of development. Published data from genome-wide ENU mutagenesis screens in the mouse indicate that ~30% progeny carry a heritable recessive phenotype, making 3-generation recessive screens the method of choice for identifying developmental malformations [18]. At least 24 3rd generation progeny per 1st generation mutant are typically screened, resulting in a >78% probability of identifying at least one fully penetrant recessive homozygous mutant. A typical recessive screen (e.g. 50 – 100 first generation mutants per year) would require the analysis of ~1200 – 2400 embryos per year. Our results show that multi-embryo MRI is eminently suitable for such throughput. Although, screening of 32 embryos overnight requires typically about 16 hours of data analysis, multi-embryo MRI mutagenesis screens can be easily performed at a reasonable scale if multiple operators analyze the data in parallel. As the data are permanently stored on DVDs, and can be analyzed easily by commercially available software, they can be without difficulty disseminated to specialists for further and more detailed analysis. Another powerful application of multi-embryo MRI will likely be the investigation and screening of potentially teratogenic drugs.
Functional role of Ptdsr in heart development
Our results presented here indicate a new, and hitherto unsuspected role for the phosphatidylserine receptor in controlling ventricular septal, outflow tract, pulmonary artery, and thymus development. This finding suggests that a novel Ptdsr-mediated pathway is required for cardiac and thymus development. Recently, we have demonstrated that in contrast to previously reported hypothetical Ptdsr functions, the Ptdsr protein is not required for the clearance of apoptotic cells [21]. Moreover, detailed analysis of apoptosis induction and apoptotic cell clearance in Ptdsr+/+ and Ptdsr-/- embryos during heart development did not reveal any difference in the number and location of apoptotic cells between the genotypes (J.B., A.D.G. and A.L. unpublished observations). This further excludes that Ptdsr has any function in apoptotic cell clearance and points to other developmental mechanisms that are affected by Ptdsr ablation. The neural crest plays an important role in the development of the cardiac outflow tract, aortic arches, and the thymus [29]. As Ptdsr-deficient embryos lack intestinal ganglia [21] which are also derived from the neural crest, these results suggest that Ptdsr-/- mice may have an underlying neural crest defect. Importantly, dysfunction of these Ptdsr-mediated pathways during development could also potentially result in heart defects in humans.
Conclusions
Our results validate the utility of multi-embryo MRI for high-throughput identification of murine models for human congenital cardiac malformations, and using this technique we have shown that Ptdsr is essential for normal cardiac development. Further experiments are needed to define exactly in which pathways Ptdsr is involved during heart development. We expect that multi-embryo MRI will be an important technology for future phenotype-driven mouse mutagenesis screens. The technology can be easily implemented at standard MRI imaging centers, thus allowing by collaboration with individual researchers or mouse mutagenesis centers, a high-throughput functional genetic dissection of mechanisms underlying cardiac development and congenital heart diseases.
Methods
Mice
Cited2-/- [19], Trp53-/- [30], and Ptdsr-/- mice [21] have been described previously. All embryos were harvested at 15 days after detection of the vaginal plug.
Embryo preparation
Embryos were fixed in 4% paraformaldehyde at 4°C for ~1 week, and then embedded in 1% agarose (Seakem) containing 2 mM gadolinium-diethylenetriamine pentaacetic anhydride (Gd-DTPA, Magnevist, Schering UK) in 28 mm nuclear magnetic resonance tubes (Figure 1). The left forelimb was removed from each embryo to facilitate the identification of the left side. In addition, embryos had other limbs and/or tails removed before embedding so that each embryo in a given layer (of four embryos) could be unequivocally identified.
Magnetic resonance imaging
Single embryo imaging was performed as described previously [15-17]. For multi-embryo imaging, we used the same 11.7 Tesla (500 MHz) vertical magnet (Magnex Scientific, Oxon, UK). This was interfaced to a Bruker Avance console (Bruker Medical, Ettlingen, Germany) equipped with a shielded gradient system with a maximal gradient strength of 548 mTesla/m (Magnex Scientific, Oxon, UK), and quadrature-driven birdcage type coils with an inner diameter of 28 mm (Rapid Biomedical, Würzburg, Germany). Compressed air at room temperature was used to reduce the heating induced by the gradients. A 3D spoiled gradient echo sequence (echo time 10 ms), a π/2 excitation pulse with rectangular pulse shape, (π/2 = 100 μs), was used with a short repetition time (30 ms) to obtain strong T1 contrast. A matrix size of 512 × 512 × 768 (bandwidth: 130 Hz/pixel) at a field of view of 26 × 26 × 30 mm achieved an experimental resolution of 51 × 51 × 39 μm when imaging up to 16 specimens. In case of 32 embryos, a matrix size of 608 × 608 × 1408 at field of view of 26 × 26 × 50 mm, yielded an experimental resolution of 43 × 43 × 36 μm. The total experimental time was ~8.75 hours for 16 embryos, and ~12.3 hours for 32 embryos (typically overnight runs) whereby each phase encoding step was averaged four times.
Data reconstruction and analysis
The raw MR data were reconstructed into a stack of 1024 (for 16 embryos), or 2048 (for 32 embryos) 2D TIFF files (16 bit pixel resolution, 2 or 4 GB total size) using purpose-written software as described previously [16]. The TIFF files were analyzed using Amira 3.1(TGS Europe, Mérignac Cedex, France). 3D reconstructions were performed using the Image Segmentation Editor, and tissue volumes for morphometric analysis were measured using the Measure Tissue Statistics tool available in Amira 3.1. The probability (p) of a Type I error was calculated using a 2-sample equal variance 2-tailed t-test in Microsoft Excel.
Histology
Embryos were dehydrated in ethanol, embedded in paraffin wax, and sections were stained with hematoxylin and eosin.
X-Gal staining of embryos
Embryos were dissected free of extraembryonic membranes and then fixed in 4% paraformaldehyde at 4°C. Expression of Ptdsr was detected by staining the embryos overnight in X-Gal according to standard protocols. The embryos were postfixed in 4% paraformaldehyde and processed for documentation or histology.
Authors' contribution
J.E.S. developed the multi-embryo MRI technique, J.B. generated both Ptdsr knockout lines and harvested embryos for MRI and histopathological analysis, S.D.B developed the sample preparation for embryonic MRI, A.D.G. carried out the histopathological analysis of Ptdsr-/- mutants, C.B. prepared the embryos for MRI, K.C. and S.N. assisted the experimental development, S.B. analyzed the MRI and histopathological data, A.L. and S.B. were responsible for the co-ordination of the study and the drafting of the paper.
Acknowledgments
We thank Tim Bardsley and Neil Hoggarth for help with information technology, and Titus Lanz (Rapid Biomedical) for developing the MR probe. These studies were funded by the Wellcome Trust, British Heart Foundation, and the EU project EUMORPHIA (QLG2-CT-2002-00930). S.B. is a Wellcome Trust Senior Fellow in Clinical Science. | [
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15619330 | Dynamic in vivo imaging and cell tracking using a histone fluorescent protein fusion in mice
Abstract
Background
Advances in optical imaging modalities and the continued evolution of genetically-encoded fluorescent proteins are coming together to facilitate the study of cell behavior at high resolution in living organisms. As a result, imaging using autofluorescent protein reporters is gaining popularity in mouse transgenic and targeted mutagenesis applications.
Results
We have used embryonic stem cell-mediated transgenesis to label cells at sub-cellular resolution in vivo, and to evaluate fusion of a human histone protein to green fluorescent protein for ubiquitous fluorescent labeling of nucleosomes in mice. To this end we have generated embryonic stem cells and a corresponding strain of mice that is viable and fertile and exhibits widespread chromatin-localized reporter expression. High levels of transgene expression are maintained in a constitutive manner. Viability and fertility of homozygous transgenic animals demonstrates that this reporter is developmentally neutral and does not interfere with mitosis or meiosis.
Conclusions
Using various optical imaging modalities including wide-field, spinning disc confocal, and laser scanning confocal and multiphoton excitation microscopy, we can identify cells in various stages of the cell cycle. We can identify cells in interphase, cells undergoing mitosis or cell death. We demonstrate that this histone fusion reporter allows the direct visualization of active chromatin in situ. Since this reporter segments three-dimensional space, it permits the visualization of individual cells within a population, and so facilitates tracking cell position over time. It is therefore attractive for use in multidimensional studies of in vivo cell behavior and cell fate.
Background
Macro- and microscopic imaging are pivotal readouts in the field of biology both for determining the normal (baseline) course of events and for observing the effects of experimental perturbations and natural aberrations [1]. Recent advances in microscopic imaging make it possible to routinely gain visual access to samples hundreds of microns thick [2]. The emergence of green fluorescent protein (GFP) as a reporter has opened up many new experimental approaches that were not previously possible [2-4]. GFP and other genetically-encoded autofluorescent protein reporters have a number of properties that make them ideal for multidimentional imaging of living specimens: no substrate (except photons) is required to generate signal, they have a high signal-to-noise ratio, are non-toxic, stable at 37°C and resistant to photobleaching. Moreover they are available in an increasingly large compendium of spectrally-distinct variants.
To construct high-resolution anatomical models of normal, mutant and pathological situations, we must establish technologies to identify and follow individual cells in three-dimensional (3D) space and in 3D over time, in four-dimensions (4D). Unfortunately, native fluorescent proteins permit tracking the position of any given cell over time only if the population of tagged cells is distributed among non-expressing cells by virtue of lineage or in a mosaic experimental situation [5-9]. In situations where groups, or all cells in a 3D field of view express a fluorescent protein label, information on the behavior of individual cells cannot be discerned. Therefore an approach is required where 3D space is segmented at cellular resolution. This is most easily achieved if each cell can be marked with an easily identifiable tag that is visible at subcellular resolution [10,11]. Since it exhibits low autofluorescence, and is a single, universal and volumetrically constrained cellular organelle, the nucleus is ideal for such labeling [12-14].
Our goal was to take advantage of this feature and to develop a non-invasive fluorescent protein marker of the nucleus for in toto imaging (all cells within the multidimensional space being imaged – discussed in Ref. [11]) of individual cells in situ in living mice [10-12]. For the unequivocal identification of individual cells, we sought a developmentally neutral, genetically-encoded autofluorescent protein-based marker that labels DNA during all phases of the cell cycle while preserving cell morphology and behavior.
As the principal structural proteins of eukaryotic chromosomes, histones are attractive targets for fluorescent nuclear labeling. Histone tagged fluorescent protein fusions have previously been shown to incorporate into chromatin without any adverse effects on the viability of cells in culture [15]. When compared to reporters containing nuclear localization sequences (nls), histone fusions exhibit an improved signal-to-noise ratio and have the distinct advantage of signal remaining bound to the target even during cell division when the nuclear envelope has broken down. In contrast nls-tagged markers (both GFP and lacZ) become dispersed throughout a cell during division, making it difficult to distinguish individual cells during mitosis.
To date GFP fusions to several histones have been generated and used for labeling nuclei in live transgenic animals, including nematode worms, fruit flies and zebrafish [13,14,16,17]. One of these is a fusion between EGFP and human histone H2B which was developed in order to label active chromatin and used to follow the segregation of double minute chromosomes in cancer cells [15]. We have investigated the expression and germline transmission of this type of fusion in mice and established its usefulness not only for imaging cell cycle dynamics [18], but also for tracking cells in living specimens. Moreover unlike native GFP variants this subcellularly localized histone fusion was found to withstand fixation while retaining both fluorescence and subcellular localization.
Results
To evaluate histone-tagged fluorescent protein fusions in embryonic stem (ES) cells and mice, we generated constructs comprising an N-terminally positioned human H2B sequence followed at the C-terminus by sequences for various fluorescent proteins both GFP and DsRed-based. We previously reported that DsRed1 was not amenable to use in ES cells or mice [22], however several improved DsRed variants have recently become available [10]. We therefore chose to evaluate DsRed2 and DsRedExpress as part of this study.
The H2B-fluorescent protein fusions we generated were introduced into vectors utilizing the CAG promoter [19] designed to drive high-level constitutive gene expression in ES cells, embryos and adult mice [20]. Standard protocols were used to establish stable lines of ES cells constitutively expressing an H2B fusion [20-22]. Several transgenic ES cell lines were generated each expressing H2B-EGFP at strong homogenous levels [23]. However, even though we did recover lines with H2B-DsRed2 and H2B-DsRedExpress expression [24], subsequent maintenance of these lines in culture revealed a continued reduction and heterogeneity in fluorescence. We were unable to establish lines with sustained homogenous H2B-DsRed2 or H2B-DsRedExpress fluorescence. Moreover our recent data suggest that mRFP1 [10,25], a rapidly-maturing monomeric form of DsRed, is amenable to use in mice, both in its native form and as a part of functional fusion proteins (AKH unpublished observations).
H2B-EGFP expressing ES cells are shown in Fig. 1. It noteworthy that with this histone fusion we observed a high signal-to-noise ratio and so could achieve high-resolution imaging of mitotic chromosomes (pink arrowheads), various states of interphase chromatin and nuclear debris (yellow arrowheads). Moreover for cells undergoing mitosis we could also discern the stage of mitosis and the plane of cell division (Fig. 1b inset). Previous work indicated that a similar fusion protein expressed in HeLa cells did not affect cell cycle progression [15], and accordingly not only could we visualize nuclear dynamics and identify the various phases of mitosis in live ES cells [26] (Fig. 2), but in doing so, we did not observe any change in growth rate or mitotic index in the transgenic ES cells compared to non-transgenic parental ES cells (data not shown). By imaging several CAG::H2B-EGFP transgenic ES cells undergoing mitosis (n = 30) we calculated the progression from early prophase to cytokinesis to take less than one hour (Fig. 2). Furthermore imaging of embryoid bodies demonstrated that individual nuclei could be discerned from a three-dimensional population of densely packed cells all of which were expressing the H2B-EGFP marker (Fig. 1c). No loss of fluorescence was observed with prolonged in vitro passage of the ES cells expressing the H2B-EGFP fusion in the absence of positive selection in the presence or absence of LIF (t > 3 months in the presence of LIF).
Figure 1
Imaging chromatin in living transgenic ES cells constitutively expressing a H2B-EGFP fusion protein. (a) Bright-field and (b) dark-field micrographs of a CAG::H2B-EGFP ES cell colony. The inset shows a detail with three nuclei in metaphase (pink arrowheads) with the metaphase plates orientated differently. The mitotic spindle of the cell at the top is closely aligned to the z-y plane whereas those for the lower two cells are more closely aligned with the x-z planes. (c) Rendered stack (3-D reconstruction) of sequential optical slices acquired using spinning disc confocal methodology, projected as a fixed angle view of an embryoid body comprised of ES cells constitutively expressing a H2B-EGFP fusion. Pink arrowheads indicate two nuclei in late-anaphase – telophase. Yellow arrowhead points to the nuclear remnant of a cell that has necrosed or apoptosed. (d – f) High-power sequential optical sections each (1 μm apart) through ES cells constitutively expressing the H2B-EGFP fusion, taken using laser scanning confocal methodology showing interphase nuclei, a mitotic nucleus (pink arrowhead) and a pycnotic nucleus (yellow arrowhead).
Figure 2
Live imaging the progression through mitosis. Laser scanning confocal x-y images taken at a single z-plane at five minute intervals for one hour. Note that not all green fluorescence (corresponding to nuclear material) will be represented in the plane being imaged. A cell progressing from anaphase to cytokinesis (pink arrowheads). A cell progressing from prophase to telophase (blue arrowheads). The average time taken to transition from early prophase to cytokinesis was calculated to be approximately 1 hour (n = 30).
We next tested the effects of widespread expression of an H2B fusion protein in mice. We generated germ line chimeras and established transgenic lines of mice constitutively expressing H2B-EGFP. We were able to breed this transgene to homozygosity, resulting in viable and fertile animals exhibiting widespread expression with no overt morphological abnormalities. The transgene has been maintained for over three years in a breeding colony of homozygous mice with no apparent effect on viability, breeding performance or lifespan. We therefore infer that this fusion protein is developmentally neutral and does not interfere with either mitosis or meiosis. Wide-field microscopic analysis of both mouse embryos and adult organs demonstrates widespread expression of the H2B-EGFP fusion in all types of nucleated cells.
We used laser scanning confocal microscopy [10,11] to image this constitutively expressed transgenic reporter at subcellular resolution in live mouse embryos. Such non-invasive visualization of chromatin in living preparations allowed us to acquire high-magnification sequential optical sections (z-stacks) that can be used to generate high-resolution anatomical volumetric (3-dimensional) images with details of interphase chromatin in addition to mitotic chromosomes and fragmenting nuclei. To do this, stacks of sequential optical sections are reconstructed into 3-dimensional projections. This methodology can be used to generate 3-dimensional (3D) image sets not only of cells propagated in culture but also of cells in situ in living animals and is illustrated here by imaging whole mouse embryos at the 4-cell stage, the blastocyst stage, and the pre-gastrula stage (Fig. 3 and Additional Files 1 and 2). These data sets can be computationally manipulated in various ways, for example for the visualization of individual xy slices from a z-stack, rendered images from the full, or part of a z-stack, and color-coded depth projections of a z-stack (Fig. 3).
Figure 3
Live embryo imaging of preimplantation and early postimplantation mouse embryos hemizygous for a constitutively expressed H2B-EGFP fluorescent fusion. (a) Single confocal optical section fluorescence overlay on a bright-field image of a 5-cell stage pre-implantation embryo. Two of the blastomeres are dividing synchronously and are in metaphase (pink arrowheads in b). (b) Dark-field projection of the entire rendered z-stack of x-y sections (n = 19), through the entire embryo shown in panel a. (c) Color-coded depth projection of the entire z-stack of x-y images for the embryo shown in the previous panels. (d) Single confocal optical section fluorescence overlay on a bright-field image of a blastocyst stage embryo. Inner cell mass (ICM) is to the top left corner and second polar body is on the bottom left, juxtaposed to the edge of the ICM. (e) Dark-field projection of half the rendered z-stack of x-y sections (n = 40, sections 1–19 were used for generating the projection), spanning half the embryo shown in panel d. Condensed chromosomes of nuclei in prophase (pink arrowheads) can be seen in three cells of the mural trophectoderm. Cells of the polar trophectoderm (green arrowhead) and inner cell mass (blue arrowhead) can also be distinguished by position within the half-blastocyst reconstruction. (f) Color-coded depth projection of the entire z-stack of x-y images for the embryo shown in the previous two panels. (g-h) Saggital views and rendered z-stacks of x-y images of an E5.75 (pre-streak stage) embryo. (g) Single optical confocal section fluorescence overlay on a bright-field image positioned half the way through the embryo. The brackets on the left illustrate the position of the embryonic (Em) and extraembryonic (Ex) regions of the embryo. (h) The same optical section with only the fluorescence image. Cells of the epiblast (blue arrowhead) and visceral endoderm (green arrowhead) can clearly be distinguished on the basis of position and nuclear morphology. Cells in mitosis can readily be distinguished within the embryo (pink arrowhead). (i) Color-coded depth projection of the stack of serial sections (n = 60), part of the series of which is shown in the previous two panels. Color-coded z-scale (upper right) applies to all projections and denotes distances along the z-axis (0–120 μm).
Data on older embryos and adult organs illustrates that larger specimens can be imaged, however not in their entirety given current limitations in optical imaging capabilities. Instead of imaging the whole specimen, larger samples are positioned so that data can be acquired from regions of interest, which can then be acquired in a tiled manner and computationally re-aligned in image acquisition and processing software.
Our data demonstrates that nuclear morphology afforded by the H2B-EGFP fusion can be used to identify different cell types. In both the raw data, and a rotated rendered stack of an embryonic day (E) 7.5 embryo, cells of the definitive endoderm, mesoderm and embryonic ectoderm can be distinguished solely on the basis of nuclear morphology and orientation in addition to their expected position (Fig. 4b–h and Additional File 3). Low magnification rendered z-stacks taken from a transversely cut section through the head of an E10.5 embryo (Fig. 4i) reveal the stereotypical 3D organization of nuclei within the region imaged (Fig. 4j), and electronically magnified views of this image illustrate a characteristic apposition of nuclei both in and around the notochord, and within the mesenchyme and endoderm of the pharyngeal region (Fig. 4k and 4l and Additional Files 4 and 5), in addition to providing information on cell division and cell death (pink and yellow arrowheads, respectively in Fig. 4l).
Figure 4
Live imaging H2B-EGFP in postimplantation mouse embryos. (a) Lateral view of the embryonic region of an E7.5 embryo (anterior to the left) with box depicting the region imaged in b and double-headed arrow depicting the x-y layering of the z-stack. (b-d) single optical x-y sections of fluorescence overlayed on bright-field images acquired at the same focal plane. Each panel is 60 μm apart from the preceding panel. These panels comprise x-y images in the z-stack depicted in panel a. The different layers of this stage of embryo including the epiblast, mesoderm, visceral endoderm and node can be distinguished on the basis of both position and nuclear morphology. (e-h) projection of a rendered z-stack of (x-y) sections (n = 90) of the dark-field component of the sections taken in the series schematized in a and of the raw data shown in b. (e) 0° rotation, (f) 60° rotation, (g) 120° rotation, and (h) 180° rotation views. (i) low-magnification frontal view of an E11 embryo that has had a transverse cut made to remove the head. The box depicts the region (at the ventral hindbrain and 1st branchial pouch) subject to laser scanning confocal imaging, with the double-headed arrow depicting the x-y layering of the acquired z-stack. (j) rendered (z-) stack of sections (n = 200, i.e. 400 μm depth) taken through the boxed region. (k) rendered stack of top 50 x-y sections (100 μm depth) taken from the region imaged around the notochord (comprising axial mesoderm and mesenchyme cells). (l) rendered stack of top 50 sections (100 μm depth) taken around the branchial pouch region (comprising endoderm and mesenchyme cells). The sections used to generate the rendered stacks in panels k and l were electronically magnified. Pink arrowheads, mitotic nuclei; yellow arrowheads, pycnotic nuclei; ect, ectoderm, en, endoderm, hf, headfold, mes, mesoderm, noto, notochord.
Wide-field microscopic examination of organs from adult animals revealed widespread fluorescence as has been reported for animals expressing native fluorescent proteins under the regulation of the CAG promoter [20,22,27]. Laser scanning confocal imaging of various organs obtained from adult animals was used to generate high-resolution images revealing stereotypical nuclear positions, reflecting different cell types and revealing other subcellular details, such as mitosis and nuclear fragments, also observed in embryos (Fig. 5 and Additional Files 3, 4, 5).
Figure 5
High resolution live imaging of the organs of CAG::H2B-EGFP adult mice. Confocal images of freshly isolated organs from a 6 week old adult male hemizygous CAG::H2B-EGFP Tg/+ animal illustrate the widespread nuclear localized expression of the histone fusion. A transverse cut was made through each organ and the cut surface was placed closest to the objective lens and imaged. Cell tracker orange was used as a vital cytoplasmic counter stain. The panels show rendered confocal z-stacks imaged through 80 μm of the brain using a 20x plan-apo objective (a-c), 568 μm of the heart using a 5x fluar objective (d-f), 142 μm of a lung lobe using a 5x fluar objective (g-i) and 346 μm of a kidney using a 5x fluar objective low power view (j-l), and high power view (m-o). Insets in panels a and d show the region of the brain and heart imaged, respectively. High resolution images of the kidney (m-o) illustrate electronic magnification of the data shown in j-l. Bron, bronchus; glom, glomeruus; med, medulla; sept, septum; ub, ureteric bud; ven, ventricle. Areas of increased fluorescence in the red channel are an artefact due to saturated pixels in regions of the sample closest to the objective.
Finally we investigated whether we could follow cell movement, division, and death in time-lapse experiments using various imaging modalities. We cultured ES cells and embryos on the stages various each of which had been modified to permit culture under physiological conditions. The different types of data routinely generated using different optical imaging modalities that are widely used and commercially available are illustrated in Figure 6. Spinning disc confocal microscopy [1] was used for short-term high-resolution 4D imaging of CAG::H2B-EGFP ES cells (Fig. 6 and Additional File 6), wide-field microscopy was used for long-term low-resolution imaging of CAG::H2B-EGFP preimplantation stage embryos (Fig. 6 and Additional File 7). Note that development proceeds normally in most embryos, and that some of the embryos imaged are undergoing cavitation to form blastocysts [28] (arrowheads). Two-photon excitation microscopy [10,29,30] was used to image cells in a whole gastrula-stage mouse embryo without perturbing the morphogenetic movements associated with gastrulation (Fig. 6 and Additional File 8). Cells can clearly be followed through the successive time points in each of these experimental situations ranging from a few minutes (short-term) to 24 hours (long-term) time-lapse duration. These studies reflect the range of resolutions at which information can be acquired using a marker of this type. We observed normal cell proliferation throughout the course of these imaging experiments and no excessive nuclear fragmentation. Also, because the on-stage cultures were comparable to parallel cultures maintained in a tissue culture incubator, we conclude that the outcome of the cultures was not affected by the various imaging modalities.
Figure 6
Dynamic time-lapse imaging of mouse CAG::H2B-EGFP transgenic ES cells, preimplantation and postimplantation embryos using different imaging modalities. (a) Rendered confocal stacks of transgenic ES cells constitutively expressing a CAG::H2B-EGFP transgene representing a 25 minute time-lapse recording of images acquired using a spinning disc confocal scan head. x-y sections with a z-interval of 0.2 μm were taken at a rate of 10/second over a total z-stack of 40 μm. Cells can be traced through the 4D rendered stack. Cells entering or completing mitosis (pink arrowheads) and the nuclear remnant of a cell that has either undergone apoptosis or necrosis (yellow arrowhead) are clearly visible. (b) Wide-field imaging of CAG::H2B-EGFP transgenic preimplantation embryos. This 24 hour image sequence illustrates cavitation leading up to the formation of the blastocyst in several embryos (violet arrowheads). (c) Rendered two-photon stacks of CAG::H2B-EGFP transgenic gastrulation stage postimplantation embryos. This 40 minute time-lapse sequence illustrates cell division and tracking within the visceral endoderm (green arrowhead) and epiblast (blue arrowheads) and the movement of mesoderm emanating from the primitive streak, which is positioned to the right, out of the field of view. Scale bar in a = 10 μm, b = 100 μm and c = 50 μm.
Discussion
Here we report the evaluation of a chromatin localized histone fusion fluorescent reporter in vivo through the generation of transgenic embryonic stem (ES) cells and mice having widespread expression of this reporter.
The transgenic mice that we have generated provide a new tool for high-resolution live imaging of a genetically tractable mammalian model organism [10,11]. They represent a resource for analyzing development and disease at the subcellular level in cells, embryos and adult tissues. The marker used facilitates the acquisition of in vivo data and allows it to be integrated onto a high-resolution anatomical framework. This type of multidimensional data is complex and thus difficult to digitize and compile into a standardized and integratable format. In toto imaging of fluorescent protein expressing specimens on a large-scale could be used for generating high-resolution digitally recorded anatomical databases where the baseline (wild-type) cell behaviors and cell fates can be contrasted to those observed in mouse mutants. However, developing in toto imaging technologies for acquiring large amounts of data will necessitate improving the speed and throughput of microscopic image acquisition and analysis. This would also be coincident with the ongoing development of improved computational approaches to mine and integrate this type of data [discussed in ref. [11]].
Much of the information generated using a fluorescent fusion reporter such as the H2B-EGFP fusion is analogous to conventional histology [21,31] except that this mode of data acquisition optically sections a sample (circumventing the need to physical section), excels in permitting computational 3D reconstructions of spatial information, and can additionally be coupled to time-lapse imaging for the capture, processing and quantitation of 4D information (Fig. 6 and Additional Files). Also, unlike conventional GFP-based reporters [20,22], the histone H2B-EGFP fusion is resilient to fixation, so samples can be processed and stored for extended periods of time without compromising signal integrity or specificity (Fig. 7).
Figure 7
High-resolution 3-dimensional imaging of fixed CAG::H2B-EGFP transgenic embryos. Confocal images of an E8.5 CAG::H2B-EGFP transgenic embryo fixed in 4% paraformaldehyde for 72 hours, then washed, stored and imaged in PBS. Low-magnification views and reconstructions of whole embryo (a-c). Boxes in a designate region imaged in d and g. High-magnification views of the headfolds (d-f) and posterior primitive streak and proximal allantois (g-i). Single xy images (a, d and g) from the z-stacks used to computationally render the data sets. These images are overlayed onto the bright field channel so as to display the outline of the embryo. Rotations through the rendered z-stacks displayed at 45° intervals (b, e and h). Color-coded depth projections of each of the z-stacks (c, f and i).
The future development and availability of mouse strains constitutively expressing spectral variant histone H2B fusions should prove useful for visualizing anatomy and tracking different populations of cells in multiple dimensions at high-resolution in mice as has previously been demonstrated in other organisms which are classically perceived as being more amenable to in vitro culture and optical imaging [13,16]. They can also be used as tagged populations of cells in chimeras [8], in addition to transplantation and cell isolation experiments [32]. Also, since fluorescence is proportional to genome content and the fluorescence intensity reflects chromosome condensation state, the reagents we have generated should permit the study of alterations in ploidy and chromosomal condensation including determination of phases of mitosis [26]. Additionally, real-time analysis of chromatin fragmentation as well as the effects of mutations on chromosome stability during disease processes can be investigated using CAG::H2B-EGFP transgenic mice.
Conclusions
The CAG::H2B-EGFP strain that we have generated takes in vivo imaging using genetically-encoded reporters in mice to sub-cellular resolution. The development of additional strains permitting spectrally-distinct high-resolution live cell in vivo imaging, coupled to advances in optical imaging modalities and the development of improved computational methods to mine imaging data should pave the way for a multidimensional understanding of biological processes.
It is anticipated that in the future, in vivo imaging approaches using transgenic animals expressing genetically-encoded fluorescent proteins will not only provide high-resolution information on cell behavior in specific biological processes [12,33], but more importantly it may lead to an exponential increase in the available multidimensional in vivo biological information which could mirror the recent explosion of available genomic data. Therefore recent advances in live imaging will need to be paired with developments in computational biology, as appropriate informatics methods will need to be developed and implemented in order to mine, present and integrate this type of in vivo biological data.
Methods
The coding sequence for the human histone H2B gene (X57127) was amplified from genomic DNA by PCR using Pfx Polymerase (Invitrogen). The resulting product was cloned into pCR4 TOPO (Invitrogen) to generate pH2B. The H2B fragment was then cloned into plasmids pEGFP-N1, pDsRed2-N1 pDsRedExpress-N1 (BD Biosciences, Inc) in order to generate plasmids pH2B-EGFP, pH2B-DsRed2 and pH2B-DsRedExpress (oligonucleotide sequences are available upon request). The resulting fusions were then re-amplified by PCR and cloned into the XhoI site of pCAGGS [19] to generate pCX-H2B-EGFP, pCX-H2B-DsRed2 and pCX-H2B-DsRedExpress. All vectors were tested by transient transfection of Cos-7 cells and R1 ES cells using Fugene 6 Transfection Reagent as per manufacturer's recommendations (Roche) and electroporation respectively. The H2B-DsRed2 and H2B-DsRedExpress fusions failed to produce sustained homogenous levels of fluorescent signal, however the H2B-EGFP fusion gave strong nuclear-localized fluorescence throughout the extended culture period without perturbing cell morphology, the rate of proliferation, or the mitotic index (Fig. 2 and data not shown).
Transgenic ES cell lines constitutively expressing H2B-EGFP were generated by co-electroporation of the linearized reporter construct and a circular PGK-Puro-pA plasmid [34] conferring transient puromycin resistance. Puromycin selection was carried out as described previously [20,22]. Briefly, drug selection was initiated 24–36 hours after electroporation, maintained for 5 days, after which time it was replaced with non-selection media for a further 24–48 hours. Fluorescent colonies were identified and picked under an epifluorescence microscope (Nikon SMZ1500). Clones were passaged in 96-well plates, and scored for maintenance and extent of fluorescence. Those exhibiting homogeneous and robust transgene expression in vitro under both stem cell conditions and conditions employed to promote their differentiation were maintained further. For stem cell conditions ES cells were grown on gelatin in the presence of LIF. For differentiation, ES cells were grown on bacteriological Petri dishes in the absence of LIF for 2–5 days to promote embryoid body formation. Thereafter embryoid bodies were re-plated onto tissue culture dishes in the presence of factors promoting directed differentiation.
To assess whether an H2B fusion can continue to be widely expressed and transmitted through the germline of mice we used H2B-EGFP expressing ES cells for chimera generation by injection into C57BL/6 blastocysts using standard procedures [21]. Chimeras were bred to outbred ICR and inbred 129/Tac mice (Taconic, Germantown, NY) for germline transmission and subsequent maintenance of the lines. Two independent clones were taken germline giving indistinguishable results. We therefore focused on one of the transgenic lines. After germline transmission, this transgene was maintained at homozygosity, suggesting that the site of integration is not perturbing essential gene function. All animals retained widespread homogenous fluorescence for at least five subsequent generations. Homozygotes were distinguished from heterozygotes either by increased fluorescence in newborn (unpigmented) animal tails, by breeding, or by intensity of an EGFP hybridizing fragment on a Southern blot.
Embryos and organs were dissected in HEPES buffered DMEM media containing 10% fetal calf serum, then cultured either in a standard tissue culture incubator or on a microscope stage under standard conditions promoting the culture of mouse embryos [35,36] in 50% rat serum: 50% DMEM buffered with bicarbonate and maintained under physiological conditions in a closed temperature-controlled, humidified and oxygenated (95% air, 5% CO2) chamber (Bioptechs Inc. or Solent Sci Ltd. or home-made). For cytoplasmic staining, samples were incubated in Cell Tracker Orange (Molecular Probes; 1:500 dilution in dissecting or culture media) for 10–20 minutes. Embryos were kept in a standard tissue culture incubator at 37°C during staining. Samples were then washed twice with warm dissecting or culture media prior to imaging.
All images shown (except Fig. 7) are of living hemizygous (Tg/+) embryos or freshly dissected (unfixed) tissues obtained from Tg/+ adults and maintained under physiological conditions. Increased fluorescence and a higher signal-to-noise ratio was observed in homozygous (Tg/Tg) specimens. The embryo presented in Figure 7 was fixed in 4% paraformaldehyde at 4°C for 72 hours, then washed, stored and imaged in PBS. Similar results were obtained in embryos fixed for up to two weeks.
Wide-field images were acquired on a Nikon SMZ1500 stereo-dissecting microscope or Nikon Eclipse 5000 inverted microscope equipped with epifluorescent illumination. Spinning disc confocal data was acquired on an UltraView RS3 (Perkin-Elmer Systems) fitted on a Zeiss Axiovert 200M microscope with illumination from a 488 nm Argon laser (Melles Griot). Laser scanning confocal and multiphoton excitation data were taken on a Zeiss LSM510 NLO on a Zeiss Axioscop 2 FS MOT microscope. Objective lenses used on the Axiovert 200M and Axioscop 2 were plan-apochromat 63x/NA1.4, C-apochromat 40x/NA1.2, a plan-apochromat 20x/NA0.75 and a fluar 5x/NA0.25. For laser scanning microscopy GFP was excited using either a 488 nm Argon laser (Lassos, Inc) at 488 nm (for single-photon excitation) or a Titanium:Sapphire laser (Coherent Mira 900F with Verdi 5W pump laser) tuned between 860 and 890 nm (for two-photon excitation). Cell Tracker Orange was excited using a 543 nm HeNe laser (for single-photon use).
Images were acquired as z-stacks comprising sequential x-y sections taken at 0.1–2 μm z-intervals. Raw data was processed using a variety of packages including Zeiss AIM software (Carl Zeiss Microsystems at ), Image J (NIH at ) and Volocity (Improvision at ). Each image series was re-animated using software to make the time-lapse movies that are available as additional files. Both rendered volume and time-lapse movies were assembled in QuickTime Player (Apple Computer, Inc at ).
Appendix
The CAG::H2B-EGFP strain of mice generated in this study will be made available through the Jackson Laboratories Induced Mutant Resource (JAX IMR at ).
Supplementary Material
Additional File 1
Rotating 3D projection of a whole live CAG::H2B-EGFP Tg/+ blastocyst. Supplementary to stills presented in Fig. 3. This file was assembled at 6 frames/second.
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Additional File 2
Rotating 3D projection of a (electronic) half blastocyst. Generated from the same raw data set used to compile Supplementary File 1. Supplementary to stills presented in Fig. 3. This file was assembled at 6 frames/second.
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Additional File 3
Rotating 3D projection the node region of a live CAG::H2B-EGFP Tg/+ E7.5 embryo. Supplementary to stills presented in Fig. 4. This file was assembled at 6 frames/second.
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Additional File 4
Rotating 3D projection of the notochord of a live CAG::H2B-EGFP Tg/+ E10.5 embryo. Supplementary to Fig. 4. This file was assembled at 6 frames/second.
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Additional File 5
Rotating 3D projection of the branchial region of a live CAG::H2B-EGFP Tg/+ E10.5 embryo. Supplementary to stills presented in Fig. 4. This file was assembled at 6 frames/second.
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Additional File 6
Time-lapse sequence of CAG::H2B-EGFP Tg/+ ES cells. Supplementary to stills presented in Fig. 6. This file was assembled at 12 frames/second.
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Additional File 7
Time-lapse sequence of CAG::H2B-EGFP Tg/+ preimplantation embryos. Supplementary to stills presented in Fig. 6. This file was assembled at 6 frames/second.
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Additional File 8
Time-lapse sequence of a CAG::H2B-EGFP Tg/+ postimplantation embryo. Supplementary to stills presented in Fig. 6. This file was assembled at 6 frames/second.
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Additional File 9
Rotating 3D projection of a fixed CAG::H2B-EGFP Tg/+ E8.5 embryo. Supplementary to stills presented in Fig. 7. This file was assembled at 6 frames/second.
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Additional File 10
High-magnification rotating 3D projection of the headfold region of the same fixed CAG::H2B-EGFP Tg/+ E8.5 embryo as shown in Movie 7. Supplementary to stills presented in Fig. 7. This file was assembled at 6 frames/second.
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Additional File 11
High-magnification rotating 3D projection of the allantois and posterior tail region of the same fixed CAG::H2B-EGFP Tg/+ E8.5 embryo as shown in Movie 7. Supplementary to stills presented in Fig. 7. This file was assembled at 6 frames/second.
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Acknowledgements
We thank J.-I. Miyazaki for the pCAGGS plasmid, and T. Swayne at the Herbert Irving Comprehensive Cancer Center Optical Microscopy Facility for instruction and assistance with laser scanning microscopy data acquisition and processing. This work was supported by NIH grants GM60561 and HD33082 (VEP). AKH was a fellow of the American Heart Association during part of this work. | [
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15784609 | Conditional and inducible transgene expression in mice through the combinatorial use of Cre-mediated recombination and tetracycline induction
Abstract
Here we describe a triple transgenic mouse system, which combines the tissue specificity of any Cre-transgenic line with the inducibility of the reverse tetracycline transactivator (rtTA)/tetracycline-responsive element (tet-O)-driven transgenes. To ensure reliable rtTA expression in a broad range of cell types, we have targeted the rtTA transgene into the ROSA26 locus. The rtTA expression, however, is conditional to a Cre recombinase-mediated excision of a STOP region from the ROSA26 locus. We demonstrate the utility of this technology through the inducible expression of the vascular endothelial growth factor (VEGF-A) during embryonic development and postnatally in adult mice. Our results of adult induction recapitulate several different hepatic and immune cell pathological phenotypes associated with increased systemic VEGF-A protein levels. This system will be useful for studying genes in which temporal control of expression is necessary for the discovery of the full spectrum of functions. The presented approach abrogates the need to generate tissue-specific rtTA transgenes for tissues where well-characterized Cre lines already exist.
INTRODUCTION
Cellular diversity in a developing organism is achieved by spatial and temporal regulation of gene expression determined by genetic programs and inductive signals. As a consequence, genes have spatially and temporally acting multiple functions, which are difficult to dissect without proper genetic tools providing similar complexity of control of expression—disruption or overexpression of a gene of interest (1,2). Uncontrolled transgenic expression of a given gene in all tissues or even in restricted cell types may be associated with embryonic lethality that precludes further studies at later stages of development or in adults.
Cell type-/tissues-specific mutation of a gene can be achieved by a two-step process; introduction of loxP sites around a functionally essential genomic part followed by a cell type-specific Cre recombinase-mediated excision of the loxP flanked sequence. The same strategy can be used for cell type-specific overexpression of a transgene, when a strong overall expressing promoter is separated from the coding region of a gene of interest by loxP flanked ‘STOP’ sequences (1,3–5). In both scenarios, a Cre recombinase transgene provides spatial control. However, once Cre expression has been switched on and recombination has occurred, the resultant change in gene expression is, in most cases, irreversible. Tetracycline-inducible transgenic systems [tetracycline transactivator (tTA) or ‘Tet-Off’ and reverse tetracycline transactivator (rtTA) or ‘Tet-On’] allow for reversible temporal regulation of transgene expression (6,7). Of the two systems, rtTA is better suited for rapid induction of gene expression.
In the following report, we describe a novel system that allows for the spatial and temporal regulation of transgene expression in vivo by combining any of the existing Cre recombinase transgenic lines with the reverse tTA (rtTA)–tet-O system. We targeted rtTA into the widely expressed ROSA26 locus, in a way where rtTA expression is conditional to a Cre-mediated excision event (8). Furthermore, we demonstrate the tightness and inducibility of this approach by systemic and cell type-specific (neuronal and podocyte) inducible expression of a dosage-sensitive gene, VEGF-A (9). The presented mouse line can be used to achieve spatially and temporally controlled transgene expression in a wide variety of settings simply by crossing to any existing mice carrying cell type-specific Cre recombinase and tet-O-regulatable responder genes.
MATERIALS AND METHODS
DNA constructs
An EcoRI fragment carrying the rtTA coding sequence and an N-terminal nuclear localization signal was excised from the plasmid pSPC-rtTA (10), blunted by Klenow enzyme and inserted into the pCALL2-IRES-EGFP plasmid (a gift from Dr Corrine Lobe) at a blunted XhoI site. The resultant vector (pCALL2-rtTA-IRES-EGFP) was first digested with BglII, filled in by Klenow polymerase and then cut with NotI, and the fragment containing rtTA-IRES-EGFP was isolated. The pBigT vector (11) was digested first with SalI, filled in and subsequently digested with NotI. The above fragment was inserted into this vector resulting in pBigT-rtTA-IRES-EGFP. The whole insert of the latter vector was released by AscI–PacI double digest and inserted into pROSA26-PA vector (11) digested by the same enzymes. The resultant target vector was named pROSA26-rtTA (Figure 1A). Before introduction into mouse embryonic stem (ES) cells, the vector was linearized with XhoI.
The murine podocin promoter was amplified from genomic murine DNA as described previously (12) and cloned upstream of the NLS-Cre transgene (a gift from B. Sauer).
ES cell manipulation
R1 mouse ES cells (13) were maintained and manipulated as described elsewhere (14). In brief, 20 μg of linearized target vector was electroporated into 107 cells using 500 μF and 250 V settings on Gene Pulser (Bio-Rad). After 24 h, the cells were selected for neomycin resistance in G418 (170 μg/ml) for 8 days. The resistant colonies were picked, expanded and split in 96-well plates, and the master plates were frozen and stored at −80°C until genetic characterization identified the correctly targeted clones. Some of these targeted clones were thawed from the master plates, expanded and subjected to further procedures described below.
To activate rtTA expression from the ROSA26 locus, the transfection of the selected targeted clones with the pCAGGS-Cre-PGK-puro vector (a gift from C. Lobe) was performed using Lipofectamine 2000 (Invitrogen). An aliquot of 2 μg circular plasmid was transfected into cells growing in a 35 mm diameter dish. During the transfection, OPTI-MEM medium was used. Puromycin selection (1.25 μg/ml) was applied, and resistant colonies expanded. Green fluorescence was detected as described previously (15).
The pBI-3 plasmid (a gift from H. Bujard) contains a bi-directional tetracycline-responsive element followed by the lacZ coding sequence. This vector was introduced into ES-cells by lipofection according to the protocol above. After 5 h of transfection, doxycycline (Sigma) was added to the medium in differing concentrations. The day after lipofection, the cells were passaged 1:5 and cultured for a further 1–2 days in the presence of doxycyline (100 ng/ml), and then stained by using X-gal.
Generation of transgenic animals
Chimeric mice were generated by aggregation of targeted ES-cells with eight-cell stage embryos as described previously (16). Germline transmitting chimeric males were crossed with outbred ICR females and the offspring were genotyped by Southern blotting and PCR for the transmission of the transgene.
The Cre-recombinase transgenic founder lines were generated as described previously (17). Regarding the Podocin-Cre transgene four individual founder lines were crossed with the Z/EG reporter strain (15) to determine the degree and timing of Cre-mediated DNA excision in podocytes.
Genotyping by Southern blotting
To detect targeting of the ROSA26 locus, genomic DNA was isolated from the ES cells grown on 96-well plates (18), digested with EcoRV and the fragments separated on 0.7% agarose gel. Southern blotting was performed as described previously (19). Briefly, the DNA was transferred onto a nitrocellulose membrane (Hybond N; Amersham) and hybridized with a probe complementary to a sequence upstream of the 5′ homology arm of the vector (external probe) or with a probe complementary to the neo gene (internal probe) (Figure 1A and B). The radioactive signal was detected by phosphorimaging. To detect the mutation in mice, 10 μg genomic DNA was extracted from mouse tail biopsies and analyzed by Southern hybridization as above.
Genotyping by PCR
PCR was performed in 25 μl reaction mixture containing standard PCR buffer, 1.0 mM MgCl2, 200 μM dNTPs, 200 nM primers, 5 U Taq polymerase and 100 ng genomic DNA isolated from mouse ear biopsies. The primers for detection of the targeted ROSA26 allele containing the STOP cassette were as follows: ROSA5, GAGTTCTCTGCTGCCTCCTG; and RTTA3, AAGACCGCGAAGAGTTTGTC. The reaction resulted in a 215 bp band. The wild-type ROSA26 allele was detected by an amplicon of 322 bp using primers ROSA5 and ROSA3: CGAGGCGGATACAAGCAATA. The PCR program was as follows: initial denaturation for 1 min at 94°C followed by 35 cycles of 1 min at 94°C, 45 s at 62°C (knock-in allele) or 64°C (wild-type allele) and 1 min at 72°C and a final extension of 5 min at 72°C. PCR products were detected on 2% agarose gel.
Histology
The X-gal staining of ES cells was performed as described previously (15). Embryonic and adult tissue samples were dissected and fixed overnight in 4% PFA. The following day, the samples were washed extensively with 1× PBS, dehydrated through graded alcohol washes and embedded in paraffin wax as described previously (20). Sections (7 μm) were deparaffinized and stained with Harris' Hematoxylin and Eosin (Sigma Immunochemicals).
RESULTS AND DISCUSSION
Generation of the ROSA26-rtTA knock-in ES cell line
We inserted an artificial exon between the first and the second exons of the mouse ROSA26 locus by gene-targeting. Of the 140 G418 resistant colonies picked and genotyped, 12 (8%) were correctly targeted (Figure 1A and B). In the artificial exon, a loxP site-flanked selectable marker (neo) precedes two coding sequences separated by an internal ribosomal entry site (IRES). The upstream sequence is coding for rtTA with a nuclear localization signal, and the downstream sequence is coding for enhanced green fluorescent protein (EGFP). Translation of the latter is ensured by the IRES sequence. Before Cre-excision, the three consecutive polyadenylation signals of the selectable marker terminate mRNA synthesis and, therefore, the downstream coding sequences are not expressed (8). After Cre-mediated deletion of the loxP flanked sequence, however, rtTA and EGFP are expressed by the ROSA26 promoter. Presence of doxycycline results in the formation of an active transcriptional activator and the activation of the responder transgene (see Figure 2).
Assessment of tetracycline-inducibility in vitro
To test the inducibility of the system in vitro, we removed the loxP flanked sequence by transfecting 10 targeted ES cell lines with a plasmid expressing Cre recombinase as well as puromycin acyltransferase (pCAGGS-Cre-PGK-Puro). We isolated stable integrants using puromycin selection and verified that they displayed green fluorescence while the parental cell lines were not fluorescent (Figure 1C and data not shown). There was no difference among the different targeted cell lines. Next, we expanded the fluorescent Cre-excised subclones of two original lines (1C12 and 1H6) and transfected them with a vector containing a lacZ transgene driven by a rtTA-responsive promoter (tet-O-LacZ). (We did not use any selection for transfectants in this experiment.) We did not observe any X-gal activity in the absence of the inducer (Figure 1D). When the medium was supplemented with 100 ng/ml doxycycline, LacZ positive cells were detected (Figure 1E).
Inducible overexpression of VEGF-A in vivo
Two ES-cell clones were used to generate chimeric mice by ES cells ⇔ embryo aggregation. In both experiments, germline transmission was obtained. Heterozygous animals of both lines were healthy, fertile and had a normal lifespan. Heterozygotes from one of the lines (1C12) were intercrossed, resulting in litters of pups with a Mendelian distribution of the F2 genotypes. Homozygous transgenic animals were healthy and fertile and were used to maintain this line in the homozygous state. These homozygous, Cre-conditional ROSA26-rtTA mice were used in consecutive experiments.
In order to test the in vivo ‘silent but inducible’ nature of the conditional ROSA26-rtTA-IRES-EGFP transgene and rtTA protein activity, male mice that were homozygous for the Cre-conditional ROSA26-rtTA allele were bred with female mice that were double heterozygous for a tet-O-VEGF-A-164 responder transgene (21) and either the ubiquitous Cre-deletor line (pCAGGS-CreTg/+) (5) or a nervous system specific (Nestin-CreTg/+) (22) or a podocyte specific (Podocin-CreTg/+) Cre line. Figure 2 describes how the three transgenes work to unite the Cre/loxP and tetracycline inducible systems.
Since the pCAGGS-Cre transgene expresses Cre in all the cells of the early embryo, it activates an overall rtTA and EGFP expression as shown for a E9.5 day embryo in Figure 3B and E. On the other hand, breeding with the Nestin-Cre or Podocin-Cre line resulted in embryos/animals with neuronal lineage or glomerulus-specific expression of rtTA and EGFP, respectively, as shown in Figure 3G for a E10.5 day embryo for Nestin-Cre and in Figure 3J for a newborn glomerulus for Podocin-Cre. In the absence of Cre-mediated excision of the loxP-flanked PGK-neo-pA sequence blocks rtTA and EGFP expression.
We made use of the tight dosage sensitivity of the early developing embryo to increased (9,23) or decreased levels of VEGF-A (20,23–26) to demonstrate the correct activation of our inducible system. In the absence of the rtTA-inducer doxycycline, triple transgenic pCAGGS-CreTg/+, ROSA26-rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+ embryos (Figure 3A and B) develop normally and show no phenotype. However, in the presence of doxycycline that was administered to pregnant females starting one day after mating (E1.5), all of these triple transgenic embryos showed lethality (Figure 3D and E and see Table 1) at E9.5. The phenotypes of these mutant embryos were quite severe with no primitive red blood cells (RBCs) observed in the developing yolk sac (Figure 3D) and embryos were blocked in development. Unlike the original homozygous VEGF-A mutants that showed a severe reduction of blood island development in the yolk sac (24,25), ubiquitous VEGF-A-164 expression in this triple transgenic inducible system led to enlarged blood islands in the yolk sac that were filled with nucleated erythroblast progenitors (Figure 3F). The normal blood islands that developed in the non-induced embryos showed clearly defined endodermal and mesodermal layers ‘sandwiching’ the developing blood islands (Figure 3C); however, in the doxycycline-induced embryos, aberrant blood islands formed (Figure 3F). All the 10 triple transgenic embryos showed the same mutant phenotype described above upon doxycycline administration to their mothers (Table 1). Out of the 22 remaining littermates that were either single or double transgenic for the three transgenes, 20 appeared totally normal. Two rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+, Cre-negative embryos (as judged by Southern hybridization and PCR, data not shown), however, were also EGFP expressers indicating that rtTA expression activation (Cre-excision) occurred by maternally produced Cre protein. This known phenomenon is a consequence of high-Cre production and accumulation during oogenesis (27).
The ubiquitous systemic administration or induction of the VEGF-A-164 protein, or injection of tumor cells expressing high levels of VEGF-A-164 protein into adult mice, results in severe edema (28–30), defects in immune cell function including destruction of the thymus (31–33) and liver pathologies that closely resemble the human syndrome known as peliosis hepatis that has been described in association with Bartonella henselae infection, long-term high-dose androgen therapy, or rarely with advanced cancers (34). In order to determine if doxycycline-induced overall VEGF-A-164 expression could mimic some or all of these previously described phenotypes, triple transgenic pCAGGS-CreTg/+, ROSA26-rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+ mice were allowed to reach adulthood and a regimen of doxycycline administration was followed (Table 1). These triple transgenic mice were born in a normal Mendelian frequency and showed no sign of any phenotypic abnormalities (Figure 4A–C and G–I, and data not shown). Upon administration of doxycycline in the drinking water to 3–4 month old pCAGGS-CreTg/+, ROSA26-rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+ mice, several phenotypes became apparent already after 2 days of treatment. The mice showed signs of edema and erythema of the face, ears and feet (data not shown). By 5 days of doxycycline administration, three of the eight triple transgenic mice had died and three were so moribund and sickly in appearance that they had to be euthanized and their organs were harvested for further histological analysis. Two mice seemed more resistant to the doxycycline administration and were given the drug for an additional 4 days at which time they too became moribund and were as well sacrificed. Of the 10 single and double transgenic littermates, 9 were completely normal and showed no adverse side affects to the doxycycline administration. Similar to what was observed in the E9.5 embryos, one rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+, Cre-negative adult mouse died during doxycycline administration. This mouse, however, was EGFP positive, indicating the rtTA expression activation by zygotic (maternal) Cre protein (27). Upon autopsy, the most obvious gross phenotypes observed were an extended blood filled liver and a dramatic decrease in the size of the thymus together with enlarged lymph nodes and signs of hyper-vascularity/permeability and edema in several other organ systems as well as blood in the intestine (data not shown).
Histological analysis of the liver of uninduced pCAGGS-CreTg/+, ROSA26-rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+ adult mice (control) revealed normal hepatic architecture (Figure 4A–C). Doxycycline-treated mutant pCAGGS-CreTg/+, ROSA26-rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+ adult livers showed a dramatic ‘peliosis-like’ phenotype (Figure 4D) and signs of blood pooling in large areas of the liver (arrows in Figure 4E). This liver peliosis phenotype is characterized by enlarged hepatic sinusoids and a total disruption of the normal liver architecture (Figure 4D–F) as well as detached sinusoidal endothelial cells that are ‘sloughing-off’ into the sinusoidal space (arrow in Figure 4F). The liver pathology described here resembles the liver peliosis-like pathology recently described in mice that had been engrafted with tumor cells expressing high levels of systemic VEGF-A-164 protein (34). In addition, LeCouter et al. (35) have observed similar (although less severe) liver enlargement phenotypes and effects on the sinusoidal endothelium of the liver with systemic administration of recombinant VEGF-A protein. It was also demonstrated that VEGF-A at lower doses can act on the sinusoidal vasculature (via Flt1) to influence secretion of hepatocyte growth factor (HGF) and may have a beneficial protective effect on hepatocytes under times of cytotoxic stress (35). Our VEGF-A inducible system with liver-specific Cre lines such as the albumin-Cre line (36) will be useful in determining the optimal dosages of VEGF-A required for its beneficial action on the liver under times of cytotoxic stress and may at higher induction levels be a useful model system to unravel the molecular mechanisms behind VEGF-A's harmful effects on the liver.
The thymus of non-induced pCAGGS-CreTg/+, ROSA26-rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+ adult mice (control) displayed a normal thymic architecture with a well-defined cortical layer rich in basophilic thymocytes and a normal epitheloid-rich medullary zone (Figure 4G–I). Total average cell counts for two control thymi were 1.33 × 108 cells/ml (1.05 × 108 and 1.6 × 108 cells/ml in the two different controls). Representative histological analysis of the thymi of VEGF-A-164-induced mutant adults (Table 1) revealed a dramatic decrease in cellularity that correlated with its decreased size (Figure 4J–L). The mutant thymi lacked major histological distinctions between the medullary and cortical layers and the basophilic cortical layer that is usually rich in thymocytes was almost completely absent (Figure 4L). Total average cell counts for three mutant thymi were 1.37 × 107 cells/ml (3.9 × 107, 8.1 × 105 and 1.14 × 106 cells/ml per animal). These numbers may under-represent the acellularity of the mutant thymi as two of the higher values were from the two animals that survived the initial five-day period of doxycycline treatment. The lower cell count of 8.1 × 105 cells/ml from one of the original sick animals that had to be euthanized in the first five-day period may be more representative of the acellularity shown in Figure 4J–L. The thymic atrophy presented in this paper phenocopies previous reports concerning the detrimental effects that systemic VEGF-A has on T-cell development (33). In addition, ubiquitous expression of VEGF-A-164 during early development seems to give rise to increased numbers of erythroblasts and may inhibit the differentiation of these progenitors. From this study, it is not clear whether this phenotype is caused by alterations in the yolk sac environment or by artificial VEGF-A-164 autocrine loop created. We are currently investigating the molecular mechanisms behind VEGF-A's suppressive effects on different hematopoietic populations through the use of this transgenic system together with additional hematopoietic lineage-specific Cre lines (37,38).
The developing nervous system is also sensitive to altered VEGF-A expression, We have recently demonstrated that downregulation of the level of this growth factor, using a Nestin-Cre transgene and a conditional targeted allele of VEGF-A, have severe consequences to cortical vessel density and structure (39). The Nestin-CreTg/+, ROSA26-rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+ triple transgenic embryos presented here developed vascular abnormalities by E12.5, associated with spinal cord and brain hemorrhages, when doxycycline was administered to their mother from 1.5 dpc of pregnancy (Figure 3H and I). Without doxycycline administration to the mother, these embryos developed normally and reached adulthood. Interestingly, when these mice received doxycycline for 5 days at 4 months of age, they did not show any obvious gross behavioral or phenotypic abnormalities. Initial vessel and histological analysis from two of the six treated adult mouse brains did not reveal any striking differences in vessel architecture or structural organization of the cortex (data not shown). This result may indicate an increased resistance of the adult CNS vasculature to VEGF-A. We are currently further examining the potential effects of doxycycline-induced increases of VEGF-A-164 and its effects on the developing and adult nervous system using this triple transgenic system.
In contrast, the Podocin-CreTg/+, ROSA26-rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+ triple transgenic mice treated with doxycycline for 7 days starting at 4 months of age developed proteinuria (albumin >5 g/l, data not shown). The presence of albuminuria in concentrations >3 g/l is designated ‘nephrotic range proteinuria’. This degree of proteinuria represents loss of the permeselective function of the glomerular filtration barrier. Increased permeability in the glomerulus leads to the loss of critical blood proteins, including albumin, blood clotting factor inhibitors and lipids, and is associated with edema and thrombotic events in patients. These mice are currently under detailed phenotypic analysis to identify the nature of changes in the kidney caused by induction VEGF-A expression in podocytes.
Table 1 displays the results of all the experiments described above. In summary, by combining existing tissue-specific Cre-transgenic mouse strains to define spatial, and the tetracycline-inducible system to define temporal aspect of transgene expression, we have developed a versatile system which allows quick and efficient investigation of multiple gene functions. This aim has also been addressed by others using hormone-activated Cre proteins (40,41). The major advantage of these earlier efforts lie in that only two transgenes have to be combined. However, our system allows for complete on/off control of expression. In addition, the existing inducible systems cannot be combined with and take advantage of the large number of existing Cre-transgenic mouse strains defining tissue/cell type-specificities. Our results show that when expressed from the ROSA26 locus, the baseline activity of rtTA in the absence of inducer is not sufficient to cause significant activation of a responsive promoter. This is in agreement with the findings of Wutz and Jaenisch (42), who used constitutive rtTA expression from the ROSA26 promoter. We expect though that the ROSA26 locus placed rtTA could be a limitation by providing only a given level of expression for rtTA, which might not be high enough to reach sufficient level of induction of different responder tet-O-genes in certain anatomical areas or cell types. Therefore, an obvious future improvement of the presented system would be the establishment of Cre conditional rtTA transgenes in several genomic positions permissive to different level of overall expression. Our ROSA26 placed Cre-conditional system could be the first member of this envisioned useful series.
Acknowledgements
We are grateful to Dr C. G. Lobe for providing the pCALL2-IRES-EGFP and to Dr H. Bujard for the pBI-3 vectors. These studies were supported in part by a National Cancer Institute of Canada grant (NCIC grant 21335). J.H. was a recipient of an NCIC postdoctoral fellowship. A.N. is a senior Canadian Institutes of Health Research (CIHR) scientist. S.E.Q. is supported by CIHR grant no. 62931. Funding to pay the Open Access publication charges for this article was provided by NCIC 021335.
Conflict of interest statement. None declared.
Figures and Tables
Figure 1
Targeted insertion of a conditional rtTA-IRES-EGFP transgene into the ROSA26 locus. (A) Targeting strategy. The exons of the ROSA26 gene are depicted as numbered boxes, and the triangles as loxP sites. The insertion point (XbaI site), informative restriction sites and diagnostic fragments are also shown. Abbreviations are explained in the text. Drawing is not to scale. (B) Southern-blot analysis of genomic DNA from ES cells digested by EcoRV and hybridized with a 5′ external probe (left) and neo probe (right) (C) After Cre-excision, cells show green fluroescence. Line 1C12 was transfected with a plasmid carrying Cre and puro expression cassettes. A puromycin-resistant colony is shown. (D and E) Tetracyclin-induction of lacZ in Cre-excised 1C12 cells. The cells were transfected with a plasmid containing a tet-O-lacZ transgene. (D) No doxycycline induction. (E) Cells after doxycycline administration (100 ng/ml for 2 days).
Figure 2
Operation of the Cre/loxP-dependent, tetracycline inducible transgenic system. (A) Mice that are homozygous for the targeted insertion of the conditional rtTA-IRES-EGFP bicistronic transgene at ROSA26 can be bred with double transgenic mice that carry a tissue-specific Cre transgene and a doxycycline-inducible rtTA-dependent tet-O-GENE responder line. A total of 25% of the pups will be triple transgenic (SpecPromoterCreTg/+, ROSA26-STOP-rtTA-IRES-EGFPTg/+, tet-O-GENETg/+). (B) In cells that do not express Cre, neither rtTA nor EGFP protein is generated; therefore, the tet-O-GENE is silent. (C) In Cre-expressing cells, rtTA and EGFP expression is turned on. However, in the absence of an inducer (e.g. doxycycline), rtTA cannot activate the expression of the tet-O-GENE target. Addition of doxycycline results in the formation of an active transactivator and expression of the target gene. In the present study, VEGF-A-164 was induced either ubiquitously or nervous system or podocyte specifically in the presence of doxycycline.
Figure 3
Tissue-specific expression of the rtTA-IRES-EGFP bicistronic transgene is Cre-dependent and the rtTA trans-activation of tet-O-VEGF-A-164 is tightly regulated by doxycycline. (A–C) Images of triple transgenic pCAGGS-CreTg/+, ROSA26-rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+ E9.5 embryo and yolk sac in the absence of doxycycline: (A), Whole mount bright field microscopy; (B), GFP fluorescence microscopy; and (C), H&E histological section of the yolk sac. In the absence of doxycycline, triple transgenic embryos develop normally. Arrow in (A) shows normal yolk sac vessel with primitive hemoglobin-containing RBCs and normal blood island (BI) formation between the primitive endoderm (e) and mesoderm (m) layers of the yolk sac (C). (D–F) Triple transgenic embryo and yolk sac, the mother treated with doxycycline. (D), Whole mount bright field microscopy; (E), GFP fluorescence microscopy; and (F), H&E histological section of the yolk sac. (D) In the presence of doxycycline the triple transgenic embryos show a lethal phenotype with no proper vessel development in the yolk sac (ys) and embryo (e), no hemoglobin-containing RBC (D) and abnormal blood island (BI) structures that are filled with excessive nucleated erythroblasts (EB). (B and E) The overall EGFP signal marks the overall rtTA expression in the yolk sac and embryo of the triple transgenic embryos. (G) E10.5 triple transgenic Nestin-CreTg/+, ROSA26-rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+ embryos develop normally in the absence of doxycycline and show nervous system specific expression of EGFP/rtTA. (H and I) E12.5 triple transgenic embryos when the mother was either non-treated (H) or treated (I) with doxycycline during the pregnancy. The VEGF-A-164 induced embryo (I) shows hemorrhages in the developing nervous system (arrows). (J) Immunostaining for EGFP shows the podocyte specificity of Podocin-Cre excision.
Figure 4
Liver peliosis-like phenotype and thymus degeneration are associated with the doxycycline-induced overall VEGF-A-164 expression in adult pCAGGS-CreTg/+, ROSA26-rtTA-IRES-EGFPTg/+, tet-O-VEGF-A-164Tg/+ mice. H&E histological analysis of livers of triple transgenic adult mice of (A–C): VEGF-A-164 non-induced (−DOX) and (D–F): VEGF-A-164 induced (+DOX) by doxycycline treatment. Without doxycyclin treatment, livers have a normal hepatic architecture with well-defined vasculature and normal hepatic venules (V) and hepatic sinusoids (HS) (C). The doxycycline-treated mutant livers show major disruptions of normal hepatic architecture with a severe dilation of hepatic sinusoids (HS) (F) and evidence of blood engorgement and pooling [arrows in (E)]. Arrow in (F) shows evidence of sinusoidal endothelial sloughing and detachment. (G–I): H&E histological analysis of VEGF-A-164 non-induced (−DOX) and (J–L): VEGF-A-164 induced (+DOX) thymus of triple transgenic adult mice. Without doxycycline treatment the thymus shows normal thymic architecture with a well-defined cortex layer (cx) that is packed with basophilic lymphocytes and a normal medullary layer (m) that contains fewer lymphocytes but an extensive epithelial framework. Doxycycline-treated mutant thymus shows massive degeneration with all three thymic lobes fitting into a single optical field (J) compared with one thymic lobe fitting into the optic field at the same magnification (G) from control mice. In addition, the mutant thymus lacks a well-defined cortical layer owing to a dramatically decreased number of basophilic thymocytes seen at higher magnifications (L) compared with (I). (A, D, G and J), 50× magnification; (B and E), 125× magnification; (H and K), 250× magnification; and (C, F, I and L), 500× magnification.
Table 1
Summary of doxycycline induction experiments during development and in adult mice | [
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15850489 | GABAA receptor γ2 subunit knockdown mice have enhanced anxiety-like behavior but unaltered hypnotic response to benzodiazepines
Abstract
Background
Gamma-aminobutyric acid type A receptors (GABAA-Rs) are the major inhibitory receptors in the mammalian brain and are modulated by a number of sedative/hypnotic drugs including benzodiazepines and anesthetics. The significance of specific GABAA-Rs subunits with respect to behavior and in vivo drug responses is incompletely understood. The γ2 subunit is highly expressed throughout the brain. Global γ2 knockout mice are insensitive to the hypnotic effects of diazepam and die perinatally. Heterozygous γ2 global knockout mice are viable and have increased anxiety-like behaviors. To further investigate the role of the γ2 subunit in behavior and whole animal drug action, we used gene targeting to create a novel mouse line with attenuated γ2 expression, i.e., γ2 knockdown mice.
Results
Knockdown mice were created by inserting a neomycin resistance cassette into intron 8 of the γ2 gene. Knockdown mice, on average, showed a 65% reduction of γ2 subunit mRNA compared to controls; however γ2 gene expression was highly variable in these mice, ranging from 10–95% of normal. Immunohistochemical studies demonstrated that γ2 protein levels were also variably reduced. Pharmacological studies using autoradiography on frozen brain sections demonstrated that binding of the benzodiazepine site ligand Ro15-4513 was decreased in mutant mice compared to controls. Behaviorally, knockdown mice displayed enhanced anxiety-like behaviors on the elevated plus maze and forced novelty exploration tests. Surprisingly, mutant mice had an unaltered response to hypnotic doses of the benzodiazepine site ligands diazepam, midazolam and zolpidem as well as ethanol and pentobarbital. Lastly, we demonstrated that the γ2 knockdown mouse line can be used to create γ2 global knockout mice by crossing to a general deleter cre-expressing mouse line.
Conclusion
We conclude that: 1) insertion of a neomycin resistance gene into intron 8 of the γ2 gene variably reduced the amount of γ2, and that 2) attenuated expression of γ2 increased anxiety-like behaviors but did not lead to differences in the hypnotic response to benzodiazepine site ligands. This suggests that reduced synaptic inhibition can lead to a phenotype of increased anxiety-like behavior. In contrast, normal drug effects can be maintained despite a dramatic reduction in GABAA-R targets.
Background
GABAA-Rs are the major inhibitory receptors in the mammalian central nervous system and can be modulated by a number of sedative, hypnotic and anesthetic drugs [1]. GABAA-Rs are putatively pentameric complexes and are members of a 'superfamily' of related ligand-gated ion channels. There are a variety of subunit families that make up GABAA-Rs; a total of seventeen distinct subunits have been cloned, α1–6, β1–4, γ1–3, δ, ε, π, and θ. Much is known of the subunits that comprise GABAA-Rs, but the contribution of individual subunits to in vivo drug responses is only beginning to be understood, largely through the use of genetically engineered mice [2-4].
The γ2 subunit is highly expressed throughout developing and adult brain and spinal cord, is a component of almost 60% of all GABAA-Rs [5,6], and is required for synaptic clustering of GABAA-Rs [7]. In vitro electrophysiologic studies have demonstrated an obligatory role of the γ2 subunit for benzodiazepine responsiveness [8]. Gunther et al. created and studied mice that globally lacked all γ2 subunits of the GABAA-R [9]. Mice homozygous for the γ2 knockout died in the perinatal period with rare survivors reaching postnatal day 18. Knockouts appeared normal at birth, but developed sensorimotor abnormalities characterized by hyperactivity, impaired grasping and righting reflex, and abnormal gait. Pharmacologically, knockout of γ2 resulted in a severe deficit in benzodiazepine binding sites (~94% reduction compared to control) while GABA binding sites were only slightly reduced. Behaviorally, in the few mice that survived long enough to be tested, diazepam failed to induce sedation and loss of the righting reflex at doses that were fully effective in wild type mice [9]. γ2 heterozygous knockout mice were normally viable, had a ~50% reduction in γ2 protein levels, and displayed increased anxiety-like behaviors [10]. Drug responses of heterozygous γ2 global knockouts have not been reported, except for the high efficacy of low diazepam doses to induce anxiolysis [10].
We set out to make mice with markedly attenuated expression of the γ2 gene (i.e., γ2 knockdown mice) that would be viable and therefore useful for studies of basic behaviors and drug induced responses. We used gene targeting and embryonic stem cell technologies to create this mouse line by inserting a neomycin resistance gene (NEO) into intron 8 of the γ2 genomic locus. Insertion of NEO into intronic DNA has previously been used to create hypomorphic alleles of several genes [11-23]. In those studies, expression of the hypomorphic allele ranged from 5–25% of control; substantially lower than the 50% reduction typically observed in heterozygous knockouts. The γ2 knockdown mouse line described in the present report had a highly variable reduction in γ2 mRNA and protein levels. γ2 knockdown mice were normally viable, had increased anxiety-like behaviors, but did not differ in the hypnotic response to benzodiazepines. In addition, since exon 8 of the γ2 gene was flanked by loxP sites in this mouse line, these mice could also be converted into a global knockout mouse by crossing to a general deleter Cre-expressing transgenic mouse line.
Results
Production of gene targeted mice
Four out of 330 embryonic stem cell clones displayed the predicted restriction fragment length polymorphisms (data not shown) indicative of the targeting event presented in Fig 1A. Because the targeting event results in a γ2 locus in which exon 8 is flanked by loxP sites, we refer to this locus as being floxed (F) in contrast to the endogenous wild type allele (+). Two correctly targeted cell lines, 26L6 and 26C8 yielded germ-line competent chimeric males. Chimeras were mated to C57BL/6J females. Offspring that were +/F were interbred to produce +/+, +/F, and F/F mice. Mice were genotyped at weaning using Southern blot analysis as indicated in Fig 1B. In this analysis, Probe A hybridized to a 9.2 kB BglII fragment from the wild type γ2 allele and a 3.4 kB fragment from the targeted allele. Of 724 mice genotyped, 196 (27%) were +/+, 372 (51%) were +/F, and 156 (22%) were F/F. These values are in accord with the 1:2:1 genotype frequency as expected by Mendelian genetics. Thus, homozygous floxed mice were normally viable. They were also healthy and overtly indistinguishable from wild type mice.
Northern blot analysis
Semi-quantitative northern blots were used to compare the amount of γ2 mRNA in wild type and F/F animals. Hybridization of adult brain RNA with a γ2 cDNA probe yielded an intense 2.8 kB band in wild type mice. In contrast, RNA from F/F mice hybridized less intensely in most mice, however the attenuation within this group was highly variable. A sample northern blot is shown in Figure 2A. We semi-quantitatively determined γ2 mRNA amounts by comparing γ2 band densities between wild type (n = 23) and F/F (n = 36) mice following normalization with β-actin. To normalize results over four different blots, the ratio of wild type band densities on each blot was averaged and normalized to 100. All samples on each blot were then compared to the average of wild type band densities. Values are shown in Figure 2B. Mean wild type ratio was 100 whereas mean ratio in F/F samples was 35 with high variability including several mice with values that overlapped with wild type values. Thus, γ2 mRNA levels in knockdown mice were dramatically attenuated on average, but the levels varied significantly between mice.
Immunohistochemistry
Immunohistochemical staining of sagittal sections probed with a γ2-specific antibody was used to assess γ2 protein amount and distribution in the brain. A sampling of wild type (Fig 3A) and knockdown (Fig 3B) brain sections are displayed. In three of the four knockdown mouse sections shown, there was a marked overall reduction of γ2 immunoreactivity in all areas of the brain except the outer layer of the olfactory bulb. However, staining intensity was variable between knockdown mice, with one having near wild type intensity (Fig. 3B, d). A total of 10 F/F mice were studied. Seven out of 10 had a marked decrease in γ2 staining compared to wild type controls, and three out of 10 had near wild-type or intermediate levels of staining. Adjacent sections from these same mice showed no change in α1 or β2/3 immunoreactivity (CPM and ALD, unpublished observations).
Pharmacological characterization
Autoradiography on brain cryostat sections with various ligands binding to selective sites of the GABAA-R revealed clear differences in the homozygous γ2 knockdowns as compared to heterozygous and wild type mice (Table 1). Total flumazenil-sensitive [3H]Ro15-4513 binding to benzodiazepine sites was reduced in the knockdown in most brain regions. The reduced binding in the 5 mice analyzed ranged on the average from 25 to 57%. In several regions, the heterozygous mice also showed significant reduction of the binding, but clearly less than the homozygous knockdown samples. A small proportion of the reduction in the total binding was apparently due to reduction in diazepam-insensitive α4 and α6 subunit-containing receptors, this component being mostly affected in the cerebellar granule cell layer.
GABA-sensitive [3H]muscimol binding to GABA agonist sites was not altered in any brain region (Table 1).
Picrotoxinin-sensitive [35S]TBPS binding to the GABAA-R channels was little affected between the mouse lines: basal binding was increased in the hippocampus and decreased in the cerebellar granule cell layer of the homozygous γ2 knockdowns (Table 1). The GABA-insensitive [35S]TBPS binding was increased all over the brain, but this component still made up less than 10% of the basal binding, except for the cerebellar granule cell layer. There, the GABA-insensitive component was 20% as compared to 10% in the heterozygous and wild type animals.
Behavioral characterization
We examined the behavioral phenotype of the knockdown mice using the elevated plus maze and forced exploration in a novel open field. The elevated plus maze is widely used to assess anxiety-like behavior in response to the aversive stimulus of an elevated exposed space. On this test apparatus, knockdown mice had a 74% reduction in open arm entries (Fig. 4A) and an 83% reduction in time on open arms (Fig. 4B) compared to controls suggesting an increase in anxiety-like behavior. In contrast, knockdown mice had the same number of total arm entries as wild types (Fig. 4C) indicating that an alteration in locomotion does not account for the differences in anxiety-like behavioral measures on this assay.
Knockdown mice were also tested for behavioral response to an aversive, brightly lit open field test apparatus using the forced novelty exploration test. Knockdown mice had a 28% reduction in locomotor activity compared to wild type mice when placed into this test arena (Fig. 4D). This reduction in exploratory behavior in the knockdown animals is also indicative of an increase in anxiety-like behavior.
Knockdown of the γ2 subunit did not alter hypnotic responses to benzodiazepine site ligands or other sedative/hypnotic drugs tested. We measured the loss of righting reflex to assess the acute sensitivity to sedative/hypnotic drugs; no significant differences were found in response to injection of diazepam, midazolam, zolpidem, ethanol, or pentobarbital (Fig. 5).
Production of γ2 global knockout mice
The gene targeting strategy that we used to produce the γ2 knockdown mice also resulted in the insertion of loxP sites that flank exon 8 of the γ2 gene (see Fig. 1A). To determine if these loxP sites were functional in vivo, heterozygous floxed γ2 mice were mated to a general deleter cre-expressing transgenic mouse line (JAX Laboratory, stock #003376) where the cre transgene was driven by a ubiquitously expressed human beta actin promoter [24]. Restriction fragment polymorphism analysis by Southern blotting confirmed the deletion of genomic DNA between loxP sites. As predicted from Fig. 1A, Probe A hybridized to a 6.6 kb BglII fragment from the recombined knockout allele (data not shown).
To determine if cre-mediated recombination and deletion of exon 8 resulted in a null allele, mice hemizygous for the cre transgene and heterozygous for the recombined γ2 allele (+/f) were mated to mice heterozygous for the floxed, non-recombined γ2 allele (+/F). 10.2% (14 of 137) of offspring from this mating strategy were homozygous for the recombined locus (f/f). This ratio is in agreement with the 12.5% predicted from Mendalian genetics. 13 of the 14 f/f mice died within three days of birth. One f/f mouse survived until postnatal day 21. This neonatal lethality of γ2 f/f mice is consistent with those found in the global γ2 knockouts of Günther et al. [9] which also had a deletion of exon 8. Therefore, we conclude that cre-mediated recombination of the loxP sites and deletion of exon 8 inactivates the γ2 gene.
Discussion
Genetic dissection of the GABAA-R system is yielding remarkable insights into GABAA-R biology and the mechanisms of action of various drugs. Here we report the establishment of a new GABAA-R mutant mouse model, one with attenuated expression of the γ2 subunit that averages ~35% of normal levels. This novel γ2 knockdown model has several unique features that make it useful. First, compared to global knockouts which die as neonates [9], knockdown mice have normal viability. Secondly, the graded levels of expression of γ2 (range 10–95% of normal) in the mutant mice could be useful in studies correlating phenotype with the amount of γ2 protein present. Finally, global knockout mice can be created from these mice by crossing to a Cre recombinase expressing global deleter mouse line. Therefore, the mice reported here represent a useful addition to the rapidly expanding arsenal of mice with genetic alterations in the γ2 subunit of the GABAA-R. In addition to homozygous and heterozygous global γ2 knockouts [9], conditional γ2 knockout mice [25], knockin mice with a point mutation in γ2 that eliminates zolpidem and inverse agonist β-carboline sensitivity [26,27], global knockout of the long splice variant of γ2 [28], and transgenic mice that express either the long or short splice variants of the γ2 subunit [29] have been described.
An important unexpected finding from this study was the extremely variable degree to which the genetic alteration of the γ2 locus changed the amount of γ2 product produced. This γ2 knockdown model was produced by insertion of NEO into intronic DNA. This strategy has been used previously to create hypomorphic alleles [11-23]. Creation of hypomorphic alleles has been useful in determining function of the gene that has been targeted, especially in cases where complete disruption of the gene is lethal [12,14-16]. Intronic insertion of NEO creates hypomorphic alleles either through cryptic splicing signals in the NEO gene that lead to a frameshift or alterations in splicing, or by down regulating gene expression through an unknown mechanism [14-16]. In the majority of these studies, the authors' reported a 5–25% level of expression from the targeted gene. However, in none of these previously described hypomorphic alleles was extensive variability reported. The γ2 targeted homozygous mice in this study clearly display a wide range of γ2 mRNA and protein levels. Semi-quantitative northern blot analysis revealed that 4 out of 5 homozygous knockdown mice showed, on average, a 70% reduction of γ2 mRNA levels, whereas 1 out of 5 mice had γ2 mRNA at a level that was similar to wild type. Immunohistochemical data were consistent with northern blot data. The reason for this highly variable expression within homozygous knockdown mice has yet to be determined. The mixed background of Strain 129S1/X1 and C57BL6/J may not be the cause given that other studies have not observed substantial variability in expression using similar background strains [11,18,20]. Gender as a cause of variability also is not likely given that a set of three male knockdown siblings exhibited the full range of γ2 levels; one being near wild type levels, another being at an intermediate level, and the third exhibiting severely attenuated expression. Further study to determine the cause of this variability needs to be undertaken. Nevertheless, investigators who desire to create a hypomorphic allele by inserting NEO into intronic DNA should take into consideration the variability of gene expression observed in this study. The extent of variability may be locus dependent.
Knockdown of γ2 subunit containing GABAA-Rs resulted in behavioral changes that are indicative of increased anxiety-like behavior. These mutant knockdown mice showed reduced exploratory behavior for a novel open field apparatus and the open arms of the elevated plus maze. Similar changes have been observed in γ2 global heterozygous knockout mice [10]. Together, these results strengthen the possibility that GABAA-R dysfunction may be an underlying cause of anxiety disorders in humans. These mutant mouse models may be useful for dissecting the etiology of this pervasive condition and for developing effective therapeutic interventions.
As expected, brains from γ2 knockdown mice demonstrated significantly decreased binding of the benzodiazepine site ligand, Ro15-4513. This was expected since benzodiazepine binding sites are located at the interface of select alpha and γ2 subunits of the GABAA-R [30]. The reduction observed in Ro15-4513 binding agree with those from global γ2 knockout mice [9] in that deficiency of the γ2 subunit abolishes the binding of the benzodiazepine-site ligands. The global heterozygous γ2 knockout mice have a reduction in total [3H]Ro15-4513 binding in most brain regions [31], and the present data on the γ2 knockdown show a very similar pattern of reduced binding. However, the 5 knockdown brains that we examined had actually a greater reduction in benzodiazepine binding than the γ2 heterozygous global knockouts published earlier [31]. Similar to the γ2 heterozygous knockouts, [3H]muscimol binding was not affected, indicating that the reduction of the γ2 subunit levels is not compensated by increased δ subunit that is largely responsible for the [3H]muscimol binding signal in brain sections [32]. The basal binding of the ion channel ligand [35S]TBPS is usually reduced and its sensitivity to GABA increased by addition of γ2 subunits [31]. Both in the γ2 heterozygous global knockout and in our homozygous knockdown, TBPS binding was increased in some brain regions, and, more consistently, there emerged a "GABA-insensitive" receptor population with widespread distribution in the brain. This indicates production of GABAA-Rs with an αβ configuration. These receptors bind strongly the agonist and have reduced channel conductance [33], perhaps reflecting only partial agonist efficacy of GABA. This would explain the reduced allosteric efficacy of GABA in abolishing TBPS binding. In addition, these receptors have most likely non-synaptic, non-clustered localization as they lack the γ2 subunit [10,33]. Therefore, the partially γ2 depleted mice may have brain region-selective alterations in synaptic and extrasynaptic receptors, and the reduced synaptic GABAA-R function might correlate with the anxious phenotype. However, surprisingly, the high-dose hypnotic effects of the benzodiazepine site ligands diazepam, zolpidem and midazolam were unchanged in the knockdown mice.
Several hypotheses can be generated to explain the discrepancy between decreased benzodiazepine ligand binding and unchanged hypnotic effects. One possibility is that only a threshold level of γ2 containing GABAA-Rs are required for hypnotic responses to benzodiazepines. Mice that completely lack γ2 had only 6% of benzodiazepine binding compared to wild type mice and were insensitive to the hypnotic effects of diazepam [9]. Likewise, mice with a point mutation in γ2 that eliminated response to zolpidem eliminated the effects of this benzodiazepine site ligand [26]. It seems likely that the 20–40% of benzodiazepine sensitive receptors that remain in the knockdown mice are enough to mediate the hypnotic effect of benzodiazepine site ligands. This is supported by positron emission tomography studies in humans, in which only a small proportion of the benzodiazepine sites need to be occupied by lorazepam and zolpidem to induce clinical effects such as sedation [34,35]. There may thus be spare receptors that can be measured by biochemical techniques but are not needed for whole animal pharmacological effects. Another hypothesis is that knockdown of γ2 does not result in a proportional reduction in all γ2 containing GABAA-Rs. Recent studies conducted in knockin mice with a point mutation at the benzodiazepine binding site have attributed α1 and α2 containing GABAA-Rs to the mediation of benzodiazepine-induced sedation and anxiolysis, respectively [4,36-38]. Perhaps γ2 knockdown mice have disproportionate decreases in non-α1 containing receptors, e.g., α2 or α3 containing receptor isoforms. It may be possible that in our γ2 knockdown mice, the number of α2βxγ2 GABAA-Rs is greatly reduced while the number of α1βxγ2 receptors is not, which remains to be studied. This could be tested by immunohistochemical methods using antibodies against α2 and α3 or by testing for changes in zolpidem insensitive binding. Changes in subunit trafficking may also play a role. Essrich et al. [7] established strong evidence that GABAA-Rs are clustered at the synapse through indirect interactions between γ2 and gephyrin. Enough clustering may remain in the knockdown mice such that drug response does not change. Alternatively, synaptic clustering of GABAA-Rs may not be important for behavioral responses to benzodiazepines, and it is possible that benzodiazepines potentiate the extrasynaptic GABAA-R responses determined by electrophysiology in brain slices [39]. It is also possible that the high variability in γ2 levels in the brain of knockdown mice masked any change in behavioral response to these drugs. In hindsight, it would have been useful to quantify γ2 levels in the same mice that were used for the behavioral sensitivity studies. It would be interesting to see if sensitivity correlated with γ2 levels on an individual animal basis. This type of approach could be exploited in future studies of γ2 receptor function. Lastly, decreases in benzodiazepine binding may not directly translate into changes in in vivo insensitivity. For example, in α1 null mice, changes in benzodiazepine binding did not change whole animal sensitivity to midazolam but increased sensitivity to diazepam [40,41]. In contrast, in mice that selectively lack the long splice variant of γ2, affinity for benzodiazepine site ligands is increased and behavioral response to the sedative effects of midazolam and zolpidem is also increased [42]. Similarly, in mice lacking the β3 subunit, decreased benzodiazepine binding and decreased whole animal midazolam sensitivity were observed [43]. Further study is required to examine these possibilities.
Conclusion
We have produced a novel genetically engineered mouse line that exhibits a reduction in γ2 mRNA levels that is highly variable between mice (range 10–95% of control) but averages ~35% of control levels. These γ2 knockdown mice are viable and have enhanced anxiety-like behavioral abnormalities. Surprising, these mice are normally sensitive to the hypnotic effects of benzodiazepine site ligands. Lastly, this novel genetically engineered mouse line can also be easily converted to a γ2 global knockout mouse line by simply crossing to a cre-expressing global deleter transgenic mouse line.
Methods
Generation of genetically altered mice
A 13.6 kB BglII Strain 129/SvJ mouse genomic DNA fragment containing Exons 8–10 of the GABAA-R γ2 gene was subcloned from a P1 phage clone (Genome Systems, St. Louis, MO). An oligonucleotide containing a loxP site and an EcoRV site was inserted into a NcoI site 647 bp 5' to Exon 8. A blunted fragment containing a NEO resistance gene with PGK-1 promoter and polyadenylation signals (obtained from pPNT [44]) fused with a loxP site was inserted into a HincII site 855 bp 3' of Exon 8. The NEO gene was inserted in the opposite orientation to γ2 transcription. A PGK driven thymidine kinase expression cassette (obtained from pPNT [44]) was then cloned 3' to the γ2 genomic DNA. The targeting construct was linearized with SalI and electroporated into R1 embryonic stem cells [45] following previously described procedures [46]. G418 (270 μg/ml; Life Technologies, Gaithersburg, MD) and gancyclovir (2 μM; Sigma) resistant cells were screened for gene targeting by Southern blot analysis of BglII digested genomic DNA. Fragments were hybridized with a 5' probe that was external to the targeting construct (see Fig. 1). Proper targeting of the γ2 locus was confirmed by Southern blot analysis of EcoRV and SstI digested DNA. Additional Southern blot analysis with probes to the neomycin cassette or internal to the 3' arm of the targeting vector (with 1–3 restriction enzymes) were also conducted (results not shown). All results were consistent with a correctly targeted γ2 locus.
Two correctly targeted embryonic stem cell clones (26C8 and 26L6) were microinjected into C57BL/6J blastocysts to produce chimeric mice. Male chimeras were mated to C57BL/6J females. Agouti offspring heterozygous for the targeted allele (F/+) were interbred to produce mice that were wild type (+/+), heterozygous, or homozygous mutants (F/F). The mice used in studies reported here were derived from the 26L6 ES cell clone. All mice were housed under conditions of lights on at 07:00 and lights off at 19:00.
Northern blot analysis
Ten to fourteen week old mice were sacrificed by cervical dislocation. RNA was isolated from whole brain using Trizol reagent (Invitrogen). 20 μg of total RNA was electrophoresed in a 1.9% formaldehyde/1% agarose gel and blotted onto Hybond-N (Amersham Pharmacia). Blots were first probed with a 32P-labeled γ2 cDNA probe then re-hybrizided with a control human β-actin probe (Clontech). Autoradiographs were digitally photographed and band densities were measured using Kodak 1D Image Analysis Software. Ratio between density of γ2 hybridization versus actin hybridization was recorded.
Immunocytochemistry
The procedure has been described elsewhere [47]. Briefly, wild type and homozygous mutants were deeply anesthetized with Avertin and transcardially perfused with Dextran-Phosphate Buffer (PB) then fixative consisting of 0.01M periodate/0.075M lysine/4% paraformaldehyde in 0.1M PB (pH 7.4). The brains were frozen and sliced in parasaggital sections (25 μm) with a freezing microtome. Slices were incubated overnight at 4°C with affinity-purified rabbit anti-γ2 [48,49] in 0.1M PB (pH 7.4) with 0.3% Triton X-100, at a concentration of (1:100). The sections were processed using an avidin-biotin-peroxidase system (Vectastain Elite; Novocastra, Burlingame, CA). Peroxidase reaction was carried out with 3-3' diaminobenzidine tetrahydrochloride in the presence of cobalt chloride and nickel ammonium sulfate as chromogens and hydrogen peroxide as oxidant. Controls were performed either by omitting the primary antibody or by incubating the primary antibody with the corresponding peptide [50].
Pharmacology
Eight-week-old mice were sacrificed by decapitation and whole brains were rapidly dissected out and frozen on dry ice. For autoradiography, 14-μm horizontal serial cryostat sections were cut from 5 mouse brains of each genotype, thaw-mounted onto gelatin-coated object glasses, and stored frozen under desiccant at -20°C. All experiments were carried out in parallel fashion with respect to mouse lines, eliminating any day-to-day variation between the assays. The autoradiographic procedures for regional localization of [3H]Ro 15-4513, [3H]muscimol, and [35S]TBPS binding were as described in [51]. Briefly, sections were preincubated in an ice-water bath for 15 min in 50 mM Tris-HCl (pH 7.4) supplemented with 120 mM NaCl in the [3H]Ro 15-4513 and [35S]TBPS autoradiographic assays, and in 0.31 M Tris-citrate (pH 7.1) in the [3H]muscimol assay. All radioligands were purchased from Perkin Elmer Life Sciences, Inc. (Boston, MA, USA).
The final incubation in respective preincubation buffer was performed with 6 nM [35S]TBPS at room temperature for 90 min, assays with 10 nM [3H]muscimol at 0–4°C for 30 min, and assays with 10 nM [3H]Ro 15-4513 in the presence and absence of 100 μM diazepam (Orion Pharma, Espoo, Finland) at 0–4°C for 60 min. After incubation, sections were washed 3 × 15 s or 2 × 30 s in an ice-cold incubation buffer in [35S]TBPS and [3H]Ro 15-4513 or in [3H]muscimol assay, respectively. Sections were then dipped into distilled water, air-dried under a fan at room temperature, and exposed with plastic [3H]- or [14C]-methacrylate standards to Kodak Biomax MR films for 1 to 8 weeks. Nonspecific binding was determined with 10 μM flumazenil (Hoffmann-La Roche, Basel, Switzerland), 100 μM picrotoxinin (Sigma) and 100 μM GABA (Sigma) in [3H]Ro 15-4513, [35S]TBPS and [3H]muscimol assays, respectively. Binding densities in the relevant brain areas were quantitated with MCID M5-imaging software (Imaging Research Inc., Ontario, Canada) and converted to nCi/mg (for 3H) or nCi/g (for 35S) radioactivity values on the basis of the simultaneously exposed standards.
The concentrations of [3H]muscimol (10 nM) and [3H]Ro 15-4513 (10 nM) were greater than or equal to the dissociation constants for a range of recombinant and native GABAA receptors [8,52-54]. Therefore, the autoradiographic images should represent the density rather than affinity of binding sites.
Hypnotic sensitivity
Drug induced hypnosis was assessed by measuring the duration of the loss of righting reflex in response to various sedative/hypnotic drugs. Diazepam (25 mg/kg; Sigma), midazolam (45 mg/kg and 75 mg/kg; ESI Lederle, Philadelphia, PA), pentobarbital (45 mg/kg; Abbott, Chicago, IL), zolpidem (60 mg/kg; Searle, Malvern, PA) or ethanol (3.5 mg/kg; Pharmco, Brookfield, CT) were administered intraperitoneally to wild type and homozygous mutants. Drugs were diluted in saline such that the injection volume was 20 μL per gram body weight. After losing the righting reflex, mice were placed in a plastic V-shaped trough and the time was recorded. When the mouse was able to right itself three times in 30 s, the measure of hypnotic effect was over. Body temperature was maintained with aid of a heat lamp. Assays were performed in a blinded manner with respect to genotype. Effect of genotype on the duration of the loss of righting reflex was compared using Student's t-test.
Elevated plus-maze test
Basal anxiety-like behavior was tested using the elevated plus-maze. All mice were between 7 and 9 weeks of age and were tested between 09:00 and 11:00. Each mouse was placed on the central platform of the maze, facing an open arm and allowed to freely explore the maze for 5 min under ambient room light. Open-arm and closed-arm entries and the cumulative time spent on the open and closed arms was recorded. A mouse was considered to be on the central platform or on an arm when all four paws were within its perimeter. The percent open-arm entries, total number of entries, and percent time in open-arms was determined. Data were analyzed using one-way ANOVA.
Forced exploration
Basal motor activity of mice was determined using the forced exploration test. 8–10 week old mice were placed into a walled arena (43.2 cm × 43.2 cm × 30.5 cm) for 5 min. Distance traveled (cm) was measured automatically using an Activity Monitor (Med Associates, St. Albans, VT). All tests were performed between 12:00 and 15:00. Effect of genotype on basal motor activity was compared using Student's t-test.
Authors' contributions
CPM and ALD carried out the immunohistochemical studies. ERK carried out the ligand autoradiography studies. All other experiments were performed by DC and GEH. All authors contributed to composing and editing the manuscript and have read and approved the final version.
Acknowledgements
Carolyn Ferguson and Ed Mallick are gratefully acknowledged for superb technical support. This work was supported by NIH grants AA10422, GM52035, GM47818, NS039287, the Sigrid Juselius Foundation, and the Academy of Finland.
Figures and Tables
Figure 1
Gene targeting of the γ2 subunit of the GABAA-R and development of γ2 knockdown mice. (A) Targeting strategy used to produce γ2 knockdown mice. Relevant region of endogenous gene (+), targeting vector, correctly targeted knockdown γ2 (F), and cre-recombined knockout alleles (f) are shown. Relative locations of relevant restriction sites, exons (numbered orange boxes), loxP sites (blue triangles), plasmid backbone (wavy line), and positive (NEO) and negative (PGK-TK) selection cassettes are shown. (B) Southern Blot analysis of BglII digested tail DNAs. Probe A hybridizes to a 9.2 kb BglII fragment from the wild type endogenous allele and a 3.4 kb BglII fragment from the targeted allele.
Figure 2
Northern blot analysis. (A) Sample northern blot analysis of wild type (+/+) and homozygous knockdown (F/F) adult whole brain total RNA hybrizided with a γ2 cDNA or a human β-actin probe as a loading control. (B) Densitometric analysis of northern blots. The ratio of band density between γ2 and β-actin hybridization for wild type (n = 23) and homozygous knockdown (n = 36) mice was plotted. On each individual blot, the ratio of wild type band densities was averaged and normalized to 100. All samples on each blot were then compared to the average of wild type band densities on that blot. A total of 4 different blots were included in this analysis. The horizontal bar in each column represents the mean for that genotype.
Figure 3
Immunohistochemical distribution of γ2 subunit of the GABAA-R in sections from individual (A) wild type and (B) homozygous knockdown mice. Note the high level of abundance of γ2 immunoreactivity in olfactory bulb (OB), cerebral cortex (CC), hippocampus (HP), substantia nigra (SN), and cerebellum (CB) of wild type samples. The amount of γ2 is variably reduced in many brain regions of the knockdown samples.
Figure 4
Behavioral characterization of knockdown mice. (A-C) Elevated plus maze. Knockdown mice (A) entered open arms less often and (B) for less time, however, (C) total entries into arms did not differ from wild type mice. (D) Knockdown mice were also less active in the forced exploration test. The bars are means ± SEM. *p < .01, **p < .001.
Figure 5
Hypnotic sensitivity. No significant differences in the hypnotic effects of ethanol, pentobarbital, zolpidem, midazolam, or diazepam were observed. The bars are means ± SEM.
Table 1
Autoradiographic analysis of GABAA receptor binding sites in horizontal sections of wild type and mutant mice.
The data are means ± standard deviations for 5 mice in each genotype, and are expressed as nCi/mg for 3H-ligands and as nCi/g for 35S-ligand. Significance of the difference from wild type (Tukey-Kramer test after ANOVA): aP < 0.05, bP < 0.01, cP < 0.001. Ctx, cortex; Th, thalamus; Hi, hippocampus; CPu, caudate putamen; IC, inferior colliculus; Gr, cerebellar granule cell layer; Mol, cerebellar molecular layer. | [
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15882093 | Neuronal Migration and Ventral Subtype Identity in the Telencephalon Depend on SOX1
Abstract
Little is known about the molecular mechanisms and intrinsic factors that are responsible for the emergence of neuronal subtype identity. Several transcription factors that are expressed mainly in precursors of the ventral telencephalon have been shown to control neuronal specification, but it has been unclear whether subtype identity is also specified in these precursors, or if this happens in postmitotic neurons, and whether it involves the same or different factors. SOX1, an HMG box transcription factor, is expressed widely in neural precursors along with the two other SOXB1 subfamily members, SOX2 and SOX3, and all three have been implicated in neurogenesis. SOX1 is also uniquely expressed at a high level in the majority of telencephalic neurons that constitute the ventral striatum (VS). These neurons are missing in Sox1-null mutant mice. In the present study, we have addressed the requirement for SOX1 at a cellular level, revealing both the nature and timing of the defect. By generating a novel Sox1-null allele expressing β-galactosidase, we found that the VS precursors and their early neuronal differentiation are unaffected in the absence of SOX1, but the prospective neurons fail to migrate to their appropriate position. Furthermore, the migration of non-Sox1-expressing VS neurons (such as those expressing Pax6) was also affected in the absence of SOX1, suggesting that Sox1-expressing neurons play a role in structuring the area of the VS. To test whether SOX1 is required in postmitotic cells for the emergence of VS neuronal identity, we generated mice in which Sox1 expression was directed to all ventral telencephalic precursors, but to only a very few VS neurons. These mice again lacked most of the VS, indicating that SOX1 expression in precursors is not sufficient for VS development. Conversely, the few neurons in which Sox1 expression was maintained were able to migrate to the VS. In conclusion, Sox1 expression in precursors is not sufficient for VS neuronal identity and migration, but this is accomplished in postmitotic cells, which require the continued presence of SOX1. Our data also suggest that other SOXB1 members showing expression in specific neuronal populations are likely to play continuous roles from the establishment of precursors to their final differentiation.
Introduction
The telencephalon is subdivided into dorsal (pallial) and ventral (subpallial) territories, which give rise to the cerebral cortex and the underlying basal ganglia, respectively. The embryonic subpallium consists of large protrusions—the ganglionic eminences. Several distinct types of neurons originate in the ganglionic eminences, and some migrate as far as the olfactory bulb, hippocampus, and neocortex [1–3], while others contribute more locally. The majority of neurons of the lateral ganglionic eminence (LGE) form the dorsal and ventral striatum (VS). The VS includes the caudate, putamen, nucleus accumbens, and olfactory tubercle (OT), which control various aspects of motor, cognitive, and emotional functions [4,5]. Little is known about the molecular mechanisms that control the emergence of various groups of neurons with distinct identities in this region.
Gene-expression studies and loss-of-function mutations in homeodomain transcription factors such as PAX6 [6,7] and GSH2/1 [8–13] confirm fate-mapping findings [14–16] that the majority of the VS neurons are specified within the progenitor domain of the LGE. The proneural basic helix-loop-helix (bHLH) factor MASH1 also marks the precursors of early-born neurons in the LGE progenitor domain, and its loss in the mouse leads to a deficit of both precursors and neurons of the telencephalon, including loss of VS neurons [17,18]. Therefore, GSH2 and MASH1 control VS precursor patterning and specification, but as they are not expressed in postmitotic cells it remained unknown to what extent they are involved in the emergence of neuronal subtypes in the ventral telencephalon, and whether different transcription factors with neuron-specific expression are required.
The SOX proteins constitute a family of transcription factors [19,20] that regulate transcription through their ability to bind to specific DNA sequences via their HMG box domains [21–24]. There are 20 Sox genes in mammals, and at least half are expressed in the developing nervous system [20,24]; however, their role in neural development is poorly understood. SOX1, SOX2, and SOX3 constitute the SOXB1 subfamily and share more than 95% identity within their HMG boxes and significant homology outside [25,26]. All three proteins are expressed in the neuroepithelium throughout central nervous system (CNS) development [25,27], and as they tend to be down-regulated upon neural differentiation they have been used as markers for neural stem cells and precursors [28,29]. Several studies suggest that SOXB1 factors function in stem cells and precursors to maintain broad developmental potential [30] and neural stem cell identity [30–32] by counteracting neurogenesis. Contradictory evidence, however, suggests that SOX1 promotes neurogenesis and cell cycle exit [33]. However, mice that are null for Sox1 [34] or Sox3 [35], or mice with one Sox2 allele deleted and the other hypomorphic [36], exhibit phenotypes associated with the loss of or functional deficit of only specific neuronal populations. As these SOXB1 factors are expressed in both precursors and neurons that are affected in these mutant mice, it was not known whether their function is required in precursors, postmitotic cells, or both.
We have previously shown that SOX1 is essential for the terminal differentiation of lens fibers and the activation of γ-crystallins [37], and for the development of VS neurons, the lack of which is associated with epilepsy [34]. Here, we show that absence of SOX1 has no effect on the generation, proliferation, and patterning of neuronal precursors. This is probably due to functional compensation by SOX2 and SOX3, which are co-expressed with SOX1 in precursors. Moreover, mice lacking only the neuron-specific expression of Sox1 in the ventral telencephalon still fail to develop VS neurons, revealing its requirement within these neurons. Consistent with this, maintenance of Sox1 expression in neurons of the ventral telencephalon is sufficient to direct them to the VS, confirming the adequacy of SOX1 function in postmitotic cells for their migration and identity. Therefore, VS-specific neuronal migration and subtype identity most likely is initiated in precursors but is completed in postmitotic cells by transcription factors such as SOX1.
Results
SOX1 Is Essential for the Histogenesis of the VS
To generate a detailed map of Sox1 expression in the developing and adult brain of mice, and to perform comparative studies between homozygotes and heterozygotes, we generated a novel targeted allele referred to as Sox1βgeo. This contains an insertion of β-galactosidase-neo (βgeo) fusion protein in-frame with the SOX1 open reading frame (Figure 1A). Mice homozygous for Sox1βgeo are null for SOX1 and exhibit the same phenotype as the previously described mice, which carry a deletion of the SOX1 coding region (Sox1M1) [34,37], namely, lens defects and epileptic seizures. Staining for β-galactosidase activity (X-gal staining) in Sox1βgeo/+ heterozygous embryos matches that for the wild-type allele as revealed by whole-mount in situ hybridization and SOX1 antibody staining (Figure 1B–1E).
To elucidate the role of SOX1 in the formation of the VS, we compared the expression pattern of Sox1 in heterozygous (Figure 1F–1I) and homozygous (Figure 1J–1M) brains from embryonic day 14 (E14) to postnatal day 0 (P0) using X-gal staining. Throughout much of the CNS, X-gal staining in the Sox1βgeo/βgeo homozygotes is double the intensity observed in heterozygotes (data not shown). To perform comparative histological studies, we equalized the levels of X-gal staining in homozygous mice with those of heterozygous animals by generating homozygous mice that harbor two different Sox1-null alleles: βgeo (Sox1βgeo) and the previously described M1-targeted allele (Sox1M1) [37], which does not express β-galactosidase.
Our analysis shows (via X-gal staining) that in the developing forebrain, Sox1 is expressed throughout the ventricular zone (VZ) and subventricular zone (SVZ) and in neurons around the anterior commissure region, where the prospective nucleus accumbens forms (red arrowheads in Figure 1K and 1M), and in the striatal bridges that link this intermediate cluster of cells with the prospective OT region toward the pial surface. X-gal-positive neurons start populating the OT area as early as E14 and continue to accumulate at least until birth (Figure 1F–1I). In the Sox1βgeo/M1-null mutants, the X-gal staining pattern of the VZ/SVZ is indistinguishable from that of the Sox1βgeo/+ heterozygotes, and there is no obvious deficit of X-gal-positive cells around the anterior commissure. On the other hand, both the striatal bridges and the OT layers are absent in the Sox1-null brain at all developmental stages (compare Figure 1F–1I to 1J–1M). It is unlikely that neurons die en masse in this region, because an apoptosis assay did not reveal any evidence of increased cell death in the mutant (data not shown). In addition, X-gal staining is increased throughout the ventral telencephalon in the mutant postnatal brain (red arrowheads in Figure 2), suggesting that the Sox1-null cells are not correctly specified and contribute to other brain regions. Interestingly, although neurons that form the core of the nucleus accumbens express Sox1 highly, they form normally (Figure 2F and red arrowheads in Figure 1) and do not depend on SOX1 for their development. Therefore, SOX1 is required for histogenesis of the OT throughout its development.
Normal Precursor Proliferation and Neurogenesis but Loss of OT Neuronal Differentiation in the Absence of SOX1
Studies with conflicting results suggest that SOX1 either, like SOX2 and SOX3, counteracts neurogenesis [32] or, unlike SOX2 and SOX3, promotes neurogenesis [33]. To examine whether the loss of SOX1 affects general neural differentiation in the area of the striatum, we used an anti-βIII-tubulin (TuJ1) antibody, which is a marker for immature neurons [38], at E13, a critical time of differentiation in the LGE. TuJ 1 immunocytochemistry did not reveal any obvious general differentiation problems in the Sox1 mutants (Figure 3A and 3B), suggesting that loss of SOX1 alone is not sufficient to compromise general neuron differentiation and maturity. The differentiation and distribution of specific mature neurons was examined in our previous study at adult stages with the expression of striatal markers such as preproenkephalin and Gad65/67 [34], and in our current study, at embryonic stages with additional markers such as Brn4 [39] (Figure 3C and 3D) and Robo [40] (Figure 3E and 3F). This analysis revealed a differentiation defect restricted in the region of the nucleus accumbens/OT, and not the rest of the striatum.
As Sox1 expression is associated with dividing cells throughout the neuroepithelium, we examined whether a proliferation defect could partially account for the cell deficit in the ventral telencephalon. We used 5-bromo-2′-deoxyuridine (BrdU) to label all proliferating precursors in wild-type and Sox1-null embryos at E13–E15, and harvested their brains 1 h later. Most of the dividing cells were found in the VZ/SVZ, and there was no increase or ectopic proliferation (Figure 3G–3L; Table S1). Therefore, SOX1 is unlikely to be required for proliferation or the exit of precursors from the cell cycle in the LGE. Collectively, the above data show that SOX1 is not essential for the proliferation of precursors and general neuronal differentiation. However, it is required specifically for the differentiation and/or migration of VS neurons.
Early- and Late-Born OT Neurons Fail to Migrate in the Absence of SOX1
To investigate the possibility that neurons migrate in the Sox1-null OT regions, but are not visible because they do not express Sox1βgeo and other differentiation markers, we exposed embryos to BrdU. This way, we permanently marked all proliferating precursors independently of Sox1 or other striatal-marker gene expression and followed them at later embryonic stages and after birth (Figure 4). Labeling the precursors at different embryonic days also provided information on the birthdates of the OT neurons, which were previously unknown for the mouse. Specifically, birth of ventral striatal neurons commences early at E13 and continues until birth (Figure 4A, 4C, 4E, and 4G; data not shown) and is consistent with data from the rat [41,42]. Furthermore, in wild-type embryos, BrdU exposure between E13 and E16 with examination of embryos either 72 h later (Figure 4A; data not shown) or after birth (Figure 4C, 4E, and 4G) showed that early-born neurons migrate more laterally than those born later.
In the Sox1-null embryos, the presence and distribution of cells labeled with BrdU between E13 and E16 shows that the olfactory cortex is largely normal (bracket in Figure 4D), but in the VS area the number of labeled cells was found to be greatly reduced (Figure 4B, 4D, 4F, and 4H). Furthermore, in the Sox1-null postnatal brain the striatal mantle is more densely populated by BrdU-labeled cells (Figure 4D–4H), consistent with the general increase of X-gal staining observed throughout the striatum (see Figure 2). The above data suggest that in the absence of SOX1, early- and late-born neurons fail to migrate to the appropriate position to form the ventral areas of the striatum.
The Generation and Patterning of LGE Precursors Is Normal in the Absence of SOX1
It is known that the majority of VS neurons derive from precursors that are born in the LGE [10,15,43]. To investigate whether the defect is in the patterning of precursors, we examined the expression pattern of various transcription factors that mark LGE progenitors and are known to have a role in OT neuronal specification (Figures 5, S1, and S2).
The homeodomain transcription factors PAX6 and GSH2 are expressed, respectively, in the pallial and subpallial precursor domains of the dorsal LGE. The boundary between them has been shown to be essential for the patterning of VS precursors [10,11]. Specifically, GSH2-null mice do not form early-born OT neurons, and the precursors of the dorsal LGE are lost as PAX6 expands ventrally into the LGE. However, loss of both PAX6 and GSH2 restores dorsoventral patterning and partially rescues OT formation [9–11]. Using double antibody immunohistochemistry in embryos at E12–E16, we found that loss of SOX1 has no effect on the expression of GSH2/PAX6 and the boundary in the dorsal LGE (Figures 5A, 5B, and S1). In addition, at this boundary, PAX6-expressing postmitotic cells form a stream (arrow in Figure 5A) that extends laterally to the VS (Figures 5A, 5C, S2E, and S2F) [6,7,44]. In the absence of SOX1, the stream of PAX6-positive postmitotic cells is normal (arrow in Figures 5B and S2F), but to characterize the PAX6- and SOX1-expressing neurons in the region of the OT, we used double antibody immunohistochemistry. Specifically, for the wild-type brain sections we used PAX6 and SOX1 antibodies, but to visualize the Sox1-expressing cells in the Sox1βgeo/M1-null brain sections we used an antibody for β-galactosidase.
Our data showed that PAX6 and SOX1 proteins were co-expressed in progenitors (Figures S2E and S2F), but in postmitotic cells of the LGE this expression became mutually exclusive (Figure 5C and 5D). Pax6-expressing neurons were clustered laterally to those expressing Sox1, at the border between the OT and olfactory cortex (Figures 5C, S2A, and S2C). In Sox1-null mice, the postmitotic Pax6-expressing cells were distributed throughout the VS area (Figures 5D, S2B, and S2D), suggesting structural disorganization. It is unlikely that these ectopically localized Pax6-expressing neurons are mis-specified Sox1-null neurons, because they should be expressing both PAX6 and βgeo from the mutant Sox1βgeo allele, but they do not (Figure 5D).
The LGE structure consists of neural progenitors with radial glial characteristics, having fibers that extend from the VZ to the pial surface. These cells also provide the substrate for the migration of neurons [45,46]. Staining with X-gal (see Figure 1) and β-galactosidase antibody in mice carrying the Sox1βgeo allele allowed visualization of the cytoplasmic compartment of the SOX1-expressing progenitors, which have radial glial morphology (Figures S2F and S3). We examined the morphology of radial glia in the Sox1-null LGE using the RC2 antibody [47], but we did not find any difference from wild-type (Figure S4). Therefore, we conclude that the abnormal distribution of Pax6-expressing neurons in the LGE of the Sox1-null mice is unlikely to be caused by abnormal morphology of the radial glial fibers or substantial loss of radial glia-like precursors.
In the ventral telencephalon, the bHLH transcription factor MASH1 [18] and the homeodomain factor DLX1 mark LGE precursors [8]. Ablation of MASH1 in mice causes loss of specific subpopulations of precursors and striatal neurons that contribute to the OT and nucleus accumbens [18]. In addition, Gsh2-null mice, which also fail to develop the OT, exhibit reduced Dlx1 expression in LGE precursors [9]. We therefore examined the expression of these two genes in Sox1-null embryos, but found no difference (Figure 5E–5H), indicating that there is no deficit of early or late LGE precursors in the absence of SOX1. Collectively, the above data show that SOX1 is not required for patterning, generation, and maintenance of LGE precursors.
Sox1 Expression from the Endogenous Sox2 Promoter Can Be Tolerated In Vivo
It has been shown that all three SoxB1 genes are expressed [25,27] and share similar functions in neural precursors [31,32]. However, in the postmitotic cells of the mantle and the VS area, antibody staining for each of the genes at E15 indicated that SOX2- and SOX3-positive neurons represent a very small population compared to that expressing SOX1 (Figure 6A–6C). Therefore, it is likely that the other SoxB1 genes compensate for the loss of Sox1 in precursors, whereas they cannot do so in the LGE postmitotic cells. Nevertheless, it remained unknown whether SOX1 functions solely in precursors for VS fate specification or in postmitotic cells for maintaining this fate and the emergence of specific subtype identity and migration. To address this, we generated mice that express Sox1 mainly in precursors and not in LGE neurons. We took advantage of the fact that Sox2 is co-expressed with Sox1 in precursors but it is down-regulated in LGE neurons, and generated mice that express Sox1 from the endogenous Sox2 promoter. We confirmed the overlap of Sox1 and Sox2 expression in the VZ/SVZ of the LGE by staining serial coronal telencephalic sections with antibodies for each of the two genes and counter-staining with the nuclear stain TOTO at E14 (Figure S5), and by performing double anti-SOX1 and -SOX2 immunohistochemistry at E13 (Figure 6D–6L).
The replacement of the SOX2 open reading frame with that of SOX1 was achieved by targeting the Sox2 allele (Figure 7A). The new allele, Sox2R, was engineered to express not only Sox1 but also βgeo via an internal ribosomal entry site (IRES). Furthermore, the coding region of SOX1 in the Sox2R allele was flanked by LoxP sites, which can be deleted using Cre-mediated recombination [48]. In this way, we generated a mouse line carrying another allele, termed Sox2βgeo2 (but referred to hereafter as Sox2βgeo), which expresses only the βgeo reporter gene from the Sox2 promoter (Figure 7A) and not SOX1. Like the Sox2βgeo/+ heterozygotes, Sox2R/+ mice were viable, fertile, and phenotypically normal, indicating that SOX1 over-expression in precursors, as well as ectopic expression in other locations where Sox2 is uniquely expressed, does not cause any obvious developmental abnormality.
X-gal staining of embryos showed that both targeted Sox2 alleles (Sox2βgeo and Sox2R) express βgeo in the CNS, but to verify that SOX1 protein was produced from the Sox2R allele, we used SOX1 antibody staining. We found that SOX1 protein was ectopically present at sites where SOX2 normally shows unique expression—for example, in the floor plate of the diencephalon (arrowheads in Figure 7B and 7C) and in the sensory placodes (arrows in Figure 7D and 7E). The intensity of the immunostaining at ectopic sites was comparable to the staining in areas with expression from two wild-type Sox1 alleles (VZ/SVZ), indicating that the level of expression was similar to the wild-type allele. Therefore, the Sox2R allele produces SOX1 ectopically in all neurons uniquely positive for SOX2 and increases the endogenous level of SOX1 in precursors and neurons that express both genes, without causing an obvious defect in mice. The fact that ectopic expression does not cause any obvious phenotype suggests either that the partner factors required for SOX1 target specificity [19] are absent in those cells uniquely expressing Sox2 or that the two proteins are interchangeable, sharing target genes. Further experiments are required to clarify this.
Sox1 Over-Expression in Precursors Does Not Increase VS/OT Neuronal Fate Specification
To investigate more subtle defects due to the over-expression of Sox1 in precursors of the Sox2R/+ mice, we examined several litters (n > 10) of mice and visualized the migration of the Sox1/Sox2-positive neurons in the VS via X-gal staining. Newborn mice carrying Sox2R/+ with two (Sox2R/+, Sox1+/+; Figure 8A) or one (Sox2R/+, Sox1M1/+; Figure 8B) Sox1 endogenous wild-type alleles were compared with those carrying Sox2βgeo/+, Sox1+/+ that do not express Sox1 ectopically (Figure 8C). All the above mice have only one wild-type Sox2 allele. We also compared the Sox1-βgeo and Sox2-βgeo neurons of the ventral telencephalon in Sox1βgeo/+ (Figure 8C and 8D) and Sox2R/+ (Figure 8A) mice, respectively. In this area of the brain, the Sox2-positive neurons are far fewer than those positive for Sox1. Therefore, for comparison purposes, we used thin tissue sections (80 μm) with short X-gal staining (3 h) for Sox1-βgeo, and thicker sections (100–150 μm) with a long staining period (48 h) for Sox2-βgeo. Consistent with the antibody staining data (see Figure 6B), Sox2-βgeo neurons contribute to the OT, indicating that they are a subset of the VS neuronal population. Therefore, the ectopic expression of Sox1 in LGE neurons is expected to be very limited. More importantly, the migration and the number of LGE neurons expressing βgeo via the Sox2 promoter were found to be the same regardless of the number of endogenous Sox1 alleles or the ectopic presence of Sox1 (compare Figure 8A, 8B, and 8C). To further investigate the differentiation of the VS neurons, we used the striatal-specific markers dopamine and cAMP-regulated phosphoprotein (DARPP-32) at postnatal stages and found them to be unaffected in Sox2R/+ mice (Figure 8E and 8F). The data therefore indicate that the over-expression of SOX1 in precursors does not increase OT neuronal specification.
Sox1 Expression in Precursors Cannot Rescue OT Neuron Development
To address directly whether SOX1 function is essential in precursors, we crossed Sox1M1/+, Sox2R/+ mice with Sox1βgeo/+, Sox2+/+ mice and examined whether offspring carrying Sox2R/+ without any wild-type Sox1 alleles (Sox1M1/βgeo) could develop OT. In these Sox1R/+ embryos that carry no endogenous Sox1 functional allele (termed here HoHe), SOX1 is expected to be expressed only via the Sox2R allele in precursors and become down-regulated in postmitotic LGE cells. However, βgeo expression from the endogenous Sox1 mutant allele marks precursors and OT prospective neurons. We followed the Sox1M1/βgeo prospective OT neurons with X-gal to determine whether they were capable of contributing to the OT in HoHe embryos (Figure 9). The Sox2R allele also expresses βgeo; however, in LGE postmitotic cells, Sox2-βgeo expression is much less than that of Sox1-βgeo and is not very visible by short (3 h) X-gal staining (only a slight increase of X-gal staining is seen in the VS; Figure 9C compared to Figure 9B).
Each brain was split into left and right hemispheres, and coronal sections of the left were used for short staining with X-gal (Figure 9A–9C) whereas sections of the right were stained with SOX1 antibody (Figure 9D and 9E). The hemispheres stained for X-gal showed characteristic staining of OT neurons in heterozygous Sox1βgeo/+ mice (red arrowheads in Figure 9A), but the hemispheres of the Sox1-null embryos (Sox1βgeo/M1) with Sox2R/+ (Figure 9B) or without (Figure 9C) did not. This indicates that Sox1-null embryos do not develop OT despite the presence of the Sox2R allele and SOX1 protein in progenitors. To verify the presence of SOX1 protein in the precursors of the HoHe mice, we examined the other hemisphere that was stained with SOX1 antibody. In the Sox1βgeo/+ embryos, we found SOX1 present in the VZ (yellow arrows in Figure 9D and 9E) and the OT neurons (red arrowheads in Figure 9D). In Sox1βgeo/M1 (null) embryos, SOX1 expression was completely absent (data not shown); in the HoHe embryos, SOX1 protein was present in the VZ (yellow arrow in Figure 9E) and in very few neurons of the LGE (Figure 9E). HoHe mice, like the Sox1-nulls, are born with small eyes, and around weaning age develop seizures associated with lethality, which, if anything, is increased compared to that of Sox1-null mice (data not shown). In the brain of P10 HoHe mice, we used staining with DARPP-32 antibody (which is a SOX1-independent striatal marker) to investigate the recovery of OT neurons, and found staining in the striatal mantle but not in the VS (Figure 9G compared to Figure 9F). We therefore concluded that SOX1 expression in precursors is not sufficient to rescue VS/OT neuron fate specification, and that the continued presence of SOX1 in postmitotic cells is required for their identity.
Sox1/Sox2 Expression in Neurons Is Sufficient for Their Migration to the VS
We have shown that in mice carrying two (Sox1+/+), or one (Sox1M1/+), Sox1 wild-type alleles (see Figure 8A, 8B, and 8C), the migration of the Sox2-positive LGE neurons is not overtly different from that observed in mice carrying the Sox2R/+ allele. However, it remained unknown whether the Sox1/Sox2 double-positive LGE neurons migrated to the VS when both Sox1 endogenous alleles were missing (HoHe mice). We used X-gal staining to follow these neurons in several litters (n > 10), including HoHe mice, which have two Sox1M1 alleles and thus βgeo expression exclusively driven by the Sox2R allele. We found that in the LGE of these mice, the double-positive neurons are generated and migrate to the OT area (compare Figure 10A and 10B), but this area is compacted in the absence of the majority of the OT/SOX1 neurons. The above data show that the continued expression of Sox1 in neurons of the LGE is sufficient to direct their migration to the OT in the absence of endogenous Sox1.
Discussion
The specification of neurons in the ventral telencephalon has been shown to depend on several transcription factors that are expressed mainly in proliferating precursors. However, it was unknown to what degree specification in precursors included the emergence of neuronal subtype identity in the ventral telencephalon, and whether expression of additional transcription factors was required. We showed that the differentiation and migration of early- and late-born neurons that constitute the VS require SOX1 expression not only in precursors but also in postmitotic cells. Furthermore, in this region, the migration and organization of other neurons such as those expressing Pax6 also depend on the presence of SOX1-positive VS neurons. The finding that SOX1 functions in neurons to control migration and identity is novel and suggests that the other SOXB1 factors, in addition to their roles in precursors, have similar functions in neurons.
Identity and Migration of Neurons in the VS
The development of subtype identity and migration of neurons in the ventral telencephalon has not been well characterized. The expression of differentiation markers reveals neurons in both VS and dorsal striatum, but we showed that SOX1 specifically marks a large population of VS neurons that form the principal layer II of the OT, the islands of Calleja, and the nucleus accumbens (see Figures 1 and 2). In addition, we showed that the neurons expressing Sox1 are born continuously from E13 until the first postnatal week and that these migrate to a ventrolateral region of the telencephalon, with later-born neurons positioned progressively to more medial positions (see Figure 4). In the absence of SOX1, the majority of neurons of the VS fail to develop. All Sox1-expressing neurons of the OT and the islands of Calleja require SOX1 for their development, but it is essential only for the shell of the nucleus accumbens, although the core also expresses it. While neurons of the adjacent striatal mantle and the olfactory cortex that do not express Sox1 develop normally in its absence, other groups of neurons within the VS appear to be disorganized. Specifically, we identified a distinct population of neurons located lateral to the OT at the border with the olfactory cortex that expresses Pax6, but not Sox1. In the absence of SOX1, these neurons migrate into more medial positions, occupying the space of the missing OT neurons (see Figure 5). These are not mis-specified Sox1-null neurons because they do not express βgeo. This indicates that Sox1-expressing OT neurons play a non-cell-autonomous role in the organization of other neurons in this region, including the production of essential signals for migration. Most likely, the disorganization of the VS in the absence of SOX1 results in abnormal local neuronal connectivity, which in turn leads to the abnormal (epileptiform) electrophysiological behavior observed in the SOX1-deficient animals [34].
SOX1 Function in Precursors
SOXB1 factors share considerable homology in both their DNA binding and C-terminal transcriptional activation domains, and they are co-expressed in precursors. It is therefore possible that in the LGE precursors, the role of SOX1 in the specification of OT/VS neurons is redundant. However, as SoxB1 genes have a broad expression in the neuroepithelium, we have to assume that their specific function at different areas of the VZ, and particularly the VZ of the LGE, is controlled by the presence of LGE-specific partner factors. SoxNeuro and Dichaete, the two Drosophila orthologs of the vertebrate SoxB1 group genes, also show overlapping functions during neural development [49]. Furthermore, in Drosophila, these two genes have been shown to genetically interact with the dorsoventral patterning genes ind (intermediate neuroblast defective) and vnd (ventral nerve chord defective) [50,51]. The vertebrate orthologs of ind and vnd are Gsh1/2 [52,53] and Nkx2.2 [54], respectively. In the mouse, Gsh2 is expressed in the VZ/SVZ, and like Sox1, its loss results in a reduction of VS neurons. As target gene specificity of SOX proteins depends on partnering with other transcription factors [20], our work, along with the data from Drosophila, supports the hypothesis that in the LGE precursors GSH1/2 may act as partners for SOXB1 factors to initiate ventral telencephalic neuronal identity.
The neurons of the VS area occupy approximately a quarter of the striatal mass [55], and migrate there over a period of at least 10 d (E13 to first postnatal week). The LGE precursors that generate the OT/VS in the LGE are expected to have an equivalent representation during this period of development. In Gsh2-null mice, which are missing early-born OT neurons, there is a deficit of precursors in the LGE, readily seen by the reduced expression of Dlx1/2 in LGE precursors [9–11]. In Sox1-null mice, the neuronal deficit is more severe than that in Gsh2 mutants, as it includes both early- and late-born OT/VS neurons. However, BrdU labeling and LGE precursor-specific marker analysis, including Gsh2, Dlx1/2, Mash1, and Pax6, in Sox1-null brains at different stages did not show any deficit in precursors. The increase of X-gal-stained (Sox1-βgeo; see Figure 2) and BrdU-labeled neurons (see Figure 4D and 4F) in the area of the septum and the striatum supports the notion that the VS/OT neurons are born but lack VS subtype identity to migrate toward ventral positions. The normal expression of TuJ1, a marker of immature neurons, excluded the possibility that loss of SOX1 delays or enhances differentiation. Therefore, in the absence of SOX1, the precursors are there and generate neurons, but these fail to migrate to the VS because they assume different identity and position. The finding that Sox1-null neurons contribute widely to different areas argues that the presence of SOX1 provides neurons with ventral identity and the ability to migrate to ventral regions.
Emergence of VS Neuron Identity
To test the role of Sox1 expression in neurons and to determine whether ventral identity emerges in postmitotic cells, we limited expression of Sox1 largely to precursors of LGE neurons. Sox2R/+ mice express Sox1 from one of the Sox2 alleles. When the Sox2R allele is present in animals with no endogenous Sox1 wild-type alleles (HoHe), SOX1 expression mimics that of SOX2—being present in VS/OT precursors but largely absent from the neurons they give rise to. HoHe mice also fail to develop the majority of VS/OT neurons (see Figure 9) and exhibit an equally severe phenotype to that of Sox1-null mice in the OT. As these mice reproduce faithfully the Sox1-null phenotype without any evidence of a partial rescue, it is unlikely that this is the result of incomplete expression of Sox1 from the Sox2 promoter in the precursors. However, to exclude the possibility that the failure of OT/VS neuron development in HoHe mice was due to a low level of expression of SOX1 protein in precursors, we used one hemisphere of the brain to assay OT development and the other for SOX1 antibody staining, linking in each animal the phenotype with the presence of SOX1 protein in precursors. We found no difference in the extent and level of expression of SOX1 protein in the VZ/SVZ of embryos with one copy of Sox1, whether it is expressed from the Sox2 locus in HoHe (see Figure 9E) or the endogenous Sox1 allele in Sox1βgeo/+ heterozygotes (see Figure 9D). Therefore, the emergence of VS/OT identity requires Sox1 expression in postmitotic cells. Consistent with the above findings, the small population of LGE postmitotic cells in HoHe mice that maintain SOX1 expression from the Sox2R allele migrate to the VS. However, the number of these neurons is small and cannot rescue the deficit in the area of the VS. In conclusion, although specification of neuronal identity is initiated in precursors, emergence of neuronal subtype and ventral migration require the continued presence of SOX1. Our findings suggest that in other brain areas, subtype identity and migration may also be controlled by the expression of transcription factors in postmitotic cells.
The current study, along with our previous one showing that SOX1 expression in the lens of the mouse is responsible for terminal differentiation and the expression of γ-crystallin genes [37], has revealed that SOX1 has important functions in postmitotic cell differentiation at two distinct sites. It is possible that the other SOXB1 factors have similar roles in postmitotic cells in which their expression is maintained.
Materials and Methods
Gene targeting.
The βgeo gene was inserted into the Sox1 single exon as previously described [37]. The resulting targeted locus produces a fusion protein consisting of the first 50 amino acids from SOX1 (which excludes the HMG box) followed by ten amino acids that are encoded by a synthetic linker sequence, and the βgeo sequence (including a polyadenylation signal). Tissue culture was carried out as described before [37], omitting the addition of gancyclovir for negative selection. The targeting vector did not contain any other selectable markers or promoters, and the targeting frequency was 1/52. As Sox1 is not normally expressed in embryonic stem (ES) cells, we used the minimum level of G418 for selection. Positive recombinants were identified by Southern blotting, using a 3′ 1-kb external probe on an EcoRI digest. Three ES cell clones were obtained, and one was successfully passed through to the germ line. All anatomical investigations were performed on mice of mixed genetic background. Sox1βgeo/+ mice were mated with mice that were heterozygous for the previously described [37] Sox1 deletion (Sox1M1) and did not express β-galactosidase.
For the Sox2 replacement vector, the 5′ and 3′ homologies used were the same as described before [30]. A SmaI-XhoI 2.2-kb Sox1 fragment containing the SOX1 open reading frame was flanked by LoxP cassettes followed by the NotI-SalI IRES–βgeo-polyA fragment (plasmid gift from Dr. A. Smith, University of Edinburgh). The replacement vector was linearized with SalI and electroporated into ES cells. Positive recombinants were identified by Southern blotting as described before [30]. Several targeted ES cells were isolated at a frequency of 1/20 and gave germ-line transmission of the mutation. Heterozygous animals carrying the Sox2Sox1βgeo allele (referred to in the text as Sox2R) expressed Sox1 and βgeo where Sox2 is normally expressed.
Deletion of the SOX1 coding region from the Sox2Sox1βgeo allele was achieved via pronuclear injection of a supercoiled plasmid expressing Cre-recombinase (gift from Dr. K. Rajewsky, Harvard Medical School). Although in the text we refer to this new allele as Sox2βgeo, it is officially named Sox2βgeo2 to distinguish it from the one previously described [30].
X-gal staining and in situ hybridization.
For β-galactosidase staining, fetal, newborn, or adult mouse brains were processed as previously described [30]. Detection of Sox1 mRNA was performed in whole embryos, as described previously [27,34]. The probes that were used on embryonic brain slices were generated by RT-PCR from embryonic brain cDNA. The position of the probes was Dlx1 (nt 41–573), Brn4 (nt 199–541), and Robo (nt 8–779). All fragments were cloned into a suitable cloning vector (pGET-easy, Promega, Madison, Wisconsin, United States), and were re-amplified using a sense oligonucleotide and an oligonucleotide upstream of either the T7 or SP6 sites. The resulting products were gel-purified, and 40 ng was used for probe synthesis. The brains were processed as described before [27,34].
BrdU labeling
A 25 mg/ml solution of BrdU (Sigma, St. Louis, Missouri, United States) was made in PBS warmed to 37 °C. The solution was sterilized through a 0.2-μm syringe filter and injected into the peritoneal cavity of pregnant mice (0.1 ml per 25 g of body weight, to give a final dosage of 0.1 mg/g). Brains were harvested 1 or 72 h after the injection, or on P16. The brains were fixed in 4% PFA in PBS at 4 °C overnight, embedded in paraffin wax, and cut into 5-μm thick sections. The sections were processed for immunohistochemistry as previously described [18].
Immunohistochemistry.
The source of antibodies and the dilutions used are as follows: PAX6, 1:10 (gift from Dr. J. Briscoe); GSH2, 1:5,000 (gift from Dr. K. Campbell); β-galactosidase 1:2,000 (Cappel); TuJ1, 1:1,000 (Novus Biologicals, Littleton, Colorado, United States); and SOX1, 1:500; SOX2, 1:500; and SOX3, 1:500 (gift from Dr. T. Edlund). MASH1, 1:2 (gift from Dr. F. Guillemot) [18], RC2 [56], and DARPP-32 [57] were used as previously described. For single or double immunofluorescence, embryonic tissue was fixed either for 1 h in 4% PFA in PBS at room temperature or for 15 min in MEMFA as described before [58]. The brains were then washed in PBS, cryoprotected overnight in 30% sucrose in PBS at 4 °C, embedded in OCT (Raymond Lamb), and cut in 10-μm and 15-μm sections using a cryostat. Sections were rehydrated in PBS, blocked in 4% goat serum and 0.1% Triton X-100 in PBS, and incubated overnight at 4 °C with primary antibodies diluted in the blocking solution. After incubation, the slides were washed in PBS and incubated with fluorescent secondary antibodies (FITC- or TRITC-labeled, 1:200 in blocking solution) for 1 h at room temperature.
Slides for double immunolabeling were first immunostained for SOX1 as described above and visualized with Alexa568 goat anti-rabbit antibody (1:500, Molecular Probes, Eugene, Oregon, United States), and then incubated with unlabeled anti-rabbit secondary antibody (1:100, Dako, Glostrup, Denmark) for 1 h at room temperature to block existing unlabeled anti-SOX1 antibody. Subsequently, the slides were incubated with anti-SOX2 antibody in blocking solution without Triton X-100 for 48 h at room temperature and visualized with Alexa488 goat anti-rabbit antibody (1:500, Molecular Probes). The cross-reactivity of the SOX1 and SOX2 antibodies when using immunohistochemistry was excluded by looking at tissues where SOX1 (lens; [37]) or SOX2 (see Figure 7B–7E) are uniquely present, and in Western blots where each antibody recognizes a different size band (data not shown). After incubation with the secondary antibodies, the slides were washed in PBS and the sections were mounted with Vectashield mounting medium (Vector Laboratories, Burlingame, California, United States) and observed under either a fluorescent or a confocal microscope.
Supporting Information
Figure S1
The GSH2/PAX6 Boundary Is Unaffected throughout Development in the Absence of SOX1
Similar to earlier stages (E12; see Figure 5), the expression of GSH2 (green) and PAX6 (red) protein in wild-type (A, C, and E) and mutant brains (B, D, and F) is the same at E14 and E15, as shown by DAPI (blue) nuclear stain. Cx, cortex; lge, lateral ganglionic eminence; mge, medial ganglionic eminence. Scale bar = 200 μm for (C) and (D).
(10 MB TIF).
Click here for additional data file.
Figure S2
Abnormal Distribution of VS Pax6-Expressing Neurons in the Absence of SOX1
Ventral telencephalic region of coronal brain sections stained with antibodies: PAX6 (brown) at E16 (A and B) and adult (C and D); SOX1 (green) and PAX6 (red) at E14 (E); and SOX1 and β-galactosidase (green) at E15 (F). Note Pax6-expressing cells are excluded from the OT region in the ventral telencephalon in wild-type (Sox1+/+) mice but not in the mutant(Sox1−/−). Arrows indicate ectopically localized Pax6-expressing cells. PAX6 and SOX1 are co-expressed in precursors but not in postmitotic cells in both mutant and wild-type. Scale bar = 500 μm for (C) and (D).
(10 MB TIF).
Click here for additional data file.
Figure S3
Sox1-Expressing VZ Precursors Have Radial Glial Morphology
Detail from immunofluorescence with β-galactosidase antibody of an E15 coronal brain section from mice carrying the Sox1βgeo allele. In the cortical ventricular zone, this staining reveals the cytoplasmic compartment of the SOX1-expressing progenitors, which have radial glial morphology.
(3.34 MB TIF).
Click here for additional data file.
Figure S4
The Distribution of Radial Glia in the LGE Is Unaffected in the Absence of SOX1
Coronal brain sections of wild-type (+/+) and Sox1-null (−/−) mice at E16. Immunostained with RC2 antibody (a radial glia marker) showing no differences.
(2.5 MB TIF).
Click here for additional data file.
Figure S5
Widespread Presence of SOX1 and SOX2 Proteins in Nuclei of the LGE VZ
Immunofluorescence (green) of E14-stage wild-type coronal brain sections at the level of the LGE stained with SOX1 (C and G) and SOX2 (D and H) antibodies and visualized under a confocal microscope. (A, B, and E–H) are stained with TOTO red-fluorescent nuclear stain. (C, E, and G) and (D, F, and H) show high magnification of the area indicated in the rectangle in (A) and (B), respectively.
(10 MB TIF).
Click here for additional data file.
Table S1
No Difference in the Number of LGE Dividing Precursors in the Absence of SOX1
BrdU-positive cells were counted using the Openlab image analysis program (Improvision, Coventry, United Kingdom). Measurements were performed in the area of the LGE (VZ/SVG) and of the pallial VZ. All data are represented as mean ± standard error of the mean. Cell counts were done in at least three different slides (sections) from each brain and in at least three separate optical fields in each slide (n = 4). To correct for tissue thickness and to obtain a better estimate of the proliferation within the LGE VZ/SVZ, the numbers of BrdU-positive cells were expressed as a ratio of the total number of LGE VZ/SVZ cells that were counted after hematoxylin staining, and as a ratio to the BrdU-positive cells of the pallial VZ/SVZ. Comparisons were made between wild-type and mutant mice using the unpaired Student's t-test (p < 0.05).
(21 KB DOC).
Click here for additional data file.
Accession Numbers
The GenBank (http://www.ncbi.nlm.nih.gov/Genebank/) accession numbers for the entities discussed in this paper are Brn4 (NM 008901), Dlx1 (NM 010053), Robo (MMU 17793), Sox1 (MN 009233), and Sox2 (MN 011443).
Acknowledgements
We thank for antibodies Drs. K. Campbell (Children's Hospital Research Foundation, Cincinnati, Ohio), T. Edlund (University of Umea, Sweden), and F. Guillemot (National Institute for Medical Research Medical Research [NIMR MRC], London). For comments on the manuscript we thank J. Briscoe (NIMR MRC) and J. Corbin (Georgetown University, Washington, DC). We are grateful to Z. Webster for the generation of transgenic mice and M. Delahaye for technical support with the mice. This work was supported by the MRC, the Wellcome Trust (grant 062197 to AC), and a Marie Curie Fellowship of the European Community Program (contract QLGA-CT-2001–50880 to AE).
Competing interests. The authors have declared that no competing interests exist.
Abbreviations
bHLH - basic helix-loop-helix
BrdU - 5-bromo-2′-deoxyuridine
CNS - central nervous system
DARPP-32 - dopamine and cAMP-regulated phosphoprotein
E[number] - embryonic day [number]
ES - embryonic stem
IRES - internal ribosomal entry site
LGE - lateral ganglionic eminence
OT - olfactory tubercle
P[number] - postnatal day [number]
SVZ - subventricular zone
TuJ1 - anti-βIII-tubulin antibody
VZ - ventricular zone
VS - ventral striatum
βgeo - β-galactosidase-neo
Figures and Tables
Figure 1
The Mouse Sox1βgeo Allele Reveals the Requirement of SOX1 in the Development of VS Neurons
(A) Strategy for targeting of the Sox1 locus by insertion of βgeo. Restriction enzymes: RV, EcoRV; K, KpnI; E, EcoRI; S, SpeI; B, BamHI. Yellow boxes indicate βgeo, green, SOX1 exon, and blue lines indicate fragments appearing in Southern blots of EcoR1-digested genomic DNA, hybridized with the external probe, which is shown with red lines.
(B–E) X-gal and SOX1 antibody staining of Sox1βgeo/+. Comparison of Sox1βgeo expression visualized by X-gal staining (B and D) and the endogenous wild-type Sox1 gene visualized by whole-mount in situ (C) and SOX1 antibody staining (E). (B and C) show E9-stage embryos and (D and E) show coronal sections of newborn ventral telencephalon.
(F–M) 100-μm coronal sections (Vibratome) were stained with X-gal to identify cells with Sox1 promoter activity. (F–I) show Sox1βgeo/+ forebrain sections from E13 to birth (P0) showing normal migration of Sox1-expressing cells from the VZ to the site of the OT, including striatal bridges. (J–M) show sections of Sox1βgeo/M1 forebrain, showing absence of X-gal staining in the OT and the striatal bridges. Red arrowheads show the anterior commissure.
Scale bar = 500 μm for (B) and (C) and 300 μm for (D–M).
Figure 2
Ectopic Distribution of Sox1-Null Neurons
X-gal staining of mouse forebrains at P16. (A and B) show intact forebrain viewed from the ventral surface, and (C–F) show 150-μm coronal Vibratome sections for Sox1βgeo/+ mice (A, C, and E) and Sox1βgeo/M1 mice (B, D, and E). Red arrows indicate the width of the OT. Red arrowheads indicate increased X-gal staining at more medial and posterior areas of the brain in (B), and in the striatum and septum in (D) and (F). White arrowheads indicate islands other than the medial islands of Calleja. an, accumbens nucleus; I, II, III, cell layers of the OT; ICjM, medial islands of Calleja; lot, lateral olfactory tract; lsn, lateral septal nucleus; ob, olfactory bulb; PC, olfactory (piriform) cortex; S, striatum; sb, striatal bridge Scale bar = 500 μm.
Figure 3
Normal Precursor Proliferation and Neurogenesis but Loss of OT Neuronal Differentiation in the Absence of SOX1
Coronal brain sections from the ventral telencephalon of wild-type (+/+) and Sox1-null (−/−) embryos. TuJ1 immunolabeling (A and B) at E13 shows no difference in early neuronal differentiation embryos; in situ hybridization at E16 for Brn4 (C and D) and Robo (E and F) shows absence of differentiation in the mutant at the prospective OT area. Red arrow in wild-type brain sections indicates OT. Telencephalic sections of wild-type (G, I, and K) and Sox1-null mutant (H, J, and L) embryonic brains were harvested 1 h after BrdU injection at E13 (G and H), E14 (I and J), and E15 (K and L) to detect actively dividing cells of the VZ/SVZ. Positive cells were visualized with anti-BrdU immunofluorescence (G–J) or with DAB staining (K and L). (K and L) show dorsal LGE area at high magnification. No differences were detected in the proliferation precursors at all stages examined, and no ectopic proliferation was observed in the mutant brains. Measurements and statistical analysis of BrdU-positive cells were performed on the DAB-stained sections, showing no significant differences (see Table S1). Scale bar = 300 μm (A and B), 300 μm (C–F), 500 μm (G–J), 500 μm (K and L).
Figure 4
Failure of Neurons to Migrate to the VS in the Absence of SOX1
This figure shows BrdU labeling of proliferating cells in the developing forebrain. Immunohistochemistry was performed on 5-μm coronal sections, cut at the level of the OT.
(A and B) Sections at E17, after BrdU injection at E14. White arrowheads in (A and B) indicate streams of migrating cells.
(C–H) Sections at P16, after BrdU injection at E13 (C and D), E14 (E and F), or E16 (G and H). The DAB reaction product (C–H) was viewed under dark-field illumination. “II” is layer II of the OT, and the red bracket indicates the olfactory cortex. Note E13-born neurons contribute laterally to the olfactory (piriform) cortex, and medially to the layer II of the OT and the striatal bridges (red arrow). E14-born neurons contribute to more medial VS structures than E15- and E16-born cells, which contribute almost exclusively to the medial islands of Calleja (red arrowheads).
Scale bar = 300 μm (A and B), 1 mm (C–H).
Figure 5
Normal Generation and Patterning of LGE Precursors in the Absence of SOX1
(A and B) Immunocytochemistry and on coronal brain sections of dorsal and ventral telencephalic markers in wild-type (+/+) and Sox1-null (−/−) embryos. PAX6 and GSH2 immunocytochemistry in the dorsal LGE at E12 shows no difference at the expression boundary in the absence of SOX1; the arrows point at the stream of PAX6-positive cells emanating from the boundary.
Double immunostaining for SOX1/PAX6 in wild-type brain (C), and for β-galactosidase/PAX6 (D) in the Sox1βgeo/− brain, at the VS area at E15. Note the presence of the PAX6-positive neurons in the area of the VS in the Sox1βgeo/− brain.
(E and F) MASH1 immunocytochemistry in the LGE of wild-type (E) and Sox1-null brain (F), at E13. No changes are detected.
(G and H) The distribution of Dlx1-expressing cells, as detected by in situ hybridization, is similar in both wild-type and mutant brains.
Scale bar = 300 μm (A and B), 200 μm (C–F), 150 μm (G and H).
Figure 6
SOX2 and SOX3 Down-Regulation in LGE Neurons and SOX1/SOX2 Co-Expression in LGE Precursors
Immunofluorescence of coronal sections at LGE levels in (A–C) E15- and (D–L) E13-stage wild-type embryos visualized on a confocal microscope: antibody staining for (A, D, G, and J) SOX1 (red), (B, E, H, and K) SOX2 (green), (C) SOX3 (green), (D–L) double SOX1 (red) and SOX2 (green), and (F, I, and L) merged. In the OT area and the LGE mantle, there are more neurons expressing SOX1 (A and J) than SOX2 (B and K) and SOX3 (C). Note the extensive co-expression of the SOX1 and SOX2 in precursors (D–I). (G–I) are higher magnifications of the areas within the rectangles. Scale bar = 300 μm.
Figure 7
The Sox2R Allele Delivers SOX1 in Sox2-Specific Expression Sites
(A) Strategy for targeted replacement of the SOX2 coding region with that of SOX1 and IRES-βgeo. Restriction enzymes: S, SalI; E, EcoRI; Sm, SmaI; X, Xho. Green boxes indicate Sox1; black arrowheads indicate LoxP sites; yellow boxes indicate IRES βgeo; blue lines indicate fragments appearing in Southern blots of EcoR1-digested genomic DNA, hybridized with the external probe, which is shown with red lines. Black arrows show the locus after recombination, homologous and Cre-mediated where is indicated.
(B–E) SOX1 immunostaining of frontal sections from E10 embryos. (B and D) Sox2+/+ and (C and E) Sox2R/+ showing the ectopic expression of SOX1 in the diencephalon (arrowheads) and the nasal pit (np) at E13.
Figure 8
The Distribution of VS Neurons Is Unaffected in Mice Over-Expressing SOX1 from the Sox2R Allele
(A–D), X-gal staining of coronal sections from the ventral telencephalon of P0 mice. Note that there is no difference in the distribution of Sox2-expressing OT neurons with SOX1 (A) or without SOX1 (C), and in Sox2R/+ mice with two wild-type Sox1 alleles (A) or one (B). Comparison of the number and distribution of neurons expressing Sox2βgeo in (C) and Sox1βgeo in (D) shows overlapping expression. (A–C) show 150-μm sections, and (D) shows a 80-μm section.
(E and F) DARPP-32 immunostaining of coronal sections from the ventral telencephalon of P10 Sox2R/+ and Sox1βgeo/+ single heterozygous mice, showing no difference in the generation and migration of OT neurons. AC, anterior commissure. Scale bar = 100 μm.
Figure 9
Sox1 Expression in Precursors Is Not Sufficient for the Emergence of OT/VS Neurons
(A–C) X-gal staining of coronal sections from the ventral telencephalon showing Sox1-βgeo-expressing OT-prospective neurons at E16-stage embryos with one wild-type Sox1 allele, Sox1βgeo/+, in (A), and none, Sox1βgeo/M1, in (B), and HoHe (Sox1βgeo/M1, Sox2R/+) in (C). Note the absence of X-gal-stained neurons in the area of the VS (red arrowheads), indicating failure of the Sox2R allele to rescue OT neuron development in the HoHe embryos. Expression of βgeo from the Sox2R allele in the HoHe (C) may account for the slight increase of X-gal staining compared to (B).
(D and E) SOX1 immunostaining at E16 embryos performed on the other halves of the brains of (A and C), respectively. Note that the level of SOX1 expression in the precursors (yellow arrows) is the same, whether it is expressed from the Sox2R allele in the HoHe (E) or from one of the Sox1 wild-type alleles in the Sox1 single heterozygotes (D). Note in the HoHe (E), this expression is not sufficient for the development of OT neurons.
(F and G) DARPP32 immunostaining of coronal brain sections from Sox1+/+ Sox2R/+ (F) and Sox1M1/M1 Sox2R/+ (G) P10 mice indicating loss of VS neurons.
Red arrowheads indicate OT; red arrows indicate anterior commissure. OC, olfactory cortex. Scale bar = 500 μm (A–E), 1 mm (F and G).
Figure 10
Sox1 Expression in Postmitotic LGE Cells Is Sufficient for Neuronal Migration in the VS
X-gal staining of coronal sections from the ventral telencephalon of P0 mice indicating the migration of neurons expressing SOX1 from the Sox2R allele in the presence (A) and absence (B; HoHe) of endogenous Sox1 wild-type genes. Black arrow points at the striatal bridges forming in HoHe mice. Scale bar = 100 μm.
Footnotes
Author contributions. AE, IK, SM, HW, and VE conceived and designed the experiments. AE, IK, SM, HW, PA, MD, and VE performed the experiments. AE, IK, SM, HW, and VE analyzed the data. AE, IK, SM, HW, DK, AC, RL, and VE contributed reagents/materials/analysis tools. VE wrote the paper.
¤a Current address: Stem Cell Biology Laboratory, Wolfson Centre for Age-Related Diseases, King's College, London, United Kingdom
¤b Current address: The Cyprus Institute of Neurology and Genetics, Nicosia, Cyprus
¤c Current address: Nature Reviews Neuroscience, London, United Kingdom
Citation: Ekonomou A, Kazanis I, Malas S, Wood H, Alifragis P, et al. (2005) Neuronal migration and ventral subtype identity in the telencephalon depend on SOX1. PLoS Biol 3(6): e186. | [
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16026622 | Quantitative trait analysis of the development of pulmonary tolerance to inhaled zinc oxide in mice
Abstract
Background
Individuals may develop tolerance to the induction of adverse pulmonary effects following repeated exposures to inhaled toxicants. Previously, we demonstrated that genetic background plays an important role in the development of pulmonary tolerance to inhaled zinc oxide (ZnO) in inbred mouse strains, as assessed by polymorphonuclear leukocytes (PMNs), macrophages, and total protein in bronchoalveolar lavage (BAL) phenotypes. The BALB/cByJ (CBy) and DBA/2J (D2) strains were identified as tolerant and non-tolerant, respectively. The present study was designed to identify candidate genes that control the development of pulmonary tolerance to inhaled ZnO.
Methods
Genome-wide linkage analyses were performed on a CByD2F2 mouse cohort phenotyped for BAL protein, PMNs, and macrophages following 5 consecutive days of exposure to 1.0 mg/m3 inhaled ZnO for 3 hours/day. A haplotype analysis was carried out to determine the contribution of each quantitative trait locus (QTL) and QTL combination to the overall BAL protein phenotype. Candidate genes were identified within each QTL interval using the positional candidate gene approach.
Results
A significant quantitative trait locus (QTL) on chromosome 1, as well as suggestive QTLs on chromosomes 4 and 5, for the BAL protein phenotype, was established. Suggestive QTLs for the BAL PMN and macrophage phenotypes were also identified on chromosomes 1 and 5, respectively. Analysis of specific haplotypes supports the combined effect of three QTLs in the overall protein phenotype. Toll-like receptor 5 (Tlr5) was identified as an interesting candidate gene within the significant QTL for BAL protein on chromosome 1. Wild-derived Tlr5-mutant MOLF/Ei mice were tolerant to BAL protein following repeated ZnO exposure.
Conclusion
Genetic background is an important influence in the acquisition of pulmonary tolerance to BAL protein, PMNs, and macrophages following ZnO exposure. Promising candidate genes exist within the identified QTL intervals that would be good targets for additional studies, including Tlr5. The implications of tolerance to health risks in humans are numerous, and this study furthers the understanding of gene-environment interactions that are likely to be important factors from person-to-person in regulating the development of pulmonary tolerance to inhaled toxicants.
Introduction
Individuals may develop tolerance to adverse health effects elicited from repeated exposure to inhaled toxicants in several different occupational and environmental situations. Pulmonary tolerance can be defined as the lung's ability to withstand the detrimental effects of a toxic compound following multiple exposures. There are several implications of tolerance to human health, having both advantages and disadvantages with respect to the development of harmful health effects. Clinical investigations of zinc oxide (ZnO)- [1], endotoxin- [2], and ozone- [3-6] induced adverse respiratory effects have demonstrated inter-individual variability in the capacity to develop pulmonary tolerance following inhalation exposure. Because these clinical studies are more tightly controlled than epidemiologic studies, they suggest that genetic background and gene-environment interactions contribute to the development of pulmonary tolerance in humans.
We initially characterized the pulmonary tolerance phenotype in an outbred mouse model by assessing levels of BAL protein and polymorphonuclear leukocytes (PMNs) following single (1X) and 5 daily repeated (5X) exposures to inhaled ZnO to begin the identification of the genes regulating the development of pulmonary tolerance to repeated toxicant exposure [7]. Significant genetic variability in the development of pulmonary tolerance to ZnO, endotoxin, and ozone was established in several inbred strains of mice in a subsequent study [8]. Of the strains tested, the BALB/cByJ (CBy) strain was tolerant and the DBA/2J (D2) strain was non-tolerant to BAL protein, PMNs, and macrophages following repeated ZnO exposure.
Because inbred mouse strains are virtually identical at all loci throughout their genome, and also share several chromosomal regions of conserved synteny with humans, they are an ideal animal model in which to investigate genotype-environment interactions and identify genes controlling pulmonary responses to inhaled toxicants where no a priori evidence for their location exists [9]. Inbred strains of mice have been successfully utilized in quantitative trait locus (QTL) analyses to identify candidate genes that control susceptibility to adverse pulmonary responses induced by a variety of inhaled gases [10-13] and particulates [14,15]. In the present study, a CByD2F2 mouse cohort was used to determine QTLs linked to the development of pulmonary tolerance to BAL protein, PMN, and macrophage phenotypes following repeated ZnO exposure.
Materials and methods
Mice
Inbred BALB/cByJ (CBy), DBA/2J (D2) and CByD2F1/J (F1) mice (6–7 weeks of age) were purchased from The Jackson Laboratory (Bar Harbor, ME). CByD2F1/J mice were crossed to produce F2 (intercross) offspring in our laboratory animal facility. All mice were acclimated for at least 1 week before exposure, housed in a positive pressure environment with a 12-hour light/dark cycle starting at 6:00 a.m., and provided with water and standard laboratory rodent chow (Purina, Indianapolis, IN) ad libitum except during exposure. All mice were handled in accordance with the standards established by the U.S. Animal Welfare Acts set forth in National Institutes of Health guidelines, and by the New York University School of Medicine Division of Laboratory Animal Resources.
Zinc oxide generation, characterization, and exposure
Mice were exposed to ZnO (1.0 ± 0.2 mg/m3, mean ± SD) in stainless steel cages placed inside a 0.07 m3 Plexiglas chamber. ZnO fumes were generated in a furnace as previously described [7,16]. The ZnO particles had a mass median aerodynamic diameter of 0.3 μm and geometric standard deviation of 1.5. Samples of the chamber atmosphere were collected approximately every 40 minutes from the manifold of the exposure system with polytetrafluoroethylene filters (Type TX40HI20-WW, Pallflex Products Corp., Putnam, CT), and the ZnO concentration was determined gravimetrically using a microbalance (Model C-30, Cahn Instruments, Cerritos, CA).
For linkage analyses, all F2 offspring (n = 299, 138 males and 161 females) were exposed at 7–12 weeks of age in a total of seven 5X exposure regimens. Development of pulmonary tolerance was assessed 24 hours after the fifth exposure, and exposure group sizes ranged from 36 to 45 animals. All ZnO exposures of F2 mice also contained at least two CBy, D2, and F1 mice for control purposes.
Bronchoalveolar lavage (BAL)
Mice were euthanized by intraperitoneal injections of ketamine HCl (100 mg/kg, Vetalar, Fort Dodge Laboratories, Inc., Fort Dodge, IA) and sodium pentobarbital (175 mg/kg, Sleepaway, Fort Dodge Laboratories, Inc.), and the posterior abdominal aorta was severed. The lungs of each mouse were lavaged two times with 1.2 ml of Dulbecco's phosphate buffered saline without Ca2+ or Mg2+ (pH 7.2–7.4, 37°C, Invitrogen, Carlsbad, CA). The collected BAL was immediately placed on ice (4°C) following recovery. Measurement of BAL protein, total cell counts, and differential cell counts were performed as previously described [8].
DNA isolation and genotyping
Genomic DNA was isolated from kidney tissue of each phenotyped F2 animal and for CBy, D2, and F1 controls (Wizard Genomic DNA Purification Kit, Promega, Madison, WI). DNA concentration was determined using a Beckman DU-650 spectrophotometer, and each sample was diluted to 10 ng/μl for genotype analysis. PCRs were performed to genotype F2 offspring for SSLPs located throughout the mouse genome. Eighty-six unlabeled primer pairs for SSLPs that differed in length by at least 5% between the CBy and D2 progenitor strains were purchased from Research Genetics (ResGen/Invitrogen). PCR was performed in 20 μl reaction volumes in 96-well low profile plates (Fisherbrand, Fisher Scientific, Fairlawn, NJ) using a PTC-100 thermal cycler (MJ Research, Watertown, MA). The final concentration for each reaction was: 10 mM Tris-HCl (pH 8.3), 50 mM KCL, 2.5 mM MgCl2, 0.2 mM of each deoxynucleotide triphosphate (Promega), 1.1X Rediload (ResGen/Invitrogen), and 0.132 μM of each SSLP primer pair. This reaction mixture was added to 100 ng of genomic DNA and 0.45 U of Taq DNA polymerase (Roche Applied Science, Indianapolis, IN). Final reaction mixtures were initially denatured at 94°C for 3 min, followed by 36 amplification cycles (94°C for 30 seconds, 57°C for 45 seconds, and 72°C for 30 seconds + 1 second/cycle). A final extension step at 72°C for 7 min was followed by refrigeration (4°C). PCR products were differentiated on 3% agarose (Invitrogen) gels and all samples were visualized by ethidium bromide staining using a ChemiImager-4400 low light imaging system (Imgen Technologies, Alexandria, VA) and called by a single investigator. Any questionable calls in reading the genotype from the image were reviewed by a second investigator and if not resolved, that sample was rerun.
Estimation of loci
The number of independently segregating loci was calculated using the following formula by Wright [17]: n = (P2 - F1)2/4(|σ2F2 - σ2F1|), where n is an estimate of the number of independent loci; P2 and F1 are the mean BAL protein responses following 5X ZnO exposure in CBy and CByD2F1 mice, respectively; σ2F2 and σ2F1 are the computed variances of the F2 and CByD2F1 mice, respectively.
Linkage analyses
A genome scan was performed to identify associations between genotypes and the BAL protein phenotype using a CByD2F2 mouse cohort. All phenotypic data were natural log normalized to generate a normal distribution to meet normality assumptions of the Map Manager QT computer program. Interval analyses were then performed by fitting a regression equation for the effect of a theoretical QTL at the position of each SSLP and at 1-centimorgan (cM) intervals between SSLPs using free, additive, recessive, and dominant regression models. The regressions and significance of each genotype/phenotype association (or likelihood χ2 statistic) were calculated by Map Manager [18]. Permutation tests were performed on the phenotypic and genotypic data using Map Manager to generate empirical thresholds for significance following the methods of Churchill and Doerge [19,20].
For the initial genome scan, the 15 most tolerant and 15 most non-tolerant F2 animals with respect to BAL protein, PMNs, and macrophages (i.e., the phenotypic extremes) were used for selective genotyping [9,21]. Interval analyses were done as stated previously, and 10,000 permutations were performed to generate significant and suggestive likelihood χ2 statistic thresholds for the BAL protein phenotype. Following the identification of a suggestive QTL for BAL protein on chromosome 1, the entire F2 cohort was examined for additional QTLs. For the BAL protein phenotype, three additional SSLPs on chromosome 1 were analyzed, and a permutation test (10,000 permutations) was performed with only chromosome 1. This method was similar to that used in previous linkage studies with inhaled particles and gases that utilized Map Manager QT [10,14,22]. All likelihood χ2 statistic thresholds corresponded to those reported in the aforementioned linkage studies.
Haplotype analysis
A haplotype analysis was carried out similar to that done previously by Prows and Leikauf to determine the contribution of each QTL and QTL combination to the overall BAL protein phenotype [15]. This method quantifies any difference in mean BAL protein levels that are linked with a particular haplotype. Haplotypes for the following SSLPs were used for this analysis: D1Mit291 (101.5 cM), D4Mit254 (82.5 cM), and D5Mit193 (1.0 cM). Mean BAL protein concentrations for groups of F2 mice with the same haplotype at each QTL or QTL combinations were calculated and compared with the mean BAL protein of F2 mice with the other haplotypes to determine the contributions of these QTLs to the overall BAL protein phenotype.
Results
Phenotypes of the CByD2F2 cohort
To further understand the role of genetic background in the development of pulmonary tolerance, an F2 (backcross) cohort derived from the CBy and D2 progenitors was phenotyped. The frequency distribution of the BAL protein, PMN, and macrophage phenotypic responses of the F2 cohort were within the ranges of similarly exposed CBy and D2 progenitor mice (Figure 1).
Figure 1
Frequency distribution of the number of BAL PMNs (×104), macrophages (×104), and protein (μg/ml) in BALB/cByJ, DBA/2J, CByD2F1/J, and CByD2F2 mice following 5 consecutive days of exposure to 1.0 mg/m3 ZnO for 3 h/day. Vertical dashed lines represent approximate separation points between BALB/cByJ and DBA/2J phenotypes.
Selective genotyping
A genome-wide scan was performed using the 15 most tolerant and 15 most non-tolerant mice to initially identify possible QTLs influencing the development of pulmonary tolerance. Permutation tests on the BAL protein-extreme data set established a suggestive likelihood χ2 statistic threshold of 9.9 and a significant likelihood χ2 statistic threshold of 17.4. These values were consistent with the genome-wide probabilities projected by Lander and Kruglyak [23]. Interval mapping identified a suggestive QTL for the BAL protein phenotype on the distal end of chromosome 1 (Figure 2). No QTLs were identified for the PMN and macrophage phenotypes from selective genotyping.
Figure 2
Genome-wide scan for QTLs associated with the BAL protein phenotype by selective genotyping of the CByD2F2 cohort. For each plot, the x-axis is the length of the chromosome in centimorgans (cM), and the y-axis is the likelihood χ2 statistic value as calculated by Map Manager. The upper and lower dashed lines represent significant (LRS = 17.4) and suggestive (LRS = 9.9) linkage thresholds, respectively.
Genotyping of the entire F2 cohort
The entire F2 cohort was genotyped with three additional SSLPs on distal chromosome 1 to further analyze the suggestive BAL protein QTL on chromosome 1 identified from selective genotyping. Interval mapping of the entire F2 cohort confirmed the QTL on chromosome 1 between 101.0 cM (D1Mit426) and 109.6 cM (D1Mit293) (Figure 3). The peak likelihood χ2 statistic value for this QTL exceeded the threshold value of 10.0 for significant linkage as determined by 10,000 permutations with all loci from chromosome 1 only.
Figure 3
Plot of a significant QTL on chromosome 1 that is associated with the BAL protein phenotype from analysis of the entire CByD2F2 cohort. The x-axis is the length of the chromosome in centimorgans (cM), and the y-axis is the likelihood χ2 statistic value as calculated by Map Manager. The lower dashed line represents the suggestive linkage threshold (LRS = 3.8), the middle dashed line represents significant linkage threshold (LRS = 10.0), and the upper dashed line represents the highly significant linkage threshold (LRS = 18.8).
The entire F2 cohort was further analyzed with the initial 86 SSLPs for the BAL protein, PMN, and macrophage phenotypes. Suggestive QTLs for the BAL protein phenotype that were not previously characterized by selective genotyping were identified on chromosome 4 between 53.6 cM (D4Mit146) and 82.5 cM (D4Mit254), and on chromosome 5 between 1.0 cM (D5Mit193) and 18.0 cM (D5Mit148) (Figure 4). Suggestive QTLs for the BAL PMN and macrophages were identified on chromosomes 1 and 5, respectively (Figure 5).
Figure 4
Plots of suggestive QTLs on chromosomes 4 and 5 associated with the BAL protein phenotype from analysis of the entire CByD2F2 cohort. The x-axis is the length of the chromosome in centimorgans (cM), and the y-axis is the likelihood χ2 statistic as calculated by Map Manager. The upper and lower dashed lines in each plot represent significant (LRS = 15.8) and suggestive (LRS = 9.2) linkage thresholds, respectively.
Figure 5
Plots of suggestive QTLs on chromosomes 1 and 5 associated with the BAL PMN and macrophage phenotypes, respectively, from analysis of the entire CByD2F2 cohort. The x-axis is the length of the chromosome in centimorgans (cM), and the y-axis is the likelihood χ2 statistic as calculated by Map Manager. The upper and lower dashed lines in each plot represent significant (LRS = 15.6) and suggestive (LRS = 9.2) linkage thresholds, respectively.
Haplotype analysis
Mean BAL protein levels of F2 mice with the same haplotype were calculated and compared with the mean BAL protein levels of F2 mice with the opposite haplotype in order to determine the contribution of each QTL and combinations of QTLs to the overall BAL protein phenotype (Figure 6). For each SSLP, F2 animals were genotyped as a homozygous CBy (CC), a homozygous D2 (DD) or heterozygous (H). For any individual SSLP, the greatest difference in mean BAL protein was found for D1Mit291. F2 mice that were DD at that locus had an average of 156 μg/ml more BAL protein than those mice that had CC or H haplotypes.
Figure 6
Differences in mean BAL protein levels of CByD2F2 mice with tolerant versus non-tolerant haplotypes at SSLPs representing the identified QTLs. Open bars, F2 mice with CC or CD genotypes (represented as H). Filled bars, F2 mice with a DD genotype (represented as D). Number within each bar is the number of F2 mice with the given genotype or haplotype. Values are means ± SE. All comparisons of tolerant (open bars) haplotypes versus non-tolerant haplotypes (filled bars) were significant (P < 0.05, t test).
The combinatorial effect of the QTLs on chromosomes 1, 4, and 5 were also examined. For any combination of two QTLs, F2 mice that had a DD-DD haplotype for markers on chromosomes 1 and 5 (D1Mit291 and D5Mit193) had an average of 310 μg/ml more BAL protein than those that were CC or H (CC/H) for those markers. The greatest difference in mean BAL protein levels were found in F2 mice that had a DD-CC/H-DD haplotype for the three QTLs on chromosomes 1, 4, and 5, (i.e., D1Mit291, D4Mit254, and D5Mit193), respectively. These mice had an average of 345 μg/ml more BAL protein than those that were CC/H at the markers across the three chromosomes.
Identification of candidate genes
Within all of the QTLs, candidate genes discovered with potential roles in controlling the development of tolerance to inhaled ZnO are presented in Tables 1 and 2. These genes were chosen as candidates from a thorough review of the existing literature and through the positional candidate gene approach, which combines knowledge of map position with the mouse transcript map [24].
Table 1
Positional candidate genes from linkage analysis of pulmonary tolerance to BAL protein following repeated ZnO exposure.
Table 2
Positional candidate genes from linkage analysis of pulmonary tolerance to BAL PMNs and macrophages following repeated ZnO exposure.
Development of pulmonary tolerance in MOLF/Ei mice
We identified toll-like receptor 5 (Tlr5) as an interesting candidate gene within the significant QTL for BAL protein on chromosome 1. A wild-derived inbred mouse strain called MOLF/Ei (M. m. molossinus) which has non-conservative mutations in Tlr5 [25] was phenotyped for BAL protein following 1X and 5X ZnO exposure to determine whether Tlr5 may function in the development of pulmonary tolerance (Figure 7). MOLF/Ei BAL protein was significantly increased above control values following 1X ZnO exposure (395 ± 38 μg/ml). However, MOLF/Ei mice exhibited a tolerant phenotype, as 5X BAL protein values (215 ± 22 μg/ml) were significantly decreased below that of the 1X exposure group.
Figure 7
BAL protein levels in wild-derived MOLF/Ei mice 24 h after single (1X) or repeated (5X) exposure to 1.0 mg/m3 ZnO or air for 3 h. Protein levels of BALB/cByJ and DBA/2J mouse strains following ZnO exposure are also shown for comparison purposes. Values are means ± SE (n = 4–5 MOLF/Ei mice/exposure group). * indicates significantly different from air-exposed MOLF/Ei controls, P < 0.05 [Student-Newman Keuls (SNK) test]. + indicates significantly different from 1X MOLF/Ei exposure group, P < 0.05 (SNK test).
Discussion
Clinical studies on the acquisition of tolerance to inhaled toxicants suggest that genetic background and gene-environment interactions contribute to the development of pulmonary tolerance in humans. We have previously determined that a genetic component exists in a mouse model of pulmonary tolerance to repeated ZnO exposure [8]. In the present study, we performed linkage analyses on an F2 mouse population derived from tolerant CBy and non-tolerant D2 strains to further ascertain the contribution of genetic background to the development of pulmonary tolerance, and identify candidate genes that may be important regulators in the acquisition of tolerance.
Initial analysis using the most tolerant and non-tolerant F2 mice with respect to BAL protein generated a putative QTL (designated as the zinc-induced tolerance (ZIT1) locus) on the distal end of chromosome 1. Further assessment of the entire F2 cohort demonstrated that the QTL on chromosome 1 attained an LRS value for significant linkage, and also identified two suggestive QTLs located on chromosomes 4 and 5. Using a variation of the Wright equation [17], a minimum of three loci were estimated to be independently segregating with the BAL protein phenotype following 5X ZnO exposure, thus in agreement with the results of the linkage analysis. There are several approaches that can be pursued to focus in on the QTL intervals that were identified. For instance, increasing the number of F2 mice used in the QTL analysis is one alternative. The major disadvantage of this, however, is that segregating QTLs contribute a great deal of phenotypic "noise," making it problematic when determining whether or not a given mouse has inherited a particular QTL [9]. Thus, in order to separate the effects of multiple loci, congenic mouse strains for each QTL could be generated, which could then be used to breed multicongenic lines to examine the existence of any epistatic effects. Additionally, future studies could employ the use of a backcross (CByD2F1 × CBy) mouse population to expand the evaluation of the QTL effects to the overall BAL protein phenotype.
To measure the contribution of each individual QTL and each QTL combination to the overall BAL protein phenotype, the protein levels for F2 mice with each particular haplotype were compared. Mice with opposite allelic combinations for QTLs on chromosomes 1 and 5 had a difference of 310 μg/ml BAL protein, which accounts for approximately one-third of the total difference in mean BAL protein between the parental CBy and D2 strains. Additionally, the mean BAL protein level of F2 mice with a DD haplotype for QTLs on chromosomes 1, 4, and 5 was over half that of the non-tolerant D2 parental strain. These analyses suggest that although three QTLs were identified, the development of pulmonary tolerance to BAL protein is a decidedly complex phenotype that is regulated by a number of different genes, some of which were likely not identified by linkage analysis using an F2 cohort.
Candidate genes within the significant QTL on chromosome 1 (ZIT1) that could play a role in controlling the development of tolerance to BAL protein following repeated ZnO exposure are presented in Table 1. The Duffy blood group (Dfy) has been shown to modulate the intensity of inflammation following endotoxin exposure [26], and has a role in enhancing inflammatory cell recruitment to sites of inflammation by facilitating movement of chemokines across the endothelium [27]. ADP-ribosyltransferase 1 (Adprt1) and tumor necrosis factor (TNF) receptor-associated factor (Traf5) are functionally associated with nuclear factor (NF)-κB, a key transcription factor in the regulation of the inflammatory process [28,29]. Additionally, activation of ADPRT1 plays a role in endotoxin-induced BAL protein increases [30,31]. Activated transforming growth factor-β has been shown to be a mediator of bleomycin- and endotoxin-induced lung permeability (i.e., BAL protein) in mice [32]. Finally, solute carrier family 30 (zinc transporter), member 1 (Slc30a1) is a metal transporter on the plasma membrane that confers resistance to zinc and cadmium toxicity in vitro via an efflux mechanism [33,34].
Candidate genes for BAL protein tolerance identified within the suggestive QTL intervals on chromosomes 4 and 5 are also presented in Table 1. Notable candidates include platelet-activating factor receptor (Ptafr) and phospholipase A2 (Pla2) that have been implicated as potential mediators of toxicant-induced BAL protein increases [35-37]. Interleukin (IL)-6 protein was increased following ZnO exposure in humans [1,38], and was hypothesized to be an anti-inflammatory suppressor of ZnO-induced lung injury. Interestingly, increased lung IL-6 levels have been shown to mediate pulmonary tolerance to ozone in rats [39]. Lastly, solute carrier family 30 (zinc transporter) members 2 and 3 (Slc30a2 and Slc30a3) have been identified as zinc transporters that protect against zinc-induced toxicity in both cell culture and animal models [40,41]. The development of pulmonary tolerance to BAL protein in our model could be regulated by any number of the aforementioned candidate genes. These candidates will be investigated in future studies of transcriptional and protein regulation to determine their roles in the development of tolerance.
The suggestive QTLs for tolerance to BAL PMNs and macrophages (on chromosomes 1 and 5, respectively) were also examined for positional candidate genes (Table 2). Chemokine (C-C motif) ligand 20 (Ccl20), chemokine (C-X-C motif) receptor 4 (Cxcr4), and interleukin-10 (Il10) were identified as candidates for BAL PMN tolerance. The movement of PMNs into inflammatory tissues is regulated by chemotactic factors (e.g., chemokines) that signal through numerous chemokine receptors [42]. PMNs are capable of producing several chemokines and proinflammatory cytokines, indicating that PMNs may be important in auto-direction of cell trafficking during inflammation. CXCR4 expression has been detected on human PMNs [43], and coordinated chemokine receptor gene expression may control the tissue-specific migration and activation status of PMNs into the lung. Human PMNs are also able to express CCL20 [44], which is able to recruit immature dendritic cells that play an important role in the initiation of the immune response as well as chronic inflammation [45-47]. Finally, IL-10 can be produced by T cells and is able to diminish PMN influx by inhibition of expression of proinflammatory chemokines [48], NF-κB (via I kappa kinase) [49], and TNF [50], as well as modulate cells and effector functions associated with an allergic response. With respect to the BAL macrophage phenotype, the epidermal growth factor receptor (EGFR) ligands amphiregulin (Areg) and betacellulin (Btc) were identified as candidate genes within the suggestive QTL on chromosome 5. Ligand-dependent activation of the EGFR by particles rich in metal content lead to activation of the MAP kinase signaling cascade and cytokine expression and secretion [51,52]. Secreted phosphoprotein 1 (Spp1, also known as osteopontin) was also identified, and it can act as a chemoattractant for macrophages [53,54]. Lastly, a host of chemokine (C-X-C motif) ligands were identified between 51 and 53 cM that could be involved in macrophage chemotaxis [55]. Again, tolerance to BAL PMNs and macrophages may be under the influence of any number of these candidate genes.
In the present study, we identified toll-like receptor (Tlr5) as a candidate gene within the significant ZIT1 QTL on chromosome 1 for tolerance to BAL protein. Toll-like receptors activate intra-cellular signaling that culminates in the induction of a multitude of effector genes [56]. Tlr5 has been shown to be an important gene in the immune response to Gram-positive and Gram-negative bacterial flagellin [57,58]. We utilized a wild-derived inbred mouse strain called MOLF/Ei which has non-conservative mutations in Tlr5 that are associated with a lower level of expression [25] to determine whether Tlr5 plays a role in the development of tolerance to BAL protein in our ZnO model. Because MOLF/Ei mice have a lower level of TLR5 mRNA expression compared to other strains [25], it was unclear whether we would observe tolerance after a single exposure to ZnO. Although the MOLF/Ei strain has no "wild-type" control strain per se, MOLF/Ei mice were tolerant to increased BAL protein following repeated ZnO exposure when compared to the non-tolerant D2 strain. These data support a role for Tlr5 in the development of tolerance to BAL protein. Interestingly, Kleeberger and colleagues identified Tlr4 as a candidate gene in a study of susceptibility to increased BAL protein in mice following a single exposure to 0.3 ppm ozone for 72 hours [22], which also suggests toll-like receptor signaling may be important in the regulation of inhaled toxicant-induced changes in BAL protein.
The mechanism through which Tlr5 signaling may regulate the development of tolerance to ZnO is unknown. While endotoxin [59] and bacterial flagellin [60] have been demonstrated as the primary ligands for Tlr4 and Tlr5, respectively, the ligand(s) that is responsible for toll-like receptor signaling following exposure to inhaled toxicants such as ozone and ZnO is unknown. Although there have been no studies on endogenous ligands for Tlr5, several endogenous ligands for Tlr4 have been identified. For example, fibronectin [61] and hyaluronic acid [62] are produced by lung cells during lung injury and are endogenous Tlr4 ligands. The downstream pathway through which Tlr5 may regulate the development of tolerance to ZnO-induced BAL protein is potentially via NF-κB, a transcription factor that is known to induce several cytokines involved in inhaled ZnO responsiveness such as IL-8 and IL-6 [1,63]. Furthermore, NF-κB-dependent gene expression is decreased in Tlr5-mediated tolerance to flagellin in vitro [64]. Thus, it is plausible that a mutation in Tlr5, a regulatory element upstream of NF-κB signaling, could modulate tolerance to BAL protein from repeated ZnO exposure. Finally, Tlr5 may function as a danger signal receptor in the development of ZnO tolerance, consistent with the "danger model" of innate immunity that explicates activation of the innate immune system by factors other than foreign antigens [65]. Whatever the case, much work is needed in understanding how toxicants function through toll-like receptor signaling mechanisms in the lung to regulate adverse responses such as BAL protein.
In summary, linkage analysis of a large F2 mouse cohort identified significant linkage of a QTL (ZIT1) on chromosome 1 associated with tolerance to BAL protein following repeated exposure to inhaled ZnO. Suggestive QTLs were also identified on chromosomes 4 and 5 for BAL protein, on chromosome 1 for BAL PMNs, and on chromosome 5 for BAL macrophages. Haplotype analysis suggested that the combinatorial effects of these three loci contributed to the overall phenotype, which agrees with the calculated number of segregating loci. Tlr5 was identified within the significant QTL for BAL protein on chromosome 1. Wild-derived Tlr5-mutant MOLF/Ei mice were determined to be tolerant to BAL protein following repeated ZnO exposure, suggesting a role for Tlr5 in the development of pulmonary tolerance to inhaled toxicants.
Conclusion
These data substantiate genetic background as an important influence in the acquisition of pulmonary tolerance following exposure to inhaled toxicants such as ZnO, and promising candidate genes exist within the identified QTL intervals that would be good targets for additional studies on the pathogenesis of tolerance.
Competing interests
The author(s) declare that they have no competing interests.
Authors' contributions
SCW: Participated in the design and coordination of the study, performed the study, and drafted the manuscript.
LCC: Participated in the design of the study and helped draft the manuscript.
TG: Conceived the study, participated in the design and coordination of the study, and helped draft the manuscript.
Acknowledgements
The authors would like to thank the following people for their contributions to this study: Karen Galdanes, Margaret Krasinski, Kathy Baker, and Dr. Moon-shong Tang (New York University); Dr. George D. Leikauf (University of Cincinnati); Dr. Daniel R. Prows (Cincinnati Children's Hospital); Dr. Steven R. Kleeberger (National Institute of Environmental Health Sciences); Dr. Carrie L. Welch (Columbia University). This study was supported by USEPA Grant R-826244, USEPA Particulate Matter Center Grant R-827351, USEPA STAR Fellowship U-91578301, NIEHS Center Grant of Excellence ES-00260, and CDC/NIOSH (via Mt. Sinai School of Medicine, New York, NY) Grant T-42CCT210425-06-01. | [
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16027110 | Identification and characterization of the mouse nuclear export factor (Nxf) family members
Abstract
TAP/hNXF1 is a key factor that mediates general cellular mRNA export from the nucleus, and its orthologs are structurally and functionally conserved from yeast to humans. Metazoans encode additional proteins that share homology and domain organization with TAP/hNXF1, suggesting their participation in mRNA metabolism; however, the precise role(s) of these proteins is not well understood. Here, we found that the human mRNA export factor hNXF2 is specifically expressed in the brain, suggesting a brain-specific role in mRNA metabolism. To address the roles of additional NXF factors, we have identified and characterized the two Nxf genes, Nxf2 and Nxf7, which together with the TAP/hNXF1's ortholog Nxf1 comprise the murine Nxf family. Both mNXF2 and mNXF7 have a domain structure typical of the NXF family. We found that mNXF2 protein is expressed during mouse brain development. Similar to TAP/hNXF1, the mNXF2 protein is found in the nucleus, the nuclear envelope and cytoplasm, and is an active mRNA export receptor. In contrast, mNXF7 localizes exclusively to cytoplasmic granules and, despite its overall conserved sequence, lacks mRNA export activity. We concluded that mNXF2 is an active mRNA export receptor similar to the prototype TAP/hNXF1, whereas mNXF7 may have a more specialized role in the cytoplasm.
INTRODUCTION
TAP/hNXF1 is the key factor mediating the nuclear export of mRNAs (1–3), and its orthologs in Saccharomyces cerevisiae, Drosophila melanogaster and Caenorhabditis elegans were shown to be essential for general cellular mRNA export (4–6). Metazoans encode additional NXF-like proteins, which together with the TAP/hNXF1 orthologs comprise a family of proteins termed nuclear export factors (NXFs) that are evolutionarily conserved from yeast to humans. Besides sequence homology, the NXFs share the domain architecture, and therefore are thought, by analogy to TAP/hNXF1, to participate in mRNA metabolism (7). For the human hNXF2 and hNXF3 proteins, the mRNA export activity has been confirmed by mRNA export assays (6,8). In Drosophila, while the NXF1 ortholog dmNXF1 is essential (9), additional NXF proteins were shown to be nonessential for general mRNA export in cell culture, suggesting roles that are more specialized than that of dmNXF1 (10). We previously found that the C.elegans NXF1 homolog Ce-NXF1 is essential for mRNA export (4). Although the RNAi depletion of Ce-NXF2 was not lethal (5), this protein was recently implicated in the post-transcriptional regulation of tra-2 mRNA, which is required for female development (11). A human hNXF5 nullisomy was linked to mental retardation (12). Taken together, these observations suggest that while the TAP/hNXF1 orthologs are essential for general mRNA export in metazoan species, additional NXF family members have more specialized roles. To understand these roles, we studied the family of the mouse NXF proteins. Here, we describe the isolation and characterization of two additional mouse Nxf genes, Nxf2 and Nxf7.
MATERIALS AND METHODS
Isolation of cDNAs encoding mNXF2
Gene-specific primers were designed using a short region of identity between the mouse BAC clone RP23 65A22 and human TAPX2/hNXF2 cDNA (13). 5′ and 3′ rapid amplification of cDNA ends (RACEs) were performed on mouse brain Marathon-ready cDNA (Clontech) using outward primers complementary to this identity region. The RACE products were cloned into TOPO vector (Invitrogen), and the cDNAs were sequenced by BigDye™ Terminator Sequencing Kit (AB Applied Biosystems). The mouse Nxf2 and Nxf7 genes were identified and sequenced from the mouse BAC clones RP23 65A22 and BAC441N13, respectively.
Recombinant DNA
Expression plasmids for the green fluorescent protein (GFP)-tagged mNXF2 or mNXF7 proteins or their deletion mutants were generated by PCR amplification of corresponding cDNAs, and subsequent insertion of PCR products into the SacII and NheI sites in plasmid pCMV-GFPsg25 (14) in-frame with GFP. GFP-βGal-tagged mNXF2 and its mutants were generated by PCR amplification, followed by insertion of SacII- and XbaI-digested PCR fragment into SacII and NheI sites in plasmid pGFP-βgal (13), respectively. Histidine-tagged mNXF2 was constructed by PCR amplification of full-length mNXF2 coding sequence, then cloned into BssHII and XhoI sites in pCMV37M1-10D (15) replacing gag. Single or multiple point mutations, using 40–80Gβgal as a template, were generated by the QuickChange XL Site-Directed Mutagenesis kit (Stratagene) according to the manufacturer's protocol. The nucleotide sequences of each mutant plasmid were confirmed by automated BigDye fluorescence sequencing. To generate glutathione S-transferase (GST)-tagged mNXF2, the coding region for amino acids 1–400 of mNXF2 was PCR amplified and subcloned into the pGEX-2T vector (Promega) for expression in Escherichia coli. Mouse p15/NXT1 expression plasmid was generated based on GenBank accession no. NM_019761, and the human p15-1 expression plasmid was obtained from E. Izaurralde.
Cell culture, transfection and microscopy
Human HeLa and 293 cells and mouse NIH3T3 and PA317 cells were transfected with FuGene 6 reagent (Roche) according to the manufacturer's protocol. All transfections for subcellular localization analysis were performed in 35 mm glass-bottom plates. Approximately 24 h post-transfection cells were fixed with 3.7% formaldehyde in phosphate-buffered saline. For some experiments, 20 h post-transfection, fresh media containing 30 nM leptomycin B, actinomycin D (2 μg/ml) or 5,6-dichlororibofuranosylbenzimidazole (DRB, 30 μg/ml) were added, and the cells were incubated for 4 h prior to fixation. As control for the drug-induced relocalization experiments, the same treatments were performed on cells expressing GFP-tagged HIV Rev protein. Microscopic analysis of GFP fluorescence was performed as described previously (13). Three-dimensional reconstruction of fluorescent images from confocal Z-stacks was performed using the maximum intensity projection renderer implemented in Imaris software (Bitplane).
Protein analysis
Chloramphenicol acetyltransferase (CAT) assays and luciferase measurements were performed as described previously (13). For western blot analyses, transfected cells were extracted in 500 μl of lysis buffer (0.5% Triton X-100 and 100 mM Tris–HCl, pH 7.4), separated on denaturing polyacrylamide gels and blotted onto nitrocellulose membrane. Polyclonal antisera against human TAPX2/hNXF2 and mouse mNXF2 were raised in rabbits using purified cellulose-binding domain (CBD) fusion protein containing amino acids 102–372 of TAPX2, or GST fusion containing full-length mNXF2 as immunogens. The TAPX2/hNXF2 antibodies were affinity purified on immobilized CBD-TAPX2 immunogen. For western blots, after probing with rabbit anti-TAPX2/hNXF2 or anti-mNXF2 antibody (1:1000) in 5% nonfat dry milk, and subsequent incubation with donkey anti-rabbit horseradish peroxidase-conjugated secondary antibody, immunoreactive proteins were visualized by enhanced chemiluminescence (ECL plus Western Blotting Detection System, Amersham) and autography. Pre-made western blots of multiple human tissues (GenoTech) and ‘mouse brain aging’ blots (RNAWAY) were quality controlled by the manufacturers to ensure equal loading and transfer efficiency.
In vitro protein binding assays
Reticulocyte-produced proteins were synthesized and metabolically labeled in coupled transcription/translation system (TNT T7 Coupled Reticulocyte Lysate System, Promega), using T7 promoter-containing PCR fragments as templates, and were adjusted with unprogrammed extract to equal molar concentrations. These stocks were used in the binding reactions that contained equimolar amounts of reticulocyte-produced proteins and 1–2 μg of E.coli-produced GST-tagged proteins that were immobilized on glutathione–Sepharose beads (Amersham). The binding was performed in 200 μl RBB buffer (15 mM HEPES, pH 7.9, 50 mM KCl, 0.1 mM EDTA and 0.2% Triton X-100) supplemented with 200 mM NaCl. Following incubation for 15 min at room temperature, the beads were pelleted and washed three times with binding buffer. Bound proteins were eluted by boiling in SDS–PAGE sample buffer, separated by SDS–PAGE and detected using Phosphoimager.
Biocomputing
Database similarity searches, multiple sequence alignments and phylogenetic analyses were performed using the standard programs of Genetics Computer Group package, with default parameters.
Nucleotide sequence accession
The sequences of mouse Nxf genes and cDNAs were submitted previously to GenBank under the accession numbers AY017476, AF490577 for Nxf2, and AY260550, AY266683 for Nxf7.
RESULTS
Identification of mouse NXF-related genes
As a first step to study the role of the NXF family of proteins, we have identified the complete cDNAs as well as the exonic structures of the mouse Nxf2 (GenBank accession numbers AY017476 and AF490577) and Nxf7 genes (GenBank accession numbers AY260550 and AY266683). To achieve the identification of the mouse homologs of human TAP/hNXF1-related proteins, database searches for NXF-related mouse expressed sequence tags (ESTs) or cDNAs were employed, which revealed a homology in the mouse BAC clone RP23 65A22 to the human TAPX2/hNXF2 cDNA (13), a hNXF2 isoform. A full-length cDNA clone, termed Nxf2, was obtained using mouse brain cDNA library as template and the intron sequences were identified from the BAC clone. By comparison of the cDNA and genomic sequences, the exon and intron structure of the mouse Nxf2 gene was determined (Figure 1A). We found that the mouse Nxf2 gene is >17 kb in length and consists of 21 coding exons with 20 in-frame AUGs present in 11 exons. Comparison of our mNXF2 protein sequence with those published by Wang et al. (16) (GenBank accession no. NM_031259) and Jun et al. (12) (GenBank accession no. NP_112549) shows 99% identity with a single E347D amino acid change. Database searches further revealed a partial cDNA sequence of an additional Nxf-related gene on another BAC clone. By PCR amplification based on EST homology, a full-length cDNA corresponding to mouse Nxf7 was obtained and its exonic structure was determined (Figure 1A). We found that the mouse Nxf7 gene contains 22 exons, spans >14 kb (GenBank accession no. AY266683) and encodes a predicted protein of 620 amino acids. Our cDNA clone has a perfect sequence match with all the exonic sequences in the mouse Nxf7 genomic DNA. The proteins encoded by Nxf-a1 (GenBank accession no. AJ305317) and Nxf-a2 (GenBank accession no. AJ305318) have a high sequence homology to mNXF7, although both protein isoforms lack 110 amino acids at the N-terminus as compared with mNXF7. Comparison of the mNXF7 protein with NXF-a1 and NXF-a2 reveals the following changes: L354P and L361P (NXF-a2); the replacement of 121-STF-123 with two valine residues (NXF-a1 and NXF-a2); NXF-a1 is a splice variant (lacking exon 10), which results in an internal deletion of 36 amino acids (nt 271–306) and removes the sequence between L271 and L306.
Taken together, we identified and isolated two genes termed mouse Nxf-2 and Nxf-7, whose sequences we have previously submitted to GenBank. Both genes are located on X chromosome, suggesting that they arose from a gene amplification, whereas Nxf1 is located on chromosome 19. Thus, the mouse NXF family consists of three members, whereas the human NXF family comprises five (7). Figure 1B shows a dendrogram of NXF family proteins from yeast to humans, illustrating significant homology both across and within species. On this tree, the mouse mNXF1 clusters with the other TAP/hNXF1 orthologs, from birds to humans, as expected (7). Interestingly, the ‘additional’ mouse factors mNXF2 and mNXF7 are close together and further cluster with the ‘additional’ human factors. We then performed a more detailed comparison of the mouse NXF family with TAP/hNXF1 protein, a prototype NXF factor (Figure 1C). While the mouse mNXF1 and the human TAP/hNXF1 share 90% identity, the comparison of mNXF1 to mNXF2 and mNXF7 shows reduced homologies of 47 and 49% amino acid identity, respectively. Despite this, the mNXF2 and mNXF7 proteins are predicted to share the domain organization with the human TAP/hNXF1 protein (7), including a non-canonical RNP-type RNA-binding domain (RBD), the leucine-rich repeats (LRRs), the NTF2-like domain (mediating interactions with p15/NXT1) and the ubiquitin associated-like domain (UBA-like) that is part of nucleoporin-binding region. mNXF2 also has an insertion of 5 tandem 12 amino acid imperfect repeats located at the C-terminus of its LRR domain, as previously noted (12), but its role was not further investigated.
Expression of mNXF2 protein in the brain
To study the mNXF2 protein expression, we generated an antiserum specific against GST-tagged mNXF2 in rabbits. Western blot analysis showed that the antiserum recognized untagged, His-tagged, GFP-tagged or GFP-βGal-tagged mNXF2 protein (Figure 2A, left panel), whereas it did not react with proteins from untransfected human 293 cells. In addition, it did not cross-react with the human TAP/hNXF1 or mouse mNXF7 proteins (Figure 2A, right panel). The untagged mNXF2 protein migrated with an apparent molecular mass of ∼75 kDa.
It has been previously reported that the mouse NXF-related mRNAs can be detected using RT–PCR in the brain (12). Using western immunoblot analysis, we found that the mNXF2 protein can also be detected in the mouse brain at different stages of development (Figure 2B). While the mNXF2 protein could be detected in a brain sample from an 18-week mouse embryo (E18), its level of expression is slightly increased at 1 week after birth and kept constant for ∼3 months and is still detectable, although at lower levels, after the age of 6 months. We were unable to address the expression of mNXF7 protein because of the lack of specific antibodies.
Similarly, we used a monospecific rabbit antiserum generated to human TAPX2 that represents an isoform of hNXF2 (13), to probe a human multiple tissue blot. This analysis revealed that TAPX2/hNXF2 was specifically expressed in the brain (Figure 2C), but could not be detected in other tissues examined, such as testis, where mRNA was readily detectable (data not shown). Taken together, these results confirmed that NXF-related proteins, such as the human and mouse NXF2, are expressed in the brain, suggesting the roles in brain-specific mRNA metabolism.
Distinct subcellular localization of mNXF2 and mNXF7
We next studied the subcellular localization of mNXF2 and mNXF7 upon transfection of human HeLa cells with plasmids expressing GFP-tagged fusion proteins. Figure 3 shows that mNXF2 localizes to the nucleus, but is excluded from the nucleolus, accumulates at the nuclear rim and is present in the cytoplasm. Its localization is similar to that observed for the human TAP/hNXF1 (13,17,18). In contrast, mNXF7 is localized exclusively to the cytoplasm (Figure 3), where it accumulates in granules. Similar localization patterns of mNXF2 and mNXF7 were found in human 293 cells as well as in mouse NIH3T3 and PA317 cells (data not shown); thus, the distinct subcellular localization is independent of the tested cell lines. The localization of both proteins was not affected in the presence of Leptomycin B, which excludes a role of CRM1 in protein export. Also, the presence of Actinomycin D or DRB did not alter the localization of mNXF2 and mNXF7 (data not shown), indicating that their nucleocytoplasmic trafficking is transcription-independent, while it affected the localization of HIV-1 Rev as expected (19).
Identification of the active nuclear localization signal of mNXF2
We next determined the nuclear localization signal (NLS) of mNXF2 using GFP-tagged deletion mutants (Figure 4A). We found that mutant 1–264 which lacks 407 amino acids from the C-terminus (Figure 4A) lost the nuclear rim association as expected, because it lacks the conserved UBA-like domain (see Figure 1C). Mutant 1–80GFP still localized to the nucleus (Figure 4A, 1–80GFP), whereas further deletion to amino acid 70 resulted in cytoplasmic accumulation of the mutant protein (Figure 4A, 1–70GFP). This suggests a NLS located between amino acid 1 and 80 at the N-terminus.
Since GFP is a small protein, a GFP-tagged small polypeptide may localize to the nucleus due to passive diffusion. To distinguish passive diffusion from active import, we used GFP-β-galactosidase (Gβgal) fusion protein as a tag (13). The Gβgal fusion protein is localized to the cytoplasm because the fusion protein has a higher molecular mass, and neither GFP nor βgal contains an active nuclear import signal (Figure 4B). In contrast, insertion of the N-terminal 80 residues of mNXF2 (1–80Gβgal) conferred nuclear localization on the otherwise cytoplasmic GβGal, demonstrating that mNXF2 contains an active nuclear import signal. We also tested this nuclear import signal using GST–GFP (13) as a tag (data not shown). The GST–GFP fusion protein is localized to the cytoplasm because GST can form a dimer, and neither GST nor GFP contains an active nuclear import signal. Fusion of GST–GFP to amino acids 2–80 or to amino acids 40–80 of mNXF2 resulted in nuclear localization (data not shown).
We further mapped the minimal NLS within amino acids 1–80 of mNXF2 and examined the localization of several deletion mutants (Figure 4B). For the N-terminal deletion mutants, we found that deletion of amino acids 1–39 did not alter nuclear localization of the mutant protein (40–80Gβgal), whereas further deletion to residue 49 (49–80Gβgal) resulted in cytoplasmic accumulation of the fusion protein. Deletion of 10 amino acids from the C-terminus (1–70Gβgal) also abolished nuclear localization of the protein. These results demonstrate that amino acids 40–49 and amino acids 71–80 define the N- and C-terminal boundaries of the mNXF2 NLS. Taken together, these results demonstrate that amino acids 40–80 of mNXF2 are necessary and sufficient to promote nuclear import and this peptide constitutes the minimal NLS.
Fine mapping of the mNXF2 NLS
Inspection of the N-terminal region (40-QGRKRGVNY-48) and the C-terminal region (71-MKRRRERCSY-80) of mNXF2 revealed stretches of basic amino acids. To further study their role in NLS function, alanine substitutions were introduced within 40–80Gβgal (Figure 4C). In mutant M4, changes of three basic amino acids in the N-terminal part of the NLS domain (R42A, K43A, R44A) led to an impairment of the NLS and significant cytoplasmic accumulation of the mutant protein. Double alanine substitutions in the C-terminal part of the NLS as in mutants M10 (K72A, R73A), M6 (R73A, R74A), and M5 (R74A, R75A) resulted in loss of NLS function, whereas mutant M3 (E76A, R77A) has a fully functional NLS. Using single alanine substitutions, we further found that the NLS function is abrogated in M11 (K72A) and M7 (R73A) or significantly affect in M9 (R75A), but the NLS function is only slightly affected in M8 (R74A). Taken together, our results demonstrate that the mNXF2 core NLS consists of two short stretches of basic amino acids (amino acids 42–44 and amino acids 72–75), which are essential for maintaining full NLS activity. Similarly, we previously found that the NLS of the human TAP/NXF1 is bipartite consisting of a C-terminal region contributing (Figure 1C, dotted line) and an N-terminal region (solid line) being essential for NLS activity (13). In both the human TAP/hNXF1 and mNXF2, the core NLS is composed of few basic residues located toward the N-terminus of the proteins.
Association of mNXF2 and mNXF7 with nuclear envelope
As previously shown for human TAP/hNXF1, mNXF2-GFP protein also localized to the nuclear envelope (Figures 3 and 5A). Fusion of the C-terminal portion 545–671 to GFP-βgal led to the accumulation of the mutant protein in the cytoplasm and the nuclear periphery (Figure 5A, 545–671Gβgal), whereas fusion to GFP resulted in diffusion of the protein to the nucleus and accumulation at the nuclear rim. Deletion mutant 565–671GFP still accumulated at the nuclear rim, whereas further deletion to amino acids 581 abolished rim association (581–671GFP). Rim association of the mutants 545–671GFP and 565–671GFP was not affected by digitonin treatment prior to fixation (data not shown), indicating strong interactions with the nuclear envelope. Therefore, mNXF2 contains a signal for rim association at the C-terminus, which shows conservation in location and sequence with other human NXF proteins (7) (Figure 1C).
Interestingly, we found that although GFP-tagged full-length mNXF7 or the N-terminal deletion 95–620GFP did not show apparent nuclear rim association, the deletion of the N-terminal 263 residues led to the accumulation of the protein in the nucleus and to its association with the nuclear rim (Figure 5B, 264–620GFP). Inspection of mNXF7 protein sequence confirmed the conservation of the C-terminal portion containing the nuclear rim association determinants (Figure 1C). By analogy, we previously reported that deletion of the N-terminal NLS of TAP/hNXF1 resulted in a mutant protein, which was found to accumulate at the nuclear rim and the nucleoplasm, suggesting that the ability of NXF to associate with mobile factors of the nuclear pore complex (NPC) facilitates its nuclear import even in the absence of the active N-terminal NLS (13,17,18). We conclude that although mNXF7 protein shares the key signals with the prototype TAP/hNXF1, other regions within the protein are responsible for its distinct localization.
Mapping of mNXF7 domain responsible for granule formation
A key property distinguishing mNXF7 from the other members of the NXF family is its localization to cytoplasmic granules. Figure 6A shows a series of N- and C-terminal deletion mutants of mNXF7 designed to identify the domains responsible for granule formation. Removal of part of the LRR, the NTF2 and the UBA-like domains as well as the N-terminal 95 residues did not abolish the localization to granules. The minimal region necessary and sufficient for granule formation spans amino acids 95–274, and further truncation to amino acids 110–210 resulted in uniform localization of the mutant (Figure 6). The localization of some mutants was further verified using 3D rendering, an approach that allows to better distinguish the granular and homogeneous localization patterns. As shown in Figure 6B, this analysis indeed led to better visualization of the granular pattern of the wild-type mNXF7 and its mutants encompassing amino acids 95–562, 1–274 and 95–274, whereas the mutant spanning amino acids 110–210 showed mostly homogeneous localization, confirming the granule phenotypes presented in Figure 6A. We note that the granule localization determinant in mNXF7 corresponds to part of TAP/hNXF1's ‘substrate-binding domain’ through which it interacts with nuclear export cofactors and assembles with mRNP complexes (2,7,20–22).
mNXF2, but not mNXF7, can promote mRNA export
To test whether mNXF2 or mNXF7 play a role in mRNA export, we used the CAT reporter plasmid pDM128. Plasmid pDM128 contains CAT coding sequence and the Rev response element located within the env intron of HIV-1, flanked by a splice donor and acceptor sites (23). In 293 cells, such transcripts are retained in the nucleus and, therefore, produce only background levels of CAT enzyme, whereas coexpression of export activators, such as HIV-1 Rev or TAP/hNXF1, leads to the stimulation of CAT production. We have previously demonstrated that TAP/hNXF1 indeed enhances the nuclear export of CAT transcripts (22); therefore, CAT production directly reflects the nuclear export of this reporter mRNA.
As a positive control, we coexpressed pDM128 with HIV-1 Rev and found a strong (∼10-fold) activation of CAT production (Figure 7A), as expected (23). We further showed that the presence of TAP/hNXF1 resulted in an ∼2-fold activation, whereas cotransfection with its cofactor, the human p15/NXT1, led to ∼10-fold activation, in agreement with previous observations (7,24), whereas the presence of p15/NXT1 alone had no effect. Using this assay, we found that cotransfection of mNXF2 alone yielded a significant ∼5-fold activation of CAT expression, demonstrating that mNXF2 is a bonafide mRNA export factor. Cotransfection with human or mouse p15/NXT1 did not significantly affect mNXF2 activity. This is in contrast to the human TAP/hNXF1, which depends on the exogenous p15/NXT1 for maximal activity in this assay. We hypothesize that the mNXF2 has a higher affinity to the NPC similar to a TAP/hNXF1 mutant having a duplication of its nucleoporin-binding sites (25) or to endogenous p15/NXT1; therefore, it can act independently of exogenous p15/NXT1. In contrast, mNXF7 failed to activate CAT production both in the absence and presence of p15/NXT1 (Figure 7A), consistent with the lack of mRNA export function of this cytoplasmic protein. This finding suggests that the mouse NXF7 protein functions at post-export steps in the mRNA metabolism. Similar results were obtained in mouse PA317 cells (data not shown).
To understand the mechanism of mNXF2's nuclear export activity, we studied its interactions with Y14/MAGOH, the REF proteins and U2AF35, which are known to bind to TAP/hNXF1 directly and are thought to facilitate the addition of TAP/hNXF1 to its export substrates (22,26–30). The bacterially produced recombinant proteins spanning the substrate-binding domains of mNXF2 and TAP/hNXF1 (amino acids 1–400) were immobilized and used in pull-down assays together with metabolically labeled binding factors. As a negative control, we used the mRNA export factor DBP5 (31), which does not bind TAP/hNXF1 directly. Figure 7B shows that MAGOH, REF2-II and U2AF35 bound readily to both TAP/hNXF1 and mNXF2, whereas the binding of Y14 and DBP5 was low or undetectable. The observed binding of MAGOH, REF2-II and U2AF35 to TAP/hNXF1 is consistent with previous findings (22,26–30). Since the MAGOH subunit binds to TAP/hNXF1 much more avidly than Y14 (28), the lack of REF1-II and Y14 binding confirmed that only strong interactions were revealed under our assay conditions. However, we cannot exclude that the reticulocyte-produced REF1-II and/or Y14 were misfolded or lacked proper post-translational modifications, and this possibility was not further addressed. Although TAP/hNXF1 and mNXF2 proteins were used at similar amounts, the overall binding efficiency of the tested proteins to TAP/hNXF1 was higher. This effect could be attributed to a better folding of this recombinant protein, and it was not further examined. In conclusion, this study shows that mNXF2 is an mRNA export factor sharing the localization and functional properties with TAP/hNXF1.
DISCUSSION
In this report, we present the identification and functional characterization of the mouse Nxf family genes and their corresponding proteins. Since some mammalian NXF factors are believed to be expressed specifically in the brain, and because human hNXF5 nullisomy is linked to mental retardation (12), we wished to understand the possible roles of such proteins. We have chosen to study the Nxf gene family in mouse, because this species is widely used as a model to study human neurological disease, yet possesses only two Nxf1-related genes.
The mouse mNXF2 and the human hNXF2 are expressed in the brain
Previously, mRNA analysis was used as evidence of brain-specific expression for the NXF factors, such as mNXF2, mNXF7, and hNXF5. Using hNXF2 as a model, we found that this approach can lead to gross overestimation of the predicted protein expression levels, because of high abundance of aberrant, non-coding splice products in certain tissues such as testis (A. Zolotukhin, S. Lindtner, S. Smulevitch and B.K. Felber, unpublished data). We therefore sought to verify the brain-specific expression of NXF family factors at the protein level and raised antibodies to mNXF2 and its human counterpart, TAPX2/hNXF2. By using these antibodies, we show here that the expression of the human TAPX2/hNXF2 protein is restricted to the brain (Figure 2C), and that the mouse mNXF2 expression levels in the brain are developmentally regulated (Figure 2B), suggesting brain-related function(s) of these proteins.
mNXF2 is an active mRNA export receptor, with properties similar to those of TAP/hNXF1
We show here that mNXF2 behaves like a typical NXF factor, since it shows mRNA export activity, localizes mostly to the nucleus and the nuclear envelope, and contains active nuclear localization and NPC-association signals that are positioned similar to those of TAP/hNXF1. We also show that mNXF2 and the human TAP/hNXF1 share the ability to interact with mRNA splicing/export cofactors, such as MAGOH, REF and U2AF35, strongly suggesting a shared nuclear export mechanism. While this manuscript was under review, another group (32) has reported that mNXF2, but not mNXF7, binds in vitro to the TAP/hNXF1 export cofactor p15, further supporting our conclusions. The few differences we observed between mNXF2 and the prototype export factor, TAP/hNXF1, include (i) a more pronounced cytoplasmic and nuclear envelope localization of mNXF2, whereas TAP/hNXF1 is mostly found in the nucleoplasm (ii) mNXF2 shows a more relaxed requirement for the cofactor p15/NXT1 in the CAT nuclear export assays used.
We believe that these small differences do not likely account for a brain-specific role of mNXF2 that is distinct from that of TAP/hNXF1. Rather, we propose that mNXF2 functions differently from TAP/hNXF1 due to its restricted substrate specificity. In this model, mNXF2, unlike the promiscuous TAP/hNXF1, only associates with a specific subset of transcripts in the brain, leading to their regulation during development. Further work is required to characterize such transcripts and the responsible determinants within mNXF2.
mNXF7 is atypical of NXF family
Unexpectedly, we found that mNXF7 is a cytoplasmic protein that is completely excluded from the nucleus. However, these localization data at steady-state do not rule out that mNXF7 can enter the nucleus, especially considering the presence of a cryptic NLS (see below). Independent of its fusion to detection tags (such as GFP and HA), species and cell lines, mNXF7 forms granules in the cytoplasm and is targeted to these granules via a region that spans amino acids 95–274. Interestingly, when devoid of this region, mNXF7 assumes the subcellular localization that is typical of a generic NXF factor, revealing nuclear import and nuclear envelope localization determinants. Apparently, these determinants are overridden in the wild-type mNXF7 protein by a potent granule localization signal. However, the conservation of active, NXF-like signals within mNXF7 points to their functional importance and suggests that the role of this protein may require nuclear entry and NPC binding. In agreement with its unusual localization, mNXF7 did not show nuclear export activity in CAT assays, probably due to its absence from or short dwelling time in the nucleus. We cannot exclude though, that mNXF7 may have an export activity that is strictly specific toward a subclass of transcripts and is not revealed using the cat mRNA reporter. In summary, we propose that mNXF7 plays a highly specialized role(s) in the cytoplasm that is distinct from that of other NXFs studied so far. Based on homology to TAP/hNXF1 and shared domain organization, it is likely that mNXF7 is engaged in mRNA metabolism. At present, we do not know whether such proposed activities include the mRNA nuclear export per se, or are restricted to post-export events. Understanding the precise biological role of mNXF7 and the underlying molecular mechanisms remains a goal for future studies.
Acknowledgements
We thank S. Lockett, J. Brenner, E. Cho and S. Costes for help with confocal microscopy, P. Aplan, G. Dreyfuss, E. Izaurralde and F. Stutz for their generous gifts of plasmids and T. Jones for editorial assistance. We are grateful to our summer students P. Sood and A. Witten, and our Werner H. Kirsten Student Intern program recipients D. Waddelow and C. Jodrie for their contributions. Funding to pay the Open Access publication charges for this article was provided by NCI.
Conflict of interest statement. None declared.
Figures and Tables
Figure 1
Mouse Nxf2 and Nxf7. (A) Exon–intron structure of mouse Nxf2 and Nxf7 genes. The cDNAs were isolated and the sequences and the genomic structure were determined. Exons are shown in black boxes. ATG, translational initiation codon; TAA and TGA, translation termination codons. Asterisks indicate location of in-frame ATG codons. (B) Dendrogram illustrating the NXF family of proteins. Abbrevations: rn, Rattus norvegicus; cj, Coturnix japonica; dm, Drosophila melanogaster; sp, Schizosaccharomyces pombe; sc, Saccharomyces cerevisiae. The mouse family members are shown in bold. (C) Multiple sequence alignment of human TAP/hNXF1 and mouse NXF proteins. The protein domains based on Herold et al. (7) are indicated. The identified domains mediating nuclear localization of human TAP/hNXF1 and mNXF2, localization to cytoplasmic granules of mNXF7 and the rim association of mNXF2 are underlined. RBD, RNA-binding domain; LRR, leucine-rich repeat; NTF2, nuclear transport factor (NTF2)-like domain; UBA-like, ubiquitin associated-like domain.
Figure 2
Specific expression of NXF-related proteins in brain tissue. (A) Western immunoblot analysis of 293 cells transfected with untagged and tagged (HIS, GFP and Gβgal) mNXF2 (left panel) and different NXF expression plasmids (right panel) using a rabbit anti-mNXF2 antiserum. (B) Western blot analysis of mouse brain tissue using monospecific rabbit anti-mNXF2 antiserum. Pre-made ‘mouse brain aging’ blots (RNAWAY) contained normalized amounts of whole brain lysates from fetus to 1-year-old as indicated. (C) Western blot analysis of human tissues using the monospecific antiserum to human TAPX2/hNXF2. Pre-made blots (GenoTech) contained normalized amounts of proteins from the indicated tissues.
Figure 3
Distinct subcellular localization of mNXF2 and mNXF7 proteins. HeLa cells were transfected with GFP-tagged mNXF2 and mNXF7 expression plasmids, as indicated and the proteins were visualized in living cells. The images were obtained by fluorescent microscopy (Axiovert135TV, Zeiss) and by the use of a CCD camera and processed using IPLab Spectrum software (13). Similar results were obtained in numerous experiments and are typical of the vast majority of expressing cells in each individual experiment. Representative cells are shown.
Figure 4
Identification of NLS of mNXF2. (A and B) HeLa cells were transfected with plasmids producing mNXF2 and N- and C-terminal deletions fused to GFP (A) or GFP-βgal (B) and the proteins were visualized as described in Figure 2. The localization of the proteins is indicated. N, nucleus; C, cytoplasm. (C) Fine mapping of the NLS. Alanine substitution in 40–80Gβgal generated a series of mutants and the mutant proteins are visualized. The protein sequence containing the NLS regions is shown and the critical residues are indicated in bold.
Figure 5
Association of mNXF2 and mNXF7 with nuclear envelope. HeLa cells were transfected with plasmids producing the indicated GFP-tagged mNXF2 and mNXF7 deletion mutants and the images were analyzed as described in Figure 2.
Figure 6
Identification of signal mediating granule localization of mNXF7. (A) A panel of N- and C-terminal deletion mutants of GFP-tagged mNXF7 deletion mutants was analyzed upon transfection of HeLa cells. N, nucleus; C, cytoplasm. Confocal images of GFP fluorescence in the mid-sections through the nuclei. (B) Three-dimensional rendering of GFP images from confocal Z-stacks.
Figure 7
mNXF2 is an active mRNA export factor. (A) Human 293 cells were transfected with the cat reporter pDM128 either alone, in the presence of HIV-1 rev expression plasmid or in the presence of the indicated untagged NXF producing plasmids. As indicated, the transfection mixtures contained NXF expression plasmids together with human p15/NXT1 or p15/NXT1 alone. Two days later, the cells were harvested and the CAT production, measured as percentage of chloramphenicol acetylation, is shown by filled bars. Expression of the cotransfected luciferase expression plasmid was analyzed for each plate and the relative luciferase values are shown in open bars. A typical experiment is shown. (B) mNXF2 binds to TAP/hNXF1 export cofactors. Bacterially produced GST-tagged NXF proteins were immobilized on glutathione–Sepharose beads and used in pull-down assays with reticulocyte-produced, metabolically labeled factors (shown to the left). The bound (B) and 1:100 aliquots of the unbound (U) fractions were separated on SDS–PAGE and visualized on Phosphoimager. | [
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16410827 | The Notch Ligand JAG1 Is Required for Sensory Progenitor Development in the Mammalian Inner Ear
Abstract
In mammals, six separate sensory regions in the inner ear are essential for hearing and balance function. Each sensory region is made up of hair cells, which are the sensory cells, and their associated supporting cells, both arising from a common progenitor. Little is known about the molecular mechanisms that govern the development of these sensory organs. Notch signaling plays a pivotal role in the differentiation of hair cells and supporting cells by mediating lateral inhibition via the ligands Delta-like 1 and Jagged (JAG) 2. However, another Notch ligand, JAG1, is expressed early in the sensory patches prior to cell differentiation, indicating that there may be an earlier role for Notch signaling in sensory development in the ear. Here, using conditional gene targeting, we show that the Jag1 gene is required for the normal development of all six sensory organs within the inner ear. Cristae are completely lacking in Jag1-conditional knockout (cko) mutant inner ears, whereas the cochlea and utricle show partial sensory development. The saccular macula is present but malformed. Using SOX2 and p27kip1 as molecular markers of the prosensory domain, we show that JAG1 is initially expressed in all the prosensory regions of the ear, but becomes down-regulated in the nascent organ of Corti by embryonic day 14.5, when the cells exit the cell cycle and differentiate. We also show that both SOX2 and p27kip1 are down-regulated in Jag1-cko inner ears. Taken together, these data demonstrate that JAG1 is expressed early in the prosensory domains of both the cochlear and vestibular regions, and is required to maintain the normal expression levels of both SOX2 and p27kip1. These data demonstrate that JAG1-mediated Notch signaling is essential during early development for establishing the prosensory regions of the inner ear.
Synopsis
Deafness and adult-onset hearing loss are significant health problems. In most cases, deafness or vestibular dysfunction results when the sensory cells in the inner ear, known as hair cells, degenerate due to environmental or genetic causes. In the mammalian inner ear, the hair cells and their associated supporting cells can be found in six different patches that have particular functions related to hearing or balance. Unfortunately, unlike in birds or fish, mammalian hair cells show little ability to regenerate, resulting in a permanent hearing or balance disorder when damaged. Here, the authors show that a protein called JAG1, a ligand in the Notch signaling pathway, is required for the normal development of all six sensory regions in the mammalian inner ear. In ears that lacked JAG1, some of the sensory patches were missing completely, whereas others were small and lacked particular cell types. The authors showed that JAG1 is required by the sensory precursors, progenitor cells that give rise to both the hair cells and the supporting cells. By understanding how the sensory areas develop normally, it is hoped that molecular tools can be developed that will aid sensory regeneration in the mammalian inner ear.
Introduction
The mammalian inner ear is a complex structure consisting of a coiled cochlea, three orthogonally positioned semicircular canals, a central vestibule, and a dorsally projecting endolymphatic duct and sac. With the exception of the endolymphatic duct and sac, the different parts of the ear all contain sensory organs populated by sensory hair cells and their associated supporting cells. There are three different categories of sensory organs: cristae, located at the base of each semicircular canal; maculae, housed within the central vestibule; and the organ of Corti, which lines the cochlear duct. Only one sensory organ, the organ of Corti, is required for hearing; the other five organs are important for balance. Unfortunately, in mammals, if these regions are damaged due to an environmental or genetic insult, they cannot regenerate, leaving a permanent hearing and/or balance impairment.
Although some progress has been made in understanding how the individual cell types within the sensory areas of the ear are formed [1,2], little is known about the molecular mechanisms that establish the prosensory lineage and how the different sensory organ types are formed. Interestingly, the molecular mechanisms that underlie sensory differentiation in the vertebrate inner ear demonstrate strong parallels with Drosophila sense organ development [3–5]. For example, during Drosophila external sense organ development, lateral inhibition mediated by Notch signaling is required to restrict the adoption of the sensory organ precursor cell fate, which then gives rise to the entire sensory organ [6–8]. Similarly, in the vertebrate ear, lateral inhibition mediated by Notch signaling appears to be important for restricting the number of cells that can adopt the hair cell fate [9–15]. Lineage analysis has also shown that, at least in the chicken, hair cells and supporting cells arise from a common progenitor [16], consistent with an equipotent epithelium that undergoes lateral signaling to specify cell fates. Unlike in Drosophila, which has a single Notch receptor and two ligands (Delta and Serrate/Jagged), in mammals Notch signaling pathway components include four receptors (Notch 1–4) and five ligands (Delta-like [DLL] 1, 3, and 4, and Jagged [JAG] 1 and 2; for reviews, see [8,17–19]). In the mouse, both DLL1 and JAG2 are expressed in nascent hair cells [10,20] and act synergistically during lateral inhibition [15]. Both DLL1 and JAG2 appear to signal through the NOTCH1 receptor [15]. In Drosophila, prosensory regions that undergo lateral inhibition are first delineated by expression of members of the atonal or acheate-scute family of basic helix-loop-helix transcription factors. Further parallels with Drosophila have arisen with the finding that an atonal homolog, the Math1 gene, is required for hair cell differentiation [21–23]. However, the situation is not entirely similar to Drosophila, as MATH1 does not appear to be required to establish the prosensory regions [22,24]. Instead, it has turned out that another type of transcription factor, the HMG-box factor SOX2, is required for sensory organ formation in the inner ear [25]. This finding does not demonstrate direct parallels with Drosophila sense organ formation, since the Drosophila SOX2 homologs SoxNeuro and Dicheate have no known role in peripheral sense organ formation. Instead, both genes play a role in the formation of neural progenitor cells in the central nervous system [26,27]. Consistent with a prosensory role, SOX2 marks sensory progenitors early in development and acts upstream of the Math1 gene during sensory organ formation in the ear [25]. Interestingly, one of the Notch ligands, JAG1, is expressed early in the prosensory regions of the ear [4,20], indicating that Notch signaling also may play a role in early sensory organ formation. Consistent with this finding, mice heterozygous for N-ethyl-N-nitrosourea-induced point mutations in the Jag1 gene show mild sensory organ defects in the ear [28,29]. Unfortunately, embryos homozygous for available mutant alleles of the Jag1 gene do not survive past embryonic day (E) 11.5 due to vascular defects, precluding an analysis of their inner ears. To circumvent this early lethality, we have created a conditional allele of the Jag1 gene using Cre/loxP technology. Using the Foxg1-Cre mouse line to express Cre recombinase in the early otocyst [30,31], we have disrupted JAG1 function in the ear and show that sensory formation in the inner ear is severely attenuated in these mutants. Analysis of the patterns of hair and supporting cell formation in the Jag1-conditional knockout (cko) inner ears suggests that fewer progenitors form in Jag1-cko inner ears. This result is confirmed by analysis of the prosensory markers SOX2 and p27kip1, which are down-regulated as early as E12.5, indicating that the Jag1 gene acts early during sensory progenitor formation. These data demonstrate an early role for Notch signaling in establishing the sensory progenitors of the inner ear.
Results
Creation of a Conditional Allele of the Jag1 Gene
We created an allele for conditional inactivation of JAG1 function by flanking the Delta-Serrate-Lag2 (DSL) domain-encoding exon (exon 4) of the Jag1 gene with loxP sites (Figure 1). The DSL domain has been shown to be the region of the DLL and JAG proteins that interacts with Notch family receptors [32,33]; therefore, removing this region of the gene should create a nonfunctional protein. To demonstrate that this allele encodes a nonfunctional protein, we crossed Jag1flox/+ mice to mice expressing Cre recombinase in the female germline under control of the Zp3 promoter (Zp3-Cre mice); the Zp3-Cre mouse strain has been shown to express Cre recombinase in the growing oocyte prior to the completion of the first meiotic division [34]. Female Zp3-Cre/+; Jag1flox/+ offspring were then crossed to male B6 mice to produce offspring that were heterozygous for the deleted region of the floxed allele (designated Jag1del2/+; see Materials and Methods). Heterozygous Jag1del2/+ mice were intercrossed, and Jag1del2/Jag1del2 homozygous offspring were analyzed for defects between E9.5 and E11.5. Jag1del2/Jag1del2 mutant embryos exhibited the same vascular phenotype we described previously in embryos homozygous for a targeted Jag1 null allele, Jag1del1 [35]. Specifically, the Jag1del2/Jag1del2 mutant embryos exhibited yolk sac vascular remodeling defects and cranial hemorrhaging, and often exhibited an enlarged pericardial sac (Figure 2A–2D). All Jag1del2/Jag1del2 mutant embryos were necrotic by E11.5, and most showed vascular defects by E10.5. RT-PCR of cDNA synthesized from the mutant embryos using primers that span the floxed exon 4 demonstrated that this region was deleted, as expected (Figure 2E). These data demonstrate that deletion of the Jag1flox allele yields a nonfunctional Jag1 mutant allele.
Figure 1
Construction of a Conditional Allele of the Jag1 Gene
(A) Schematic diagram showing the strategy for generating Jag1flox mice. The targeting vector was designed to insert loxP sites (black arrowheads) on either side of exon 4, the DSL domain-encoding exon (white area in exon 4). The neomycin resistance cassette (for positive selection) was flanked by FRT sites (gray arrowheads) so that it could later be removed by crossing to FLPe-expressing mice. A diphtheria toxin gene was included for negative selection. Dotted lines depict the recombination events that occur when either the FLPe or Cre recombinases are present. Primer positions for genotyping are shown as small black arrows (a, DSLF; b, J1LoxR1; c, J1FlpF1; see Materials and Methods for sequences). DT, diphtheria toxin; R, EcoR1; X, Xba1.
(B) Southern blot analysis of EcoRI-digested DNA from ES cells using the external probe shown in (A). Left lane shows the wild-type band (12.3 kb), and the center and right lanes show correctly targeted ES cells that have both a wild-type band and a smaller mutant band (9.4 kb).
Figure 2
Jag1del2/Jag1del2 Embryos Exhibit Vascular Defects and Lethality Consistent with Loss of JAG1 Function
(A and B) E10.5 embryos demonstrating the loss of large blood vessels (white arrows in [A]) in the Jag1del2/Jag1del2 yolk sacs (B) similar to other Jag1 loss-of-function mutants. (C and D) E11 embryos demonstrating a small, necrotic Jag1del2/Jag1del2 embryo (D). White arrow in (D) indicates an enlarged pericardial sac (enlarged, inset), which is frequently observed in mutants exhibiting cardiovascular defects. RT-PCR results using primers that span exon 4 using RNA extracted from E10.5 control (+) and ZP3-Cre deleted embryos (−) (E). The upper band (541 bp) indicates that the wild-type allele is present. The lower bands (286 bp) indicates the Jag1del2 mutant allele that does not contain exon 4.
Inactivation of Jag1 Function within the Ear
To disrupt JAG1 function within the inner ear, we crossed Jag1flox/Jag1flox mice with mice doubly heterozygous for the Foxg1-Cre allele (these mice express Cre recombinase throughout the otocyst, as well as forebrain, eye, and foregut) [30] and the Jag1del1 allele [35]. Offspring with the genotype Foxg1-Cre/+; Jag1del1/Jag1flox (hereafter designated Jag1-cko) survived through E18.5 and were analyzed for inner ear defects. We examined the patterns of Cre-mediated excision in Jag1-cko embryos at E10.5 and in cochleae at E16.5–E18.5 by in situ hybridization using a probe that specifically detected the deleted exon 4 (Figure 3). These results showed that expression in the otocyst was weak or absent by E10.5 (Figure 3B). In addition, analysis of cochlear expression at later stages showed no expression at E16.5 (Figure 3F) and E18.5 (unpublished data). These data indicate that, as previously shown for conditional deletion of a Fibroblast growth factor receptor 1 (Fgfr1) floxed allele [31], the Foxg1-Cre line efficiently deletes the Jag1flox allele early during inner ear development.
Figure 3
Conditional Jag1 Inactivation Using the Foxg1-Cre Line
(A–D) Low- and high-power views of E10 embryos processed for whole-mount in situ hybridization using a Jag1 exon 4-specific probe. White arrows (A) point to the Jag1 signal in the otocyst (left arrow) and the eye (right arrow), two structures where Cre recombinase is expressed. In Jag1-cko mutants at E10, this signal is either absent or extremely weak. Black arrows (A and B) point to expression in the spinal cord and nephric duct, regions where Cre recombinase expression has not been reported in Foxg1-Cre mice. However, expression is consistently weaker in these areas in Jag1-cko embryos, indicating that there may be low levels of widespread expression of Cre recombinase in Jag1-cko embryos. In (D), the otocyst and the eye are outlined by a dotted line. Very little expression is observed in these regions, consistent with Foxg1-Cre expression.
(E and F) In situ hybridization of E16.5 cochleae demonstrating Jag1 expression in wild-type (E) and Jag1-cko cochleae (F), where expression is entirely absent. Scale bars = 500 μm.
Malformation of the Inner Ear in Jag1-cko Mutants
To examine the morphology of the Jag1-cko inner ears, paintfilling of the inner ears of mutants and controls was performed at E15.5 (Figure 4). Results of this analysis showed a severe disruption in the structure of the Jag1-cko inner ears compared to their littermate controls (Figure 4C and 4D). Specifically, the semicircular canals were largely absent, with the exception of a portion of the anterior and lateral semicircular canals. In addition, the utricle appeared small, the saccule was misshapen, and the cochlea was undercoiled. In contrast, the parts of the inner ear that are not associated with sensory formation, including the endolymphatic duct and sac and the common crus, appeared relatively unaffected.
Figure 4
Inner Ear Dysmorphology in Jag1+/− and Jag1-cko Embryos
E15.5 inner ears that have been paintfilled to display their overall morphology.
(A) Wild-type inner ear showing normal morphology. Structures are labeled as follows: aa, anterior ampulla; asc, anterior semicircular canal; cd, cochlea duct; ed, endolymphatic duct; la, lateral ampulla; lsc, lateral semicircular canal; pa, posterior ampulla; psc, posterior semicircular canal; sac, saccule; ut, utricle.
(B) Jag1 heterozygote inner ears (either Jag1del1/+ or Foxg1-Cre/+; Jag1flox/+) display truncated posterior semicircular canals and missing ampullae (asterisk). Arrows point to the anterior and posterior ampullae, which are small compared to the wild-type control (A).
(C and D) A much more severe phenotype is observed in Jag1-cko animals. There are no ampullae and little semicircular canal development; an asterisk indicates a remnant of the anterior canal. In (D), there is also a remnant of the lateral canal (arrow). The utricle and saccule are smaller and the cochleae are shorter and undercoiled. Scale bar = 500μm.
Sensory Defects in Jag1-cko Mutant Inner Ears
Since the Jag1 gene is expressed in the sensory areas of the ear, and because the structural malformations observed in Jag1-cko mutant inner ears appeared to primarily affect regions of the ear that contained sensory organs, we examined the sensory regions of the ear for defects. We examined the organ of Corti, the sensory organ of the cochlea, at E18.5 by scanning electron microscopy (SEM) (Figure 5). By this stage, all hair cells within the organ of Corti have exited the cell cycle, and most are well-differentiated although not fully mature [36]. Severe hair cell patterning defects were apparent by SEM within the Jag1-cko mutant cochleae. This phenotype was most striking in the basal turns of the cochlea, where no hair cell formation was observed (Figure 5D). In the midbasal regions of the organ of Corti, hair cells formed in patches, within which there was no clear formation of rows or distinction between inner and outer hair cells (Figure 5F). More apically, hair cells appeared more continuous along the organ of Corti (Figure 5H). However, although hair cells were present in the apical region, their numbers were clearly reduced; rather than the normal, perfectly ordered four rows of hair cells, there were only two rows of loosely arranged hair cells of indistinct type.
Figure 5
Hair Cell Patterning Defects in the Cochlea
Scanning electron micrographs demonstrating the different patterns of hair cell production along the length of the cochlea in Jag1-cko embryos.
(A–D) Low-power views of the apical and basal cochlear turns. The boxed-in area along the base in (A) and (B) is shown at higher magnification in (C) and (D). Note the absence of hair cells in the base of the Jag1-cko cochlea, except for a small patch of cells in the more apical portion (arrow). Scale bars = 500 μm.
(E and F) In the midbasal region, more hair cells are observed, but they are arranged in patches, with no clear distinction between inner and outer hair cells.
(G and H) In the apical turn, hair cells are continuous but generally arranged in only two rows. Scale bar = 100 μm.
Abnormal Hair and Supporting Cell Patterns in Jag1-cko Inner Ears
To determine which sensory cell types were differentiating in the Jag1-cko mutant cochleae, specific markers were used to identify hair cell and supporting cell subtypes throughout the ear (Figure 6). In the cochlea, we used an antibody against MYO7A to label all hair cells and an antibody against S100A1 to label inner hair cells, Deiter's supporting cells, and inner phalangeal supporting cells [24]. When both markers were used in combination, inner hair cells, outer hair cells, and some supporting cell types could be distinguished (Figure 6A–6F). This analysis showed that in the apex of the cochlea, inner hair cells were present and usually formed as doublets (Figure 6B). Their associated supporting cells, the inner phalangeal cells, were also present. Outer hair cells and their associated supporting cells, the Deiter's cells, were not present in this region. In the middle portions of the cochlea, both inner and occasionally outer hair cells were present, although their patterning was clearly abnormal (Figure 6D). In addition, the tunnel of Corti was not apparent, and there were often doublets of inner hair cells and increased numbers of outer hair cell rows without accompanying Deiter's supporting cells. As shown by SEM, both hair cells and supporting cells were absent in the very basal regions of the cochlea (Figure 6F).
Figure 6
Hair and Supporting Cell Markers Demonstrate Sensory Areas Are Reduced or Absent in the Jag1-cko Inner Ear
Immunocytochemistry using two markers, myosin VIIA (red; all hair cells) and S100a (green; inner hair cells, Dieter's cells, and inner phalangeal cells) demonstrate patterns of hair and supporting cell production at E18.5 in control and Jag1-cko inner ears.
(A–F) Sections through the indicated turns of the cochlea. Note the different hair cell patterns in the apical, middle and basal turns of the Jag1-cko cochlea. Normal morphology is shown (A) along with labeled structures, as follows: GER, greater epithelial ridge; IHCs, inner hair cells (color-coded yellow); LER, lesser epithelial ridge; OHCs, outer hair cells (color-coded red); SCs, supporting cells (color-coded green).
(G–J) Patterns of hair and supporting cell production in the vestibular system. The utricular macula is extremely small with very few hair cells (H) while the saccule (J) shows robust hair and supporting cell production although the shape of the organ is smaller and malformed.
Using the same markers we also examined the vestibular sensory organs in Jag1-cko mutant inner ears (Figure 6G–6J). Consistent with the lack of semicircular canal and ampulla formation observed by paintfilling, there was no evidence of crista formation. The Jag1-cko utricular macula was extremely small with very few differentiating hair cells (Figure 6H). Surprisingly, the saccule and its macula were only mildly affected in the Jag1-cko inner ears (Figure 6J). Hair cell differentiation appeared relatively unaffected, although the entire saccular structure was shaped differently than in the controls, a feature that was also observed in the paintfilled specimens (see Figure 4C and 4D). These data show that all sensory organs within the inner ear are affected to varying degrees in Jag1-cko inner ears. However, some sensory organs, such as the cristae, appear to be more sensitive to the loss of JAG1 function.
To examine whether aberrant hair cell patterning in Jag1-cko cochleae was due to defects in hair cell formation or in subsequent differentiation, we examined hair cell patterning at an earlier stage (E16.5). Using a lectin that binds to hair cell stereocilia, we examined whether the patterns of hair cell formation at E16.5 looked similar to the patterns at E18.5 (Figure 7). At E16.5 in wild-type cochleae, a gradient of hair cell differentiation was evident (Figure 7A, 7C, 7E, and 7G); in the basal regions both inner and outer hair cells could be recognized (unpublished data), while in the middle regions only inner hair cells were clearly detected by most markers (Figure 7A and 7C). In the more apical regions, little to no hair cell differentiation had taken place by this stage (Figure 7E and 7G). In Jag1-cko cochleae, the patterns looked similar to those at E18.5, with patches of hair cells in the midbasal regions (Figure 7B and 7D) and a complete absence of hair cells in the very basal regions (Figure 7B). These data suggest that the Jag1-cko mutants have defects in hair cell formation rather than differentiation. In addition, the apical regions in the Jag1-cko cochleae did not appear more differentiated than the controls (Figure 7E–7H), arguing against precocious differentiation as an explanation for the reduced numbers of hair cells observed in the mutant cochleae.
Figure 7
Early Analysis of the Patterns of Differentiation in the Jag1-cko Cochlea Indicates the Defects Are Caused by a Failure in the Formation of Sensory Cells and Not Subsequent Degeneration
Lectin staining of whole-mount cochlea at E16.5.
(A) Normal patterning in wild-type control cochlea. GER, greater epithelial ridge.
(B) Both the basal and middle portions of the cochlea are shown, although because it is much longer in the control (A), the very basal portion of the cochlea has been removed. Note the lack of hair cells in the basal portion of the Jag1-cko cochlea.
(C–H) Boxed-in areas of (A) and (B) are shown at higher magnification in (C) and (D). Arrows in (D) indicate the abnormal patches of hair cells also observed at E18.5. Similarly, the boxed-in regions of (E) and (F) are shown at higher magnification in (G) and (H), demonstrating the few hair cells that are just beginning to differentiate in this region in both the control and the mutant (arrowheads). Scale bars = 500 μm for the corresponding panels.
Disrupted Prosensory Development in Jag1-cko Inner Ears
To determine how the JAG1 ligand functions during sensory development, we used several markers of the prosensory domain, including p27kip1 and SOX2, and examined their expression patterns in both wild-type and Jag1-cko mutant cochleae (Figure 8). At E14.5, the majority of hair cells and supporting cells in the organ of Corti have completed their final division, and hair cells are beginning to differentiate in the basal portions of the cochlea [36]. p27kip1, a cell-cycle inhibitor, is required for the cochlear sensory progenitors to exit the cell cycle on time, and is an established marker of the prosensory domain in the cochlea [22,37]. p27kip1 begins to be expressed in a discrete domain within the cochlea as the hair cells and supporting cells exit the cell cycle around E13.5 to E14.5 (Figure 8A, 8B, 8D, and 8E). Recently it has been shown that the SRY-related transcription factor SOX2 is required for establishment of the prosensory regions in the inner ear [25]. Using fluorescence immunocytochemistry double labeling, we examined the relationship between these markers and JAG1 protein expression in both wild-type and Jag1-cko cochleae. As previously reported [22], JAG1 was not expressed within the prosensory domain as assessed by p27kip1 expression at E14.5, but instead was expressed immediately adjacent (possibly with some slight overlap) in the inner (neural) portion of cochlea (Kölliker's organ; Figure 8A and 8D). In contrast, SOX2 did show a largely overlapping domain with p27kip1 (Figure 8B and 8E), as originally described [25]. However, the SOX2 expression domain was slightly larger than the p27kip1 domain, extending into Kölliker's organ and overlapping with the JAG1 domain. Despite the fact that JAG1 was not expressed within the prosensory domain at E14.5, both p27kip1 and SOX2 expression was absent in the basal regions of the cochlea (Figure 8C), indicating that prosensory formation is already disrupted in these ears. In the apex, weak expression of both markers was observed (Figure 8F), consistent with the fact that some sensory differentiation occurs in this region of the Jag1-cko cochlea.
Figure 8
At E14.5 JAG1 Is Expressed Directly adjacent to the Prosensory Domain That Is Disrupted in Jag1-cko Inner Ears
Immunocytochemistry at E14.5 using two markers of the prosensory domain, Sox2 and p27Kip1, in combination with JAG1 in both the basal and apical turns of the cochlea. Note that the JAG1 domain (red) does not overlap the p27Kip1 domain (green) (A and D), whereas the SOX2 domain does largely overlap with p27Kip1 (yellow) (B and E). Both SOX2 and p27Kip1 are down-regulated in the Jag1-cko cochlea (C and F), although there is weak expression of both markers in the apex (F). GER, greater epithelial ridge; LER, lesser epithelial ridge.
In order to determine if JAG1 is ever expressed in the prosensory region of the cochlea, we examined an earlier age (E12.5) and compared the JAG1 domain to the SOX2 domain (since p27Kip1 is not expressed in the inner ear prior to E13.5 to E14.5). Adjacent sections from both wild-type and Jag1-cko cochleae were immunostained to detect either JAG1 or SOX2 protein (Figure 9). This analysis showed that in the basal regions of the wild-type cochlea, where sensory progenitors were still dividing, JAG1 expression did overlap with the SOX2 domain (Figure 9A and 9B), indicating that JAG1 is initially expressed within the prosensory domain. However, in the apical regions, where the sensory precursors have ceased dividing, expression of JAG1 and SOX2 did not overlap (Figure 9D and 9E). In the Jag1-cko cochlea, SOX2 was absent from the basal regions and significantly down-regulated in the apical regions (Figure 9C and 9F). These data demonstrate that JAG1 is expressed within the prosensory domain of the cochlea at early stages, and that, in the absence of JAG1 function, sensory formation is disrupted prior to cell cycle exit and differentiation of sensory hair cells and nonsensory supporting cells.
Figure 9
At E12.5 JAG1 Is Expressed within the Prosensory Domain and SOX2 Expression Is Down-Regulated within This Domain in Jag1-cko Cochleae
(A, B, D, and E) Alternate sections from a control embryo processed for immunocytochemistry using antibodies against either JAG1 or SOX2. Note the similar domain occupied by both JAG1 and SOX2 in the base of the cochlea (A and B; brackets). In the apical region, the two proteins are not colocalized (D and E). SOX2 is not expressed in the basal portions of the Jag1-cko cochlea (C) and shows only weak expression in the apex (F). bv, blood vessel; cd, cochlear duct.
We also compared JAG1 and SOX2 expression in the vestibular regions of the inner ear in both wild-type and Jag1-cko mutant embryos. JAG1 and SOX2 exhibited largely overlapping expression domains that corresponded to the locations of the five sensory organs in the vestibular portion of the ear (Figure 10). The two expression domains only differed significantly in the anterior and posterior cristae, where JAG1 expression had a negative patch in the middle of its expression domain, whereas SOX2 expression did not show this same patch (Figure 10A, 10B, 10G, and 10H). The JAG1-negative region may correspond to the eminentia cruciatum, a nonsensory region present in the middle of both the anterior and posterior cristae, although it is not clear why SOX2 would be expressed there. In the Jag1-cko vestibular sensory patches, SOX2 expression was consistent with the patterns of sensory differentiation observed at E18.5. For example, the Jag1-cko saccule displayed fairly normal SOX2 expression (Figure 10C), consistent with the almost normal development of the saccular macula. In contrast, SOX2 expression in the utricle was very weak and the expression domain was much smaller than in controls (Figure 10F), consistent with the severe disruption of differentiation of the utricular macula in Jag1-cko inner ears. There was no SOX2 expression in the Jag1-cko cristae, and in fact the entire ampullae appeared to be missing or severely disrupted even at this early stage (Figure 10C and 10I; dotted line regions), consistent with the lack of cristae and ampullae observed at later stages.
Figure 10
JAG1 and SOX2 Mark the Prosensory Regions of the Vestibule, and SOX2 Expression Correlates with Impaired Sensory Formation in the Jag1-cko Vestibule
(A and B, D and E, G and H) Alternate sections demonstrating either JAG1 or SOX2 expression in the vestibular regions of control inner ears.
(C, F, I) Similar sections through the Jag1-cko inner ear demonstrating SOX2 expression. Dotted lines indicate regions where the cristae and ampullae are missing in the Jag1-cko inner ear. ac, anterior cristae; lc, lateral cristae; pc, posterior cristae; sac, saccular macula; ut, utricular macula.
Discussion
We have demonstrated that Notch signaling, mediated by the JAG1 ligand, is required early in development for the formation of the sensory regions of the ear. By comparing expression of JAG1 to two markers of the prosensory domain, SOX2 and p27kip1, we have shown that JAG1 marks all prosensory regions of the ear from early time points (E12.5), but becomes down-regulated in the organ of Corti by E14.5, when the sensory progenitors exit the cell cycle and begin differentiating into hair cells and supporting cells. Both SOX2 and p27kip1 are down-regulated in the affected prosensory regions of the Jag1-cko inner ear, demonstrating that JAG1 is necessary for the development of early sensory progenitors in the inner ear.
Distinctive Patterns of Hair Cell Formation in Jag1-cko Inner Ears Suggest Progenitor Cell Numbers Are Reduced
One intriguing result from our studies was that the six sensory regions were not equally affected by the loss of Jag1 function. For example, in the Jag1-cko vestibular system, the cristae were lacking altogether, and only a small number of hair cells differentiated in the utricular maculae. In contrast, the saccular maculae exhibited little disturbance in hair cell formation, although the overall shape of the organ was abnormal. In the Jag1-cko cochlea, hair cell differentiation patterns varied based on their apical or basal location. For example, in the apical regions of the cochlea only inner hair cells formed, and these were often arranged in multiple rows rather than the normal single row. In the middle and midbasal turns of the cochlea, patches of hair cells with nonsensory intervening regions were frequently observed. Within these patches, outer hair cells were sometimes present, although the patterning was abnormal and S100A1-labeled Dieter's cells were not present. In the very basal regions of the cochlea, neither hair cells nor supporting cells were present.
The patches of hair cells found in the basal regions of the cochlea and the differential effect of the mutation on the basal and apical portions of the cochlea were particularly interesting, as similar defects have been found in at least two other mouse mutants of genes known to play a role in the generation of the sensory precursors of the ear. For example, both a hypomorphic allele and a conditionally deleted allele of the Fgfr1 gene exhibited patches of hair cells in portions of the cochlea [31]. Similar to the Jag1-cko phenotype, these patches in the Fgfr1 conditional mutants contained mostly inner hair cells that were often arranged in multiple rows, with very few outer hair cells. Unlike the Jag1-cko phenotype, Fgfr1 function was required only in the cochlea. Another mouse mutant, a hypomorphic allele of the Sox2 gene (yellow submarine; Sox2ysb), also displayed patches of hair cells in the basal portions of the cochlea and a milder phenotype in the apical regions of the cochlea [25]. More similar to the Jag1-cko phenotype, SOX2 was required for both the cochlear and vestibular sensory regions [25]. The finding that primarily inner hair cells differentiate in these mutant cochleae may be due to the fact that inner hair cells are the first to differentiate [10,37], suggesting that if there are reduced numbers of progenitor cells, they would likely differentiate as inner rather than outer hair cells. Similarly, the milder phenotype in the apical regions of these mutants may be due to the fact that cells exit the cell cycle earliest in the apex [36]. This may mean that, if there are reduced numbers of progenitor cells, they would reside in the apical rather than the basal portions of the cochlea.
The multiple rows of inner hair cells observed in Jag1-cko and other mutants could be explained by a number of different scenarios. One possibility is that the multiple rows are not a result of actual increases in inner hair cell numbers, but rather are caused by defects in their eventual arrangement due to the shorter Jag1-cko cochlea. Recent studies of mouse mutants with defects in planar cell polarity and convergent extension (a term referring to the intercalation of cells, leading to growth of tissue in one dimension in the absence of proliferation) indicate that multiple rows of hair cells can be obtained in this way and are frequently observed in the apical regions of the cochlea [38–41]. An alternative possibility is that the multiple rows are a result of a second, later function of the JAG1 ligand, distinct from its prosensory role described here. A third possibility is that the down-regulation of p27kip1, a protein that inhibits continued proliferation of the precursor cells in the cochlea, leads to continued cell division of the remaining sensory progenitors, ultimately resulting in excess numbers of inner hair cells in the regions where they form. Taken together, these data suggest that sensory progenitors are reduced in the Jag1-cko inner ears and that Notch signaling, fibroblast growth factor signaling, and the transcription factor SOX2 all act in either common or parallel pathways involved in the production of sensory progenitors in the inner ear.
A Prosensory Role for Notch in the Ear
An examination of early prosensory markers, including p27Kip1 and SOX2, demonstrated that prosensory establishment is disrupted in Jag1-cko inner ears, consistent with the suggestion that progenitors are reduced in these mutants. Our data show that JAG1 plays an early prosensory role in ear development, quite unlike the role played later during development by the other Notch ligands, DLL1 and JAG2, which are involved in lateral signaling and differentiation [10,15]. These data are consistent with an early role for Notch signaling in progenitor cell maintenance in the inner ear. In a number of other systems, including the nervous system and more recently in the intestinal epithelium, it has been demonstrated that Notch signaling is involved in maintaining cells in an undifferentiated state [42–46]. In the mammalian nervous system it has been shown that loss of Notch signaling leads to premature differentiation and a reduction in the progenitor pool [42]. Consistent with these findings, in vitro studies have demonstrated that the frequency of neurosphere production was reduced in Notch signaling mutants [47,48], indicating a loss of stem cell potential. Moreover, studies have also shown that Notch signaling promotes radial glial identity, a cell type that has been shown to act as a progenitor cell in the central nervous system [49–52]. Our results suggest that, similar to the nervous system, Notch signaling via JAG1 is important for sensory precursor formation or maintenance in the inner ear. However, unlike the nervous system, we see no evidence for precocious differentiation, suggesting instead that JAG1 may affect the specification, survival, or proliferative capacity of the sensory precursors.
Recent evidence from the chick indicates that JAG1 may be important for the initial sensory specification events. By expressing a constitutively active form of Notch (Notch1-ICD), Daudet et al. [14] demonstrated that ectopic sensory patches could be induced, indicating that early Notch signaling may be important for the induction of sensory areas, and not just for their maintenance. However, it should be noted that ectopic sensory areas formed only in certain areas of the ear, indicating that some sensory competence is required for this effect. A similar result was obtained by overexpressing an activated form of β-catenin, an essential component of the canonical Wnt signaling pathway, in the chicken inner ear [53]. As in the Notch1-ICD studies, ectopic sensory regions were obtained, but again, only in certain regions of the ear. However, unlike the Notch gain-of-function studies, overexpression of β-catenin also led to a change in sensory region character (i.e., cochlear to vestibular), indicating that Wnt signaling governs not only whether a sensory region will form but also the type of sensory region that will form. In Drosophila, interactions between Notch and Wingless, a member of the Wnt family of signaling molecules, are well established [54,55], and evidence of an interaction has begun accumulating in vertebrates as well [45,56,57]. Bone morphogenetic protein (BMP) signaling may also be important for sensory formation, particularly for the sensory cristae, as BMP4 has been shown to mark the mouse cristae from very early in development [58]. Experiments in the chicken have shown that blocking BMP signaling sometimes leads to disturbances in sensory development [59]. Taken together, these data indicate that, based on expression patterns, previous studies, and the evidence presented here, JAG1 is the ligand responsible for the prosensory function of the Notch pathway in the ear. Furthermore, the Notch pathway likely interacts with other signaling pathways such as the Wnt, FGF, and BMP pathways to create sensory organs of the proper size, organization and character.
Sensory Formation Still Occurs in Jag1-cko Inner Ears
One somewhat puzzling question is that, if JAG1 is important for sensory progenitor development, why does any sensory formation occur in Jag1-cko inner ears? One possibility is that another Notch ligand is compensating for the loss of JAG1 function. This explanation seems unlikely since none of the other Notch ligands shows a similar expression pattern to JAG1 in the ear. For example, both the Dll1 and Jag2 genes are expressed in nascent hair cells after they exit the cell cycle and begin differentiating. However, in addition to hair cell expression, there is also early expression of the Dll1 gene in the anteroventral portion of the otocyst at about E10.5 [4,20], that likely overlaps with at least part of the JAG1 domain (see Figure 2) [60]. This expression domain has previously been thought to be related to the formation of the neuroblasts that delaminate from the otic epithelium and later differentiate into the neurons that will innervate the hair cells [4]. It has been shown in zebrafish that correct neuroblast formation requires Notch-mediated lateral signaling [9]; however, in mammals it has not been shown definitively that this is the role that the Dll1 gene plays at early stages. This leaves open the possibility that this early domain of DLL1 expression may be at least partially involved in prosensory specification, similar to the JAG1 expression domain.
Nonsensory Defects in Jag1-cko Inner Ears
In addition to the defects in sensory formation in the Jag1-cko inner ears, the mutant inner ears also exhibited nonsensory defects. Specifically, the semicircular canals were largely absent, with the exception of portions of the anterior and lateral canals. In addition, all three ampullae were absent, the utricle was small, and the cochlea was undercoiled. Based on recent studies, it is likely that these defects are secondary to the sensory defects. For example, it has been shown that FGFs expressed in the sensory cristae promote semicircular canal formation through up-regulation of BMP2 [61]. Thus, loss of the cristae would be expected to have a severe affect on canal formation. Emerging evidence from mouse mutants has demonstrated that genes involved in sensory formation result in severely malformed inner ears. For example, mutations in the Sox2 gene lead to malformations very similar to those described here in Jag1-cko mutants. The inner ears of embryos homozygous for two different mutant alleles of Sox2, Sox2lcc/lcc and Sox2ysb/ysb, showed disrupted canal formation; smaller utricular and saccular compartments; and thinner, undercoiled cochleae [25]. In addition, FGF10 mouse knockouts also showed disrupted cristae development associated with loss of canal structures [62]. However, unlike the canal structures, cochlear formation does not appear to be strictly dependent on development of the organ of Corti, as a cochlea, albeit short and thin, will form in the absence of any sensory formation [25]. However, normal cochlear length appears to be dependent on sensory formation, at least partially through convergent extension. Recently, a number of genes have been found in the cochlea that lead to defects in planar cell polarity as well as a shortened cochlea, presumably because of defects in convergent extension [38,41]. Therefore it is likely that the shortened cochlea observed in Jag1-cko mutants is at least partially a result of failure of convergent extension caused by a reduction in the number of sensory precursors.
The data presented here demonstrate that the Jag1 gene is required for sensory precursor development in the inner ear. Further studies are required to establish the exact role that JAG1-mediated Notch signaling plays in early sensory progenitors, and also its relationship to the roles played by FGF signaling and SOX2 expression. Understanding how the sensory precursors form is an important prerequisite for regeneration studies that may provide molecular tools to treat hearing loss and vestibular disorders [63].
Materials and Methods
Construction of the Jag1floxneo allele.
To construct the Jag1floxneo allele, bacterial artificial chromosome clones containing the Jag1 genomic locus were isolated from a RPCI-22 (129S6/SvEvTac) mouse bacterial artificial chromosome library (filters obtained from Research Genetics) by hybridization to a 1.8-kb mouse Jag1 cDNA probe. To make the shorter 5′ homology region of the targeting vector, a 2.2-kb KpnI fragment upstream of exon 4 was isolated, blunt-ended, and subcloned into the SmaI site of a modified pBS vector that contained a loxP-FRT-PGKneo-FRT cassette. A 1.5-kb KpnI fragment that contained exon 4 was also subcloned into the loxP-FRT-PGKneo-FRT cassette. To construct the longer 3′ homology region, a 3.5-kb KpnI-SmaI fragment containing exon 5 was blunt-ended and subcloned into the EcoRV site of another modified pBS vector that contained a single loxP site. A 3.5-kb SmaI-SalI fragment from this construct was then cloned into the SmaI-XhoI site of a pKO 905 vector containing a diphtheria toxin gene for negative selection. A 5.7-kb SalI-NotI fragment from the loxP-FRT-PGKneo-FRT construct was then cloned into the SalI-NotI sites of the pKO 905 vector containing the 3′ homology region to generate the final Jag1floxneo targeting vector (see Figure 1).
Generation of Jag1flox mice.
The Jag1floxneo targeting construct was linearized with Not1 and electroporated into CJ7 embryonic stem (ES) cells, as described previously [64]. DNA from 288 ES cell clones was screened by PCR using an internal/external primer set, and positive clones were then confirmed by Southern blot by probing EcoRI-digested DNA with an external 1.7-kb Stu1-EcoRI fragment located 3′ to the targeting construct (see Figure 1). This probe also detected partial recombination events in which the distal loxP site was lost; in these cases a slightly larger fragment (11 kb rather than 9.3 kb) was obtained (see Figure 1A). The presence of the distal loxP site was further confirmed by PCR using primers that flanked the loxP site (DSLF and J1LoxR1; see below for sequences). Correctly targeted clones were injected into C57BL/6J (B6) blastocysts, and chimeric mice were obtained. Chimeric male mice were mated to B6 females and the agouti progeny were assayed for the presence of the Jag1floxneo allele by PCR using Jag1floxneo specific primers. Jag1floxneo/+ mice were intercrossed to create homozygous Jag1floxneo/Jag1floxneo offspring. Homozygous Jag1floxneo/Jag1floxneo mice appeared normal and healthy, suggesting that the neomycin resistance cassette (PGKneo) did not adversely affect Jag1 expression. To control for any possible effects from the presence of the PGKneo cassette, the FRT-flanked PGKneo cassette was deleted by mating Jag1floxneo mice to FLPe-recombinase expressing mice (Gt[ROSA]26Sor tm1(FLP1)Dym; Jackson Laboratory, Bar Harbor, Maine, United States) to produce Jag1flox mice. Both Jag1floxneo and Jag1flox mice were used in these experiments, and no differences in the resulting phenotype were observed. To differentiate the deleted form of this allele from our previously reported Jag1 null mutant allele (Jag1del1) [35], we designate the Jag1 allele generated by Cre recombinase-mediated deletion of the Jag1flox or Jag1floxneo alleles the Jag1del2 allele.
Mouse husbandry and genotyping.
Foxg1-Cre mice ([30]; gift of Rob Burgess) were maintained on an outbred Swiss Webster background. ZP3-Cre mice ([34]; gift of Mimi de Vries and Barbara Knowles) were maintained on a B6 background. Typically, males that were heterozygous for both a Foxg1-Cre allele and our previously constructed Jag1 null allele (Jag1del1) [35] (maintained on a B6 background), were crossed to Jag1flox/Jag1flox females that were maintained on a B6/129 background. Mice of the genotypes Foxg1-Cre/+; Jag1del1/Jag1flox and Foxg1-Cre/+; Jag1del1/Jag1floxneo were used interchangeably, and are designated as Jag1-cko mice in this report.
To genotype the Jag1flox mice, the primers used were: DSLF, 5′-TCAGGCATGATAAACCCTAGC-3′ (forward) and J1LoxR1, 5′-CTACATACAGCATCTACATGC-3′ (reverse); these primers flank the 5′ LoxP site. To genotype for CRE-mediated recombination, a primer upstream of the 3′ LoxP site was used: J1FlpF1, 5′-CAGGTTGAGTGCTGACTTAG-3′, along with the J1LoxR1 reverse primer. To genotype for the Foxg1-Cre and ZP3-Cre alleles, Cre-specific primers were used: Cre1, 5′-TGATGAGGTTCGCAAGAACC-3′ (forward) and Cre2, 5′-CCATGAGTGAACGAACCTGG-3′ (reverse). Jag1del1 primers were as follows for the mutant: JGKO1, 5′-TCTCACTCAGGCATGATAAACC-3′ (forward) and SOL1, 5′-TGGATGTGGAATGTGTGCGAG-3′ (reverse). A different reverse primer, JGKO2, 5′-TAACGGGGACTCCGG ACAGGG-3′ was used to detect the wild-type allele. Littermates (wild type, Jag1del1/+ or Jag1flox/+) were used as controls for all experiments.
Paintfilling and scanning electron microscopy.
The paintfilling of the Jag1-cko inner ears was performed at E15.5. The technique was performed as previously described [28]. Inner ears were prepared for SEM as described previously using a version of the osmium tetroxide-thiocarbohydrazide method [65]. Specimens were examined with a Hitachi 3000N scanning electron microscope (Hitachi, Tokyo, Japan).
Immunohistochemistry and lectin staining.
For immunohistochemistry, embryonic heads were bisected and fixed for 1–2 h in 4% paraformaldehyde in PBS. Half heads were embedded in paraffin wax, and immunocytochemistry was performed on standard 7-micron sections. Antibodies used included anti-Myo7a (1:1,000; gift of A. EL-Amraoui and C. Petit), anti-p27kip1 (1:100; Neomarkers, Stratech, Soham, Cambridgeshire, United Kingdom), anti-SOX2 (1:2,000; Chemicon, Temecula, California, United States; AB5603), anti-JAG1 (1:100; Santa Cruz Biotechnology, Santa Cruz, California, United States; H-114) and anti-S100A1 (1:50; Dako, Glostrup, Denmark). For all antibodies used, an antigen retrieval step was performed by boiling the sections for 10 min in 10 mM citric acid. Secondary antibodies used were either Alexa-Fluor 488 or 546 goat anti-mouse or rabbit (1:400; Invitrogen, Carlsbad, California, United States). Slides were coverslipped in Vectashield HardSet Mounting Medium with DAPI (Vector Laboratories, Burlingame, California, United States). Lectin staining was performed using the Griffonia simplifonia I lectin (Vector Laboratories) essentially as described [15].
In situ hybridization and RT-PCR.
Since the Jag1del2 mutant allele creates an in-frame deletion of the DSL domain, an in situ probe was designed for detection of the floxed region of the Jag1flox allele by in situ hybridization. The probe was created by amplifying a 433-bp product encompassing exon 4 (primers: J1-420F, 5′-CGACCGTAATCGCATCGTAC-3′ and J1-853R, 5′-ATGCACTTGTCGCAGTACAG-3′) and subcloning the product into the PCR II vector (Invitrogen). For whole-mount embryos in situ, embryos were fixed overnight in 4% paraformaldehyde. In situ hybridization was performed essentially as described [66]. For cochlear in situ, inner ears were dissected from the head and fixed overnight in 4% paraformaldehyde. After washing in PBS, the bony shell and stria were removed from the cochleae and the samples were dehydrated in methanol. In situ hybridization was performed as described [67], with the exception of the posthybridization washes, which were done according to [68]. For RT-PCR, total RNA was extracted from the E10.5 control and Jag1del2/Jag1del2 embryos, using an RNAeasy kit (Qiagen, Valencia, California, United States) and following the manufacturer's instructions. First-strand cDNA synthesis was performed using the AMV reverse transcriptase (Promega, Madison, Wisconsin, United States) with specific primer J1-961R (5′-AGTCCCACAGTAATTCAGATC-3′). Products were amplified from cDNA using primers that flanked exon 4 (J1-420F, 5′-CGACCGTAATCGCATCGTAC-3′ and J1-961R).
Acknowledgements
We thank Drs. A. El-Amraoui, C. Petit, R. Burgess, M. de Vries, and B. Knowles for reagents; Peter Finger of the Jackson Laboratory Histopathology and Microscopy Services for help with the SEM processing; and the Jackson Laboratory Cell Biology and Microinjection Core Facility for the blastocyst injections. This work was supported by grants from the National Institutes of Health to AEK (DC05865) and TG (NS036437 and DK066387), and from the Jackson Laboratory (CA034196).
Abbreviations
BMP - bone morphogenetic protein
cko - conditional knockout
DLL - Delta-like
DSL - Delta-Serrate-Lag2
E[number] - embryonic day [number]
ES - embryonic stem
FGF - fibroblast growth factor
JAG - Jagged
SEM - scanning electron microscopy
Footnotes
Author contributions. AEK and TG conceived and designed experiments. AEK and JX performed experiments. All authors analyzed the data. AEK wrote the paper.
Competing interests. The authors have declared that no competing interests exist.
A previous version of this article appeared as an Early Online Release on December 1, 2005 (DOI: 10.1371/journal.pgen.0020004.eor).
Citation: Kiernan AE, Xu J, Gridley T (2006) The Notch ligand JAG1 is required for sensory progenitor development in the mammalian inner ear. PLoS Genet 2(1): e4. | [
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16517939 | An isoform of ZBP-89 predisposes the colon to colitis
Abstract
Alternative splicing enables expression of functionally diverse protein isoforms. The structural and functional complexity of zinc-finger transcription factor ZBP-89 suggests that it may be among the class of alternatively spliced genes. We identified a human ZBP-89 splice isoform (ZBP-89ΔN), which lacks amino terminal residues 1–127 of the full-length protein (ZBP-89FL). ZBP-89ΔN mRNA was co-expressed with its ZBP-89FL cognate in gastrointestinal cell lines and tissues. Similarly, ZBP-89ΔN protein was expressed. To define its function in vivo, we generated ZBP-89ΔN knock-in mice by targeting exon 4 that encodes the amino terminus. Homozygous ZBP-89ΔN mice, expressing only ZBP-89ΔN protein, experienced growth delay, reduced viability and increased susceptibility to dextran sodium sulfate colitis. We conclude that ZBP-89ΔN antagonizes ZBP-89FL function and that over-expression of the truncated isoform disrupts gastrointestinal homeostasis.
INTRODUCTION
Alternative pre-mRNA splicing and multiple promoter usage are common mechanisms for increasing genetic complexity in humans (1) and mice (2). Genome-wide analyses indicate that the majority of human genes express alternative splice isoforms (3) and some variants contribute to neoplasia or other disease processes (4,5). For example, truncated isoforms of the p53 gene family, including p63 (6) and p73 (7–9), oppose the tumor suppressor activity of their full-length cognates and are over-expressed in tumors.
ZBP-89 (ZNF148; Zfp148; BFCOL1), a Krüppel-type zinc-finger protein (10), is both structurally (11) and functionally complex (12–14). It regulates diverse biological functions through direct promoter binding (10,14) and through multiple protein–protein interactions. It interacts directly with the tumor suppressor p53 through its zinc-finger domain (13) and indirectly with the histone acetyltransferase and transcriptional co-activator p300 through its amino terminus (12). ZBP-89 induces cell growth arrest through a p53-dependent mechanism (13) and apoptosis through a p53-independent mechanism (12). Mice haploinsufficient for ZBP-89 (Zfp148) are sterile owing to aberrant spermatogenesis (15). In addition, embryonic stem cells harboring a single ZBP-89 allele fail to exhibit p53 phosphorylation at Ser 15 (15).
ZBP-89 forms a complex with p300 during butyrate induction of p21waf1 in human colorectal cell lines (12). The amino terminus of ZBP-89 contains an acidic domain that is required for p300-mediated induction, since an expression construct lacking this domain (Δ amino acids 1–113) loses its ability to enhance butyrate induction of p21waf1 (12). The acidic domain of ZBP-89 is contained entirely within exon 4 of the human and mouse genes (11). Given the heterogeneous ZBP-89 mRNA expression pattern (10), the functional importance of the p300 interaction domain, and the fact that the ZBP-89 null mouse is likely an embryonic lethal (15), we generated a mouse homozygous for the ZBP-89ΔN isoform. Homozygous ΔNter mice experienced growth delay, reduced viability and increased susceptibility to Dextran sodium sulfate (DSS) induced colitis, suggesting that over-expression of the ΔNter isoform disrupts normal gastrointestinal homeostasis.
MATERIALS AND METHODS
Database deposition
National Center for Biotechnology Information (NCBI) reference sequences for ZBP-89 genomic loci are Homo sapiens chromosome 3 locus NC_000003 and BAC RP11-775J23 (AC108688); Mus musculus chromosome 16 contig NT_039624 and Pan troglodytes chromosome 3 genomic locus NW_104931. The human ZBP-89ΔN mRNA and exon 4B genomic sequences reported here are found in GenBank as entries DQ090088 and DQ090089, respectively. The P.troglodytes genomic sequence homologous to human exon 4B is deposited in Genbank DQ144540.
Organization of the human (ZNF148), chimp and mouse (Zfp148) ZBP-89 loci
A human genomic clone contig encompassing the ZBP-89 locus was previously described (14). Bacterial artificial chromosome (BAC) clones were used as templates to sequence intron/exon boundaries spanning the ZNF148 locus. Exon 4B was identified by gene prediction sequence analysis (16,17). Similarly, a BAC and bacteriophage lambda clone contig spanning the mouse Zfp148 locus was assembled and sequenced to determine the genomic organization of the Zfp148 locus. Primers Ptr-4B-F: 5′-TTCACCTCCCTGTCCTGTTC-3′ and Ptr-4B-R: 5′-TATCTGTCCCGTTTGCCTG-3′ were used to amplify and sequence chimp (P.troglodytes) genomic DNA containing exon 4B, after BLAST analysis (18) had revealed a sequence gap in this region. Chimp genomic DNA was obtained from the Integrated Primate Biomaterials and Information Resource through the Coriell Institute for Medical Research (Camden, NJ).
RT–PCR analysis
Total cellular RNA was isolated from cultured cells using TRIzol reagent (Invitrogen, Carlsbad, CA). Archived whole cell RNA samples from paired colon cancer and normal mucosal samples were also analyzed. Two-step RT–PCR was performed using SuperScript III RT (Invitrogen), however reverse transcription was performed at 55°C in order to obtain adequate yields of human exon 4B-containing cDNA. RT reactions were primed with random nonamers. Human primer sequences (5′ to 3′) were 4A-F: ATGAACATTGACGACAAACTGGAAG; 5-R: TTCCATATGATATTTTTGTATGAAT; 4B-F: TAGGGATGGTCAGCACTG; 6-R: GTTCTTTTGTGCCTTTCC. Mouse primer sequences (5′ to 3′) included Ex2-F: GCGGATAGAAGAGAAGAATCAGTGG; Ex4-F: CATTGACGACAAACTGGAAGG; Ex4-R: ACTTCGATCTTGAAGTACTGACTC; Ex5-R: CAGGAGAGCGTTGTTTCCG; Ex3/5fusion-F: CAGCCTCAGATAAGTGTA and Ex9-R: TTGTGGCATCTGGTGAAG. RT–PCR products were purified and subjected to DNA sequence analysis (University of Michigan DNA Sequencing Core).
5′-RACE of human exon 4B
The 5′ end of the exon 4B variant was identified using the GeneRacer™ system (Invitrogen, Carlsbad, CA), gene specific primer (5′ to 3′) 4B211-Rev: CAGTGCTGACCATCCCTATC CTACTTG and 2 µg of Jurkat whole cell RNA. Nested PCR products, generated with adaptor primers included with the GeneRacer™ system, were cloned using the TOPO-TA system (Invitrogen, Carlsbad, CA), and sequenced.
Targeting vector
We constructed vector pΔEx-4 to replace ZBP-89 exon 4 with a PGK-Neo cassette by homologous recombination. High fidelity long-range PCR was used to amplify targeting arms from mouse BAC clone pBmZBP-89, using primers (5′ to 3′): (Kpn)Int3-F, GATAGGTACCGCATTGGATGGCACAAGTGACTGAGAGG with (Xho)Int3-R, CTCGAGCCCGGGCTTAAGTATAACTGCCTAGAAAG for the left arm and (Apa)Int4-F, CTCGAGGGGCCCGTAAGTACTAAACTAGAAATG with (Apa)Int4-R, CTCGAGGGGCCCAAGAGCCTTGCTGACTCATAG for the right arm. The left targeting arm, encompassing exon 3 and intron 3, was 8 kb in length. The right targeting arm consisted of the proximal 2 kb of intron 4. The neomycin cassette, including a transcriptional stop signal, was isolated from pPNT (19), generously provided by Dr Richard Mulligan.
Generation of targeted ES cells and ΔNter mice
Electroporation of embryonic stem (ES) cells with the pΔEx-4 targeting vector and microinjection of blastocysts with targeted ES cells were performed by the University of Michigan Transgenic Animal Model Core (). Genomic PCR was utilized to genotype targeted 129 Sv/J ES clones, chimeric founders, and progeny resulting after germline transmission. Primer sequences (5′ to 3′) were Int3-F1, GGAGTATTCTCTGTCCGTT ATG; Int4-326R, GCAAGAACTACACAGAGAAACCAC and R506Neo, TGAGGAA GAGGAGAACAGCG.
Two-dimensional western blot analysis
Whole cell protein extracts were prepared from mouse spleen using T-PER tissue protein extraction reagent (Pierce, Rockford, IL), supplemented with Complete Mini protease inhibitors (Roche, Indianapolis, IN) according the manufacturers' recommendations. Whole cell protein extracts from human Jurkat cells were similarly prepared using M-PER mammalian protein extraction reagent (Pierce, Rockford, IL). Isoelectric focusing (IEF) was performed with 7 cm ZOOM® Strip ph3-10L (linear) immobilized pH gradient gels (Invitrogen, Carlsbad, CA), as recommended by the manufacturer. The focused proteins were then separated on NuPAGE® 4-12% acrylamide gels (Invitrogen). Electroblot transfer to PVDF membrane and immunoblot procedures with ZBP-89 antiserum are as previously described (10,12).
Dextran sodium sulfate colitis
DSS colitis was induced in 6–9 month old ZBP-89ΔN and littermate control mice by the addition of 4% DSS to drinking water for a period of 5 days (20). Treated mice were returned to normal drinking water for 2 days prior to necropsy for histopathological scoring. Hematoxylin and eosin (H&E) stained colon sections were prepared by the Swiss roll method (21) and were scored for colitis index by a modification of a previously described method (22). Briefly, crypt damage was scored from 0 to 3 as none, basal only, moderate damage and complete erosion, respectively. Inflammation was scored from 0 to 3 as none, minor, moderate and severe leukocyte infiltration. Submucosal edema was scored from 0 to 3 as none, minor, moderate and severe, respectively. Similarly, hemorrhage was scored from 0 to 3 as none, minor, moderate and severe. Each parameter was multiplied by an extent factor (1–3), <10%, up to 25%, 25 to 50% and >50%, respectively. Samples with transmural involvement received an additional four points. Therefore, the maximum possible colitis index score was 40. Animals that died during treatment because of colitis injury were also given the maximum score. Individuals scoring the samples were blinded to the genotype.
Cell culture
The following cell lines were obtained from ATCC (Manassas, VA) and maintained on the recommended growth media: Human gastric epithelial cells Kato III and MKN45; human colon cancer cells Colo 320DM, CaCo-2 and HCT116; and human Jurkat leukemia cells. Cells were cultured in a humidified atmosphere of 5% CO2 and 95% air at 37°C. All culture media were supplemented with penicillin G (100 U/ml) and streptomycin (100 µg/ml).
RESULTS
Identification of human ZBP-89ΔN isoform
To determine if isoforms of ZBP-89 exist, we screened the human ZNF148 genomic locus in silico (16,17). After localizing candidate alternative exon 4B within 4 kb upstream of exon 5 (Figure 1A), we used RT–PCR analysis to show that exon 4B is expressed (Figure 1B). Exon 4B mRNA (ΔN) was co-expressed with exon 4A-containing message (FL) in colon cancer cells (ColoDM2, CaCo2, HCT116), and in primary tissue from normal colon and colon adenocarcinoma. Both forms also were abundantly expressed in Jurkat cells (data not shown). This suggested that at least two forms of ZBP-89 exist.
To better understand the function of the isoform, exon 4B-containing cDNA was sequenced. In addition to exon 4B, the variant mRNA included exons 5 and 6 (Figure 2A), as well as exons 7–9 (not shown). In contrast, RT–PCR with forward primers from exons 1-4A failed to generate products with exon 4B antisense primers, suggesting that an independent promoter regulates exon 4B expression. This was confirmed by 5′-rapid amplification of cDNA ends (RACE), which showed that transcription was initiated immediately upstream of exon 4B. The cDNA sequence showed that exon 4B was spliced to exon 5 resulting in an alternative reading frame relative to the cDNA encoded by exon 4A (Figure 2A). Exon 4B was 329 nt in length and composed of untranslated sequences when fused to exon 5. These data predicted that alternative promoter usage upstream of exon 4B resulted in the expression of an amino-terminally truncated ZBP-89 isoform, ZBP-89ΔN, with an alternative initiation codon corresponding to M128 of full-length mRNA (Figure 2B). This isoform lacks the acidic domain and p300-interaction region (12) found in full-length (ZBP-89FL) protein (10). Identical results were obtained with cDNA derived from esophagus, stomach, colon and Jurkat T-cells, suggesting that the exon 4B alternative promoter mechanism is common.
Detection of human ZBP-89ΔN protein
We found that the electrophoretic mobility of ZBP-89ΔN protein, despite its shorter length, overlapped with the mobility of its ZBP-89FL cognate (Figure 2C). Both forms, isolated from Jurkat cells, migrated at 100 kDa on a 4–20% gradient gel. Similar SDS–PAGE anomalies have been reported with other proteins, including CTCF (23) and XPA (24). An alternative approach to separate the protein isoforms was suggested by comparing their predicted (25) isoelectric points (pI). Loss of the acidic domain in ZBP-89ΔN protein predicted a pI of 7.8, compared to 6.0 for ZBP-89FL. This difference could be revealed by two-dimensional (2D) gel electrophoresis, followed by western blot analysis (Figure 2C). ZBP-89 antiserum detected two protein species, with apparent electrophoretic mobilities of 100 kDa, but with a pI difference of ∼1.5, confirming that the more basic form is ZBP-89ΔN.
ZBP-89ΔN alternative promoter mechanism is restricted to hominids
Although the human (ZNF148) and mouse (Zfp148) ZBP-89 loci share many features of genomic organization (11), exon 4B is absent in mice and most other mammals (data not shown). In contrast, we identified a 133 bp segment of chimpanzee (P.troglodytes) chromosome 3 sequence with striking DNA sequence homology to human exon 4B, but there was a gap in the existing online genomic sequence. Sequencing across the gap after genomic DNA amplification, we found that the chimp shares 99% DNA sequence identity to the 329 bp human exon 4B. This suggested that the ZBP-89ΔN alternative promoter mechanism is restricted to hominids.
Generating a mouse model of ZBP-89ΔN expression
Since mice lack exon 4B, and therefore do not express a ZBP-89ΔN isoform, we targeted the mouse exon 4 domain by homologous recombination (Figure 3A). The left targeting arm contained exon 3 and intron 3, and the right targeting arm contained 1.8 kb of the 5′ margin of intron 4. Homologous recombination occurred in 4 of 1200 ES clones identified by PCR (Figure 3B). Of the four homologous recombinants, three demonstrated stable karyotypes and were therefore injected into blastocysts. Germline transmission of the exon 4-targeted locus resulted in two founder lines, 4E10 and 8C6, while a third founder, 9C6, failed to transmit the recombinant locus. Germline transmission was obtained first in the 4E10 lineage. Near-Mendelian ratios were observed at the age of weaning (3 weeks) in offspring of heterozygous intercrosses (Figure 3C). The data suggested that ∼20% of Δexon4 homozygous embryos die in utero or perinatally, however the difference between predicted and observed ΔExon4/ΔExon4 yields was not statistically significant (χ2-square analysis; P > 0.05). A similar pattern was observed in the 8C6 lineage. These results also suggest that the mutant locus was at least partially functional, since heterozygosity for a null ZBP-89 (Zfp148) allele results in failed germ cell development and therefore is not transmitted (15).
Alternative splicing of exon 3 to exon 5 in recombinant mice
To determine the effect of exon 4 deletion on ZBP-89 expression, we analyzed cDNA derived from normal, heterozygous and exon 4-targeted animals (Figure 4A). Whereas exon 4 sequences were absent from recombinant mRNA, both upstream and downstream exons were expressed. RT–PCR with primers spanning exon 4 generated a 690 bp cDNA from wild type and a 339 bp cDNA from mutant alleles. The size difference between WT and targeted cDNA was 351 bp, the size of exon 4. This suggested that exon 3 was spliced directly to exon 5, excluding the intervening Neo cassette, and this was confirmed by DNA sequencing (Figure 4B). Deletion of exon 4 removed the initiation codon found in FL message and also resulted in an alternative reading frame. Gene feature sequence analysis (16,17) predicted that the alternative initiation codon corresponded to M128 of the FL protein (Figure 4B), analogous to the human ZBP-89ΔN variant (Figure 2B).
Mouse ZBP-89ΔN protein expression
2D western blot analysis (Figure 4C) showed that ZBP-89FL and ZBP-89ΔN proteins are expressed in an allele-dependent manner. The two forms have similar electrophoretic mobilities of 100 kDa, but are readily separated by their pI differences, similar to the human isoforms (Figure 2C). As mentioned above, similar electrophoretic anomalies have been observed with other protein isoforms (23,24). The immunoblot data also suggest that the ΔN protein isoform may be present at higher levels than the FL form in spleen tissue, however mRNA levels appear comparable, suggesting a possible difference in protein stability. Collectively, the mRNA and protein expression data confirmed that we knocked the ZBP-89ΔN variant into the mouse genome. Thus, the mouse model offers the advantage of assessing the biological effect of ZBP-89ΔN expression exclusively.
ZBP-89ΔN/ΔN mice experience growth delay and reduced viability
At weaning, by age 3 weeks, male ZBP-89ΔN/ΔN mice weighed an average of 5.9 ± 0.5 g (Figure 5A), 34% smaller than ZBP-89FL/FL (12.1 ± 0.7 g) and 49% smaller than ZBP-89FL/ΔN heterozygous littermates (15.1 ± 0.8 g). From ages 4–8 weeks, the size differential between ZBP-89ΔN/ΔN and control mice was diminished. A similar effect was observed with female mice (data not shown). As shown above (Figure 3C), ZBP-89ΔN/ΔN mice experienced 20% perinatal mortality as determined by genotype ratios at weaning. Reduced viability persisted and became statistically significant (Figure 5B), with 50% mortality at 48 weeks and 69% mortality by 104 weeks. The 2 year survival ratios were 20:20 ZBP-89FL/FL, 39:40 ZBP-89FL/ΔN and 5:16 ZBP-89ΔN/ΔN mice. Histopathological organ and tissue surveys revealed no obvious abnormalities in ZBP-89ΔN/ΔN mice, with the possible exception of the colon, where we noted a trend toward slightly increased lymphocytic infiltrates (data not shown). Semi-quantitative RT–PCR analysis of colon mRNA levels (Figure 5C), suggested that FL and ΔN forms were expressed in an allele-dependent manner. Collectively, these data demonstrated that balanced expression of ZBP-89FL and ZBP-89ΔN isoforms, as observed in heterozygous mice, supports normal growth and viability. In contrast, exclusive expression of the ZBP-89ΔN isoform in transgenic mice resulted in growth delay and reduced viability.
ZBP-89ΔN confers DSS colitis susceptibility
We previously showed that the amino terminal region of ZBP-89 (amino acids 1–111) is required to potentiate butyrate induction of p21waf1, suggesting that the amino terminal region mediates the butyrate-dependent anti-proliferative activities of ZBP-89 in the mammalian gastrointestinal tract (12). Butyrate suppresses colonic inflammation when induced by DSS (26). Since DSS accentuates a tendency to develop colitis, we challenged ZBP-89ΔN/ΔN mice acutely with 4% DSS for 5 days, as previously described (20). Histopathogical analysis (hematoxylin and eosin stained sections) demonstrated the presence of severe colitis that correlated with a ZBP-89ΔN gene dosage effect (Figure 6A–C). ZBP-89FL/FL mice (Figure 6A) had localized areas of lymphocytic infiltration and minimal submucosal edema. ZBP-89FL/ΔN mice (Figure 6B) exhibited more extensive infiltration, with displaced normal crypt architecture in addition to submucosal edema. ZBP-89ΔN/ΔN mice (Figure 6C) had areas with complete erosion of crypt architecture, grossly visible hemorrhage and extensive submucosal edema. The ZBP-89ΔN gene dosage effect also correlated with the duration of gastrointestinal bleeding (Figure 6D). Moreover, only ZBP-89ΔN/ΔN mice died during DSS treatment, with 50% mortality by the conclusion of the 7 day regimen (Figure 6E). Composite colitis scoring similarly paralleled the ZBP-89ΔN gene dosage (Figure 6F). These data suggest that increased expression of ZBP-89ΔN is associated with increased susceptibility to colitis.
DISCUSSION
Through structural and functional analyses, we identified a human-specific ZBP-89 splice isoform, ZBP-89ΔN, which was generated by alternative promoter usage upstream of an alternative exon 4B. As a consequence, ZBP-89ΔN protein lacks a transcriptional domain (12) found in its full-length ZBP-89 cognate. This is the first characterization of a ZBP-89 isoform and is consistent with previous observations that gene complexity is lower in mice than in humans. As many as 59% of human genes are alternatively spliced (27), while the highest estimate to date for the mouse is 33% (28). Comparative genomic analysis suggested that the ZBP-89ΔN splicing mechanism has been conserved between humans and chimps, indicating that this specific mechanism of regulating ZBP-89 function is restricted to hominids.
Alternative mRNA expression increases genetic diversity through regulatory mechanisms that also are implicated in cancer (1,3–5,29) and other disease processes (30,31). The human ZBP-89ΔN variant described here renders the colonic mucosa more susceptible to injury. Our previous studies showed that the N-terminal domain is required for ZBP-89 to mediate butyrate-dependent activation of p21waf1 by cooperating with p300 (12). This result demonstrated that the N terminal domain is functionally important in vitro. Generating a mouse model in which only the truncated form was expressed revealed that loss of the p300-interacting domain results in delayed growth and a shortened lifespan. Although the overall effect on the health of the organism was impressive, an initial survey of several organs did not reveal obvious abnormalities, with the exception of the colon, which appeared to exhibit slightly more inflammatory infiltrates than the wild type mice. The propensity of the ZBP-89ΔN expressing mice to develop colitis was subsequently uncovered when the animals were challenged with DSS. Therefore, we concluded from these studies that expression of ZBP-89FL tends to protect against colitis. The gene dosage dependence of colitis susceptibility further suggests that the ZBP-89ΔN isoform functions as a dominant negative antagonist of ZBP-89FL. Similar antagonistic interactions have been reported between FL and ΔN isoforms within the p53 gene family, including ΔNp63 (29) and ΔNp73 (7–9).
The ZBP-89ΔN knock-in model reported here, along with an earlier Zfp148+/− model (15), help to elaborate how individual protein domains mediate the in vivo functions of ZBP-89. Mice heterozygous for a null ZBP-89 allele demonstrate complete failure of male germ cell development and are defective in p53-dependent embryogenesis (15). This finding suggests that the ability of ZBP-89 to interact with p53 is exquisitely sensitive to gene dosage. In contrast, homozygous ZBP-89ΔN expression is compatible with embryonic and postnatal survival, albeit at reduced levels. We previously showed that the DNA-binding, zinc-finger region of ZBP-89 mediates its interaction with p53 (13), and this domain is retained in the ZBP-89ΔN isoform. Therefore, the Nter domain, which includes the p300 interaction domain, is dispensable for p53-dependent embryonic development and postnatal survival, but is essential for normal gastrointestinal function.
Acknowledgements
The authors gratefully acknowledge the expertise of the University of Michigan Transgenic Animal Model Core, especially Thom Saunders, Linda Samuelson and Elizabeth Hughes. The authors also thank members of the DNA Sequencing Core and the Unit for Laboratory Animal Medicine at the University of Michigan, and Kathy McClinchey for assistance with histology. Proteomics data were provided by the Michigan Proteome Consortium () which is supported in part by funds from the Michigan Life Sciences Corridor. Specifically, valuable assistance with 2D gel electrophoresis was provided by Jennifer Callahan. This research is supported in part by the National Institutes of Health through the University of Michigan's Cancer Center Support Grant (5 P30 CA46592) and the University of Michigan Gastrointestinal Peptide Research Center (DK-34533). Art Tessier and Gail Kelsey provided administrative support. Stacey Ehrenberg provided technical assistance. This work was supported by grants from the National Institutes of Health to D.J.L. (R21 DK65004-01) and J.L.M. (RO1-DK55732). Funding to pay the Open Access publication charges for this article was provided by NIH grant RO1-DK55732 (J.L.M.).
Conflict of interest statement. None declared.
Figures and Tables
Figure 1
Identification of a novel exon within the human ZBP-89 (ZNF148) locus. (A) The human ZBP-89 (ZNF148) locus spans 142 kb (upper panel) and encompasses three untranslated (1–3) and six coding (4–9) exons. A potential alternative exon 4B (shaded box) was identified at a position 4.2 kb upstream of exon 5. Arrows indicate the locations of forward (F) and reverse (R) RT–PCR primers used for expression analysis of FL ZBP-89 and the exon 4B variant. Selected intron sizes (kb) are indicated. (B) RT–PCR analysis of full-length (FL) and variant (ΔN) ZBP-89 mRNA expression, using actin (Ac) as control. The cDNAs were generated with cultured cell lines using primers 4A-F/5-R (726 bp product); primers 4B-F/6-R (432 bp product) and (Ac), Actin control primers (400 bp product). Whole cell RNA was isolated from ColoDM2, CaCo 2, and HCT116 human colon cell lines as well as paired normal colon and colon adenocarcinoma tissue. Full-length message is expressed from promoter 1 (Pro 1) and variant (ΔN) ZBP-89 mRNA is expressed from promoter 2 (Pro 2), as described in Figure 2.
Figure 2
Variant ZBP-89 transcript encodes a truncated protein, ZBP-89ΔN. (A) The 5′ end of exon 4B-variant cDNA, determined by 5′-RACE, is shown. Underlined italics, exon 4B-encoded; lower case italics, exon 5 untranslated (frameshift relative to ZBP-89FL cDNA); upper case italics, exon 5 coding sequence commencing at M128 relative to FL protein; upper case underlined, beginning of exon 6-encoded. The remainder of the cDNA and amino acid sequences are identical to the corresponding segments of FL cDNA and protein. (B) Comparison of FL (upper) and ΔN (lower) protein isoforms. The amino terminus of ZBP-89FL includes an acidic and p300-interaction domain (PID), which is absent in ZBP-89ΔN. The ZBP-89ΔN isoform retains the DNA binding and C-terminal domains. (C) 2D western blot analysis of Jurkat whole cell protein extract. ZBP-89FL (solid triangle) and ZBP-89ΔN (open triangle) peptides share an electrohoretic mobility of ∼100 kDa and are separated on the basis of their distinct isoelectric points (pI). The acidic end of the focusing gel is at the left.
Figure 3
Targeting ZBP-89 exon 4 in mice. (A) Replacement of exon 4 with a Neomycin resistance cassette (NEO) by homologous recombination in ES cells. Top panel: bracket shows location of features encoded by exon 4, including initiation codon, acidic domain (black rectangle) and p300 interaction domain. Open boxes, basic domains; hatched boxes, zinc-fingers; stippled region, untranslated. Middle panel; Zfp148 genomic locus. Intron sizes (kb) are shown. Lower panel; targeting strategy. (B) Tail biopsy DNA genotyping. WT (Int 3 F/Int 4 R) and Δexon4 (Int 3 F/Neo R) amplimers. (C) Near-Mendelian inheritance of ΔExon-4 alleles. The differences between predicted and observed values were not significant.
Figure 4
ΔExon4 locus encodes mouse ZBP-89ΔN. (A) RT–PCR analysis was used to determine the variant mRNA structure in recombinant mice. Spleen whole-cell RNA was reverse-transcribed and amplified with primers within the indicated exons. RT–PCR of another ubiquitously expressed zinc-finger transcription factor, Sp1, was used for comparison and showed that the RNA samples were of uniform quality. Control experiments showed that PCR products were RT-dependent (data not shown), indicating that they were indeed derived from mRNA rather than possibly resulting from amplification of ZBP-89 related processed pseudogene sequences, such as ΨBERF1 (32). (B) DNA sequencing of recombinant cDNA showed that deletion of exon 4 resulted in direct splicing of exon 3 to exon 5, excluding the Neo cassette. The resulting reading frameshift predicts an alternative initiation codon corresponding to M128 of FL protein, similar to the naturally occurring human ZBP-89ΔN variant. (C) 2D immunoblot analysis of whole-cell spleen protein extracts from FL/FL, FL/ΔN and ΔN/ΔN mice, using conditions identical to those used for analysis of human samples (Figure 2C).
Figure 5
Growth delay and decreased survival in ΔNter mice. (A) Growth curve for male ZBP-89FL/FL (gray dashed line, n = 20), ZBP-89FL/ΔN (black dotted line, n = 36), and ZBP-89ΔN/ΔN (black solid line, n = 16) offspring. A similar pattern was seen with female mice (not shown). ** P < 0.01; * P < 0.05. A trend toward lower body weights in ZBP-89ΔN/ΔN persisted after 12 weeks, but was no longer statistically significant (data not shown). (B) Kaplan–Meier analysis of survival interval to the onset of morbidity or death. (C) Semi-quantitative RT–PCR analysis of ZBP-89FL and ZBP-89ΔN mRNA levels in the colon.
Figure 6
Increased susceptibility to DSS colitis in ZBP-89ΔN mice. (A–C) Representative H&E stains of normal (A), heterozygous (B) and ZBP-89ΔN/ΔN (C) mice. Inf = infiltrating lymphocytes; SE = submucosal edema; TM = transmural inflammation. (D) ΔNter expression accelerates onset of gastrointestinal bleeding, measured by fecal occult blood screening. (E) Mortality during DSS treatment: 50% of ZBP-89ΔN/ZBP-89ΔN mice die during 4% DSS treatment; all ZBP-89FL/ZBP-89FL and ZBP-89FL/ZBP-89ΔN mice survived (six in each group). (F) Colitis index scoring of DSS-treated mice, as described in Materials and Methods; six mice in each genotype cohort. | [
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16611361 | Generation of mice harbouring a conditional loss-of-function allele of Gata6
Abstract
The zinc finger transcription factor GATA6 is believed to have important roles in the development of several organs including the liver, gastrointestinal tract and heart. However, analyses of the contribution of GATA6 toward organogenesis have been hampered because Gata6-/- mice fail to develop beyond gastrulation due to defects in extraembryonic endoderm function. We have therefore generated a mouse line harbouring a conditional loss-of-function allele of Gata6 using Cre/loxP technology.
LoxP elements were introduced into introns flanking exon 2 of the Gata6 gene by homologous recombination in ES cells. Mice containing this altered allele were bred to homozygosity and were found to be viable and fertile. To assess the functional integrity of the loxP sites and to confirm that we had generated a Gata6 loss-of-function allele, we bred Gata6 'floxed' mice to EIIa-Cre mice in which Cre is ubiquitously expressed, and to Villin-Cre mice that express Cre in the epithelial cells of the intestine. We conclude that we have generated a line of mice in which GATA6 activity can be ablated in a cell type specific manner by expression of Cre recombinase. This line of mice can be used to establish the role of GATA6 in regulating embryonic development and various aspects of mammalian physiology.
Background
The mouse Gata6 gene encodes a 45 kD protein containing two highly conserved zinc-finger DNA binding domains with a Cys-X2-Cys-X17-Cys-X2-Cys motif that directs binding to the nucleotide sequence element (A/T)GATA(A/G) 1. GATA4, 5 and 6 make up a subset of GATA factors that have been implicated in the development of several organs including the heart, lung, gastrointestinal tract and liver [1-4]. Development of GATA6 null embryos arrests during gastrulation as a consequence of defects in extraembryonic endoderm function [2,5]. This early embryonic lethality can be rescued by complementing the GATA6 null embryos with a wild type extraembryonic visceral endoderm using tetraploid embryo complementation, and embryos derived by this process can survive until E10.5 [3]. Analyses of such Gata6-/-ES cell-derived embryos has revealed defects in hepatogenesis, which supports the proposal that GATA6 is an important developmental regulator. Although the ability to generate embryos from Gata6-/-ES cells by tetraploid embryo complementation has provided important insight into the contribution of GATA6 during early embryogenesis, this approach is not compatible with studying the role of GATA6 at later stages in development or its role in controlling differentiation of specific cell types. In the current report, we describe the generation of mice containing a conditional null allele of Gata6 that can be used for cell type-specific removal of GATA6 by Cre-mediated recombination.
Results and discussion
To ensure the elimination of GATA6 activity, we chose to flank Gata6 exon 2 (based on nomenclature described by Brewer et al [6,7]) with loxP elements because this exon encodes the majority of the GATA6 protein (Fig. 1A). We decided to use a recombineering approach to facilitate the accurate placement of the loxP elements following the procedure described by Lee et al [8]. The final targeting vector contained a loxP element located between exons 2 and 3. In addition, a cassette containing a loxP element lying immediately 5' to a neomycin phosphotransferase (neo) gene, which conferred resistance to G418, was introduced between exons 1 and 2 (Fig. 1A). FRT sites flanked the neo gene, in order to allow removal of neo by Flp recombinase. Two novel restriction endonuclease cleavage sites (EcoRI and PacI) were also introduced into the Gata6 targeting vector to allow the identification of correctly modified Gata6 alleles by Southern blot analyses. The targeting vector was introduced into R1 ES cells [9] by electroporation, and G418-resistant ES cell clones were tested for homologous recombination at the Gata6 locus by Southern blot analysis. Fig. 1B shows an example of a correctly targeted ES (Gata6loxP(FRTneoFRT)/+) cell line. Following digestion of ES cell genomic DNA with EcoRI, a probe that lies 3' to the short arm of homology used in the targeting vector identified an expected 8.1 kb wild type Gata6 DNA fragment, while correctly targeted cells contain an additional 3.2 kb fragment due to the introduction of a new EcoRI site. When NdeI digested DNA was probed with a DNA fragment lying 5' to the expected position of the introduced loxP site, it identified the predicted 13.2 kb wild type fragment and a novel 15.2 kb fragment, which resulted from introducing the neo cassette into the Gata6 locus. Five correctly targeted ES cell clones were recovered and three of these cell lines were used to generate chimeric mice by morula aggregation [10]. These chimeric animals were then mated with CD-1 mice and successful germline transmission of the Gata6loxP(FRTneoFRT) allele was confirmed by Southern blot analysis of tail DNA (Fig. 1B). Mice carrying a single Gata6loxP(FRTneoFRT)/+ allele were viable and fertile. However, we were unable to obtain mice that were homozygous for this allele, which suggested that the inclusion of the neo cassette disrupted GATA6 function and resulted in embryonic lethality as observed for Gata6-/- embryos [2,5]. We therefore established a mouse line, Gata6loxP/+, that lacked the neo cassette by inducing recombination between the FRT sites in vivo [11]. This was achieved by mating Gata6loxP(FRTneoFRT)/+ mice to a transgenic mouse, B6;SJL-Tg(ACTFLPe)9205Dym/J, in which Flp recombinase is widely expressed from the human beta actin gene promoter [11]. Correct excision of the neo cassette was confirmed in offspring by Southern blot and PCR analyses (Fig. 1B & C). Gata6loxP/+ mice were finally interbred to generate Gata6loxP/loxPmice, which are healthy, fertile and have reached maturity.
Figure 1
Generation of a Gata6 conditional null allele. (A) Schematic showing a map of the Gata6 genomic locus and the targeting vector with exons represented by open boxes. The relative position of Southern blot probes (lines), PCR primers (small arrowheads), loxP (large arrowheads) and FRT (ovals) sites, as well as cassettes encoding neomycin phosphotransferase (neo) and diphtheria toxin (DT), are included. Sizes of relevant EcoRI (e), NdeI (n), and PacI (p) restriction endonuclease fragments are shown in kilobase pairs (kb). (B) Southern blot analysis of genomic DNA isolated from ES cells (lanes 1, 2, 8 and 9) or mouse tails (lanes 3–7 and 10). An example of an ES cell line containing a correctly targeted Gata6loxP(FRTneoFRT) allele is shown in lanes 2 and 8. Mice harbouring the modified Gata6 allele were generated from these ES cells (lanes 3 and 10). Exon 2 of Gata6 was deleted (Gata6del) or the neo cassette alone was deleted, leaving Gata6 exon 2 flanked by loxP elements (Gata6loxP), by breeding Gata6loxP(FRTneoFRT) mice to transgenic mice expressing either Cre (lane 6 and 7) or Flp (lane 4) recombinases, respectively. The size of restriction fragments identified by 5' and 3' probes (Fig. 1A) was deduced from their position relative to standard DNA fragments. (C) The genotypes of mice and embryos were also determined by PCR amplification of genomic DNA. Primers were designed that differentiated between the Gata6+ (gt4F/4R; 159 bp), GataloxP (gt4F/4R; 250 bp) and Gata6del (k/o2F/2R; 568 bp) alleles, as well as neo (Neo A/B; 225 bp), flp (Flp F/R; 750 bp), and cre (Cre 1/2; 300 bp) transgenes.
We next addressed whether removal of GATA6 could be induced in vivo by expression of Cre recombinase. We chose to disrupt the Gata6 gene in the gastrointestinal tract because it is highly expressed in the epithelium and may have roles in controlling gut physiology or development. To produce Gata6loxP/+Villin-Cre mice, we bred Gata6loxP/loxP mice with transgenic mice (Tg(Vil-cre)997Gum) in which Cre expression was directed to the epithelial cells of the small intestine by the Villin (Vil1) promoter [12]. Using this transgenic strain of mice, Cre activity can be detected in the epithelium of the small intestine from E14.5 [12]. Gata6loxP/+Villin-Cre males were mated with Gata6loxP/loxP females to obtain Gata6loxP/+Villin-Cre control and Gata6loxP/loxPVillin-Cre experimental offspring. In previous experiments, we observed that the efficiency through which Cre mediates recombination between loxP sites varies between different transgenic male mice [13]. We noted a similar situation when using various Gata6loxP/+Villin-Cre males. In the majority of cases (4/5 stud males) we were unable to obtain Gata6loxP/loxPVillin-Cre offspring (n = 95 mice genotyped), and this lethality associated with presumptive loss of GATA6 in the intestine is currently under investigation. However, one Gata6loxP/+Villin-Cre male did produce offspring (12/90) whose genotype was determined to be Gata6loxP/loxPVillin-Cre by PCR analysis of tail DNA. We, therefore, used RT-PCR with Gata6 primers corresponding to nucleotide sequences predicted to be deleted after recombination between Gata6 loxP elements, to compare the steady-state levels of Gata6 mRNA in intestines isolated from control and experimental mice generated by this particular Gata6loxP/+Villin-Cre male. Fig. 2A shows that Gata6 mRNA could be detected in control intestine but not in intestine isolated from a subset of Gata6loxP/loxPVillin-Cre offspring. The remaining Gata6loxP/loxPVillin-Cre mice were found to continue to express Gata6 mRNA at varying levels (data not shown). We also examined expression of GATA6 protein in the intestines of Gata6loxP/loxPVillin-Cre offspring derived from the same Gata6loxP/+Villin-Cre male. Fig. 2B shows that GATA6 was detected as an abundant nuclear protein in the epithelial cells of control intestines. However, Gata6loxP/loxP Villin-Cre mice displayed a marked reduction in the abundance of intestinal GATA6 (Fig. 2B). From these results, we conclude that the Gata6 gene can be conditionally disrupted in Gata6loxP/loxP mice by cell-type specific expression of Cre recombinase.
Figure 2
Gata6 can be successfully ablated in the intestine of Gata6loxP/loxP mice by expression of Cre from the Villin promoter. (A) The steady-state level of Gata6 mRNA was compared in intestines isolated from four control Gata6loxP/+Villin-Cre (lanes 2–5) or experimental Gata6loxP/loxPVillin-Cre (lanes 6–9) mice. Amplification of Pol2 (Polr2a) mRNA showed that similar levels of starting material were utilized in each reaction. (B) Immunohistochemistry showing that GATA6 was detected as an abundant nuclear protein in epithelial cells (arrows) of control intestines, but was detected at greatly reduced levels in Gata6loxP/loxPVillin-Cre intestines.
Cre-mediated deletion of exon2 in Gata6loxP/loxP mice was predicted to result in loss of GATA6 function. If this were true, we expected that embryos homozygous for the deleted allele (Gata6del) should undergo developmental arrest, as was described for Gata6-/- embryos [2,5]. Gata6del/+ mice were produced by mating Gata6loxP(FRTneoFRT)/+ mice with an EIIa-Cre transgenic mouse (B6.FVB-Tg(EIIa-cre)C5379Lmgd/J). The EIIa-Cre mouse expresses Cre recombinase in nearly all tissues, including those of pre-implantation embryos, and has been used previously to mediate recombination between loxP sites in germ cells [14]. Gata6+/del progeny were identified by Southern blot analyses of NdeI digested tail DNA. As shown in Fig. 1B, Southern blots hybridized with the 5' Gata6 probe (Fig. 1A) revealed the conversion of a 15.2 kb NdeI fragment from the Gata6loxP(FRTneoFRT) allele to an expected 12 kb NdeI fragment from the Gata6del allele. Gata6+/del were then mated inter se, and the genotype of embryos collected between E8.5 and E11.5 was determined by PCR analysis of genomic DNA. Of 77 embryos recovered, 27 were Gata6+/+ and 50 were Gata6+/del (Table 1). In addition, 23 partially resorbed empty decidual masses were recovered, which we believe likely resulted from the early developmental arrest of Gata6del/del embryos. We finally determined whether the developmental lethality associated with loss of GATA6 could be induced by expression of Cre in pre-implantation embryos. To achieve this, Gata6loxP/loxP mice were bred to Gata6+/delEIIa-Cre mice, which are heterozygous for the EIIa-Cre transgene. The genotype of resulting embryos ranging from E8.5 to E11.5 was again determined by PCR. While we recovered 14 Gata6loxP/+, 18 Gata6loxP/del, and 16 Gata6loxP/+EIIaCre embryos, no Gata6loxP/delEIIaCre embryos were identified although 13 empty resorbing decidual masses were observed (Table 1).
Table 1
Embryonic lethality associated with loss of GATA6 function
Conclusion
In summary, we conclude that we have generated a line of mice in which GATA6 transcriptional activity can be ablated by expression of Cre. We believe that the availability of this mouse will be useful to elucidate the contribution of GATA6 during organogenesis as well as its physiological role in adult mice.
Methods
Plasmid construction
A bacterial artificial chromosome (BAC # RP23-410I12) that contained Gata6 genomic DNA was obtained from BACPAC Resources, Children Hospital Research Institute, Oakland, CA. Plasmid pNeb-DT-GATA6 was generated by cloning a 12.2 kb EcoRV/PacI genomic fragment containing Gata6 exons 1 and 2 into the PacI/HincII site of a vector, pNEB193-DT, which contained a diphtheria toxin (DT) expression cassette to enrich against random integration of the targeting plasmid in ES cells. A cassette containing loxP-neo-loxP was amplified by PCR from the plasmid pSV-LNL (modified from Zhang et al)[15] using the following primers: Gata6ET1:GCTTGCTGTTTGAGTCTACCCCATTTCTGCCTGTTTCTTGACATCCCTTCGAATTCTGGTACCGGCGCGCCTAGTCGAC, Gata6ET2:ATCCATTATTGTCAATGTCTAAAGATGGAATTGTCTCTGCACAAGCTATCTTCTCAACTCGAGCCCTTAATTAACCGGT. These oligonucleotides contained 55 bp of sequence from Gata6 intron 2 (underlined). This amplicon was introduced into Gata6 intron 2 sequence in pNEB-DT-GATA6 by homologous recombination in E.coli following the procedure described by Lee et al [8] The loxPneoloxP cassette was then converted to a single loxP site by expression of Cre recombinase [8]. A loxP(FRTneoFRT) cassette was then amplified from pSV-LFNF using primers Gata6ET3:CACGCTGGTGGTTGTAAGGCGGTTTGTGTTTAAGGTGTGCGGTTGGCCTGGACGTGTGGTACCGGCGCGCCTAGTCGAC, Gata6ET4:GAAAAAGTTACCTAGCCCAGAGAAAGTGAGATGCCAGGAAAGGCATAAGGATATCAACTCGAGCCCTTAATTAACCGGT. These oligonucleotides contain 56 bp (Gata6ET3) and 52 bp (Gata6ET4) of sequence from Gata6 intron 1, respectively. This cassette was introduced into Gata6 intron 1, again using homologous recombination in E.coli. to generate the final targeting vector (Fig. 1A).
ES cell targeting and animals
Linear targeting vector (100μg) was introduced into R1 ES cells by electroporation, and the genotype of colonies resistant to 350μg/ml of Geneticin (Gibco BRL) was determined by Southern blot (Fig. 1B). Chimeric mice were generated by aggregation of ES cells with CD-1 morulae as described previously [10] and the modified allele was passed through the germline by breeding chimeras to CD1 mice. Gata6loxP/loxP mice were produced by breeding Gata6loxp(FRTneoFRT)/+ mice to B6;SJL-Tg(ACTFLPe)9205Dym/J mice [16] (Jackson Labs) to delete the FRTneoFRT cassette by Flp-mediated recombination in the germline. The ACTFLPe transgene was removed by breeding F1 Gata6loxP/loxP mice into CD-1 mice. Gata6+/del mice were generated by mating Gata6loxp(FRTneoFRT)/+ animals with B6.FVB-Tg(EIIa-cre)C5379Lmgd/J transgenic mice [17] (Jackson Labs) to allow Cre-mediate recombination between loxP elements in the germline. The EIIa-Cre transgene was removed by breeding Gata6+/del F1 mice with CD1 mice. The MCW IACUC committee approved all procedures using animals.
Southern blot, PCR and RT-PCR
Southern blot analyses were performed using standard conditions with probes indicated in Fig. 1. Genotypes were determined by PCR using the following oligonucleotide primer pairs: Gata6 gt4F/4R, GTGGTTGTAAGGCGGTTTGT, ACGCGAGCTCCAGAAAAAGT; Gata6 k/o2F/2R, AGTCTCCCTGTCATTCTTCCTGCTC, TGATCAAACCTGGGTCTACACTCCTA; Flp F/R, GGTCCAACTGCAGCCCAAGCTTCC, GTGGATCGATCCTACCCCTTGCG [16]; Cre 1/2, GTTCGCAAGAACCTGATGGACA, CTAGAGCCTGTTTTGCACGTTC [18]; Neo A/B, GCCAACGCTATGTCCTGATAGCGGT, AGCCGGTCTTGTCGATCAGGATGAT. RT-PCR was performed as described previously [19] with the following primer pairs: Gata6 9/10; AGTTTTCCGGCAGAGCAGTA, AGTCAAGGCCATCCACTGTC, Pol2 F/R; CTGATGCGGGTGCTGAGTGAGAAGG, GCGGTTGACCCCATGACGAGTG.
Immunohistochemistry
Immunohistochemistry was performed using antigen retrieval in citrate buffer as described previously [13] using an anti-GATA6 antibody (AF1700 R&D Systems,1/1000 dilution).
Abbreviations
Neo, neomycin phosphotransferase
Authors' contributions
C.P.S. generated the targeting vector and carried out analyses of conditional knockout mice, as well as contributed to experimental design and draft of the manuscript. J. L. generated aggregation chimeras. S.A.D. conceived of the study, contributed to experimental design and interpretation of results, and coordinated the project and writing of the manuscript.
Acknowledgements
We would like to thank Drs. Robert Burgess and Francis Stewart for providing plasmids and Dr. Neal Copeland for providing bacterial strains used in recombineering. Pregnant mare serum gonadotrophin (PMSG) used in superovulation was obtained from Dr. A.F. Parlow at the National Hormone and Peptide Program (Torrance, CA). We are also grateful to Dr. Michele Battle for critically evaluating the manuscript and for providing plasmids. This work was supported by an NIH training grant from the MCW cardiovascular research centre to C.P.S and NIH grants to S.A.D. | [
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16787536 | Targeted disruption of cubilin reveals essential developmental roles in the structure and function of endoderm and in somite formation
Abstract
Background
Cubilin is a peripheral membrane protein that interacts with the integral membrane proteins megalin and amnionless to mediate ligand endocytosis by absorptive epithelia such as the extraembryonic visceral endoderm (VE).
Results
Here we report the effects of the genetic deletion of cubilin on mouse embryonic development. Cubilin gene deletion is homozygous embryonic lethal with death occurring between 7.5–13.5 days post coitum (dpc). Cubilin-deficient embryos display developmental retardation and do not advance morphologically beyond the gross appearance of wild-type 8–8.5 dpc embryos. While mesodermal structures such as the allantois and the heart are formed in cubilin mutants, other mesoderm-derived tissues are anomalous or absent. Yolk sac blood islands are formed in cubilin mutants but are unusually large, and the yolk sac blood vessels fail to undergo remodeling. Furthermore, somite formation does not occur in cubilin mutants. Morphological abnormalities of endoderm occur in cubilin mutants and include a stratified epithelium in place of the normally simple columnar VE epithelium and a stratified cuboidal epithelium in place of the normally simple squamous epithelium of the definitive endoderm. Cubilin-deficient VE is also functionally defective, unable to mediate uptake of maternally derived high-density lipoprotein (HDL).
Conclusion
In summary, cubilin is required for embryonic development and is essential for the formation of somites, definitive endoderm and VE and for the absorptive function of VE including the process of maternal-embryo transport of HDL.
Background
Cubilin is a 460-kDa peripheral membrane protein expressed by a number of absorptive epithelial cells including those of the renal proximal convoluted tubule, ileum and yolk sac extraembryonic visceral endoderm (VE) [1]. The first described function of cubilin was as the receptor for intrinsic factor-vitamin B12/cobalamin (Cbl), serving a critical role in the intestinal absorption of Cbl [2,3]. Cubilin was later shown to be an endocytic receptor for apolipoprotein A-I (apoA-I)/high density lipoprotein (HDL), mediating uptake of HDL in the kidney and VE [4,5]. Other cubilin ligands include albumin, transferrin, immunoglobulin light chains, vitamin D-binding protein, myoglobin, galectin-3 and Clara cell secretory protein [6].
Three cell surface integral membrane proteins have been shown to interact with cubilin. The first identified was megalin, an endocytic receptor belonging to the low density lipoprotein receptor (LDLR) family [7,8]. Megalin functions together with cubilin to mediate endocytosis of apoA-I/HDL, presumably facilitating endocytosis of the cubilin-apoA-I/HDL complex. The cation-independent mannose 6-phosphate/insulin-like growth factor II-receptor (CIMPR) is another endocytic receptor that binds to cubilin [9], although the functional significance of its interaction with cubilin remains to be established. The ~48-kDa type I transmembrane protein, amnionless (AMN), is the most recent integral membrane protein found to interact with cubilin [10]. AMN is essential for efficient transport of cubilin to the apical cell surface as well as for membrane anchoring of cubilin [11,12],
Given the fact that cubilin is expressed by trophectoderm and VE [13,14], it is believed to play an important role in maternal-embryonic transport of nutrients. Several additional pieces of evidence support this hypothesis. First, cubilin has been shown to mediate VE uptake of holoparticle HDL, HDL-associated cholesterol and apolipoprotein A-I [8,14]. Furthermore, the work of Sahali et al. [15] demonstrated that cubilin monoclonal antibodies infused into circulation of pregnant rats (9 dpc), bound to VE cubilin and induced a spectrum of developmental abnormalities and embryonic resorption within 24–48 hours of infusion. The abnormalities included retarded embryonic growth, craniofacial defects involving the eye, ear and neural tube, hydrocephaly and telencephalic hypoplasia. Similarly, growth retardation and morphological anomalies were obtained when rat embryos (10 dpc) were cultured ex utero in the presence of cubilin antibodies [16]. Here we characterize the consequences of targeted deletion of the mouse cubilin gene on embryonic development.
Results
Generation of cubilin-/- mice
To generate mice with targeted disruption of the cubilin gene (and concomitant knock-in of the EGFP reporter gene) we cloned and fully mapped the exon-intron structure of the 5' portion of the murine cubilin gene. A mouse cubilin gene-targeting vector was designed to create a null mutation through deletion of exons 1–6 (Fig. 1). After electroporation of the construct and G418 positive selection and FIAU negative selection, ten ES clones (out of 132 screened) were identified that had the desired recombination based on Southern analysis (using both upstream and downstream flanking probes) (Fig. 1). Targeted ES clones were injected into C57BL/6J blastocysts and the blastocysts were transferred to foster mothers to obtain chimeric mice. Two male chimeras were obtained from the first targeted ES cell line tested and found to be germ line competent through the generation of heterozygous offspring. As shown in Figure 1C, Southern analysis confirmed that offspring from one of these mice contained the properly targeted cubilin allele.
Figure 1
Targeted deletion of the mouse cubilin gene. A, is a diagram of the structural and functional domains of cubilin. Ligand binding regions indicated in the diagram are based on published studies [12, 33, 34]. B, is a diagram of the gene targeting strategy showing the organization of cubilin gene exons (vertical lines in uppermost model and rectangles in expanded region shown below), the wild-type and targeted alleles and the location of 5' and 3' DNA probes used for Southern blot analysis. White portions of boxed exons depict untranslated sequences. The promoter-less EGFP (containing Kozak and ATG sequences) and loxP-floxed neoR cassettes were inserted into exon 1, 33 bp upstream of the cubilin ATG and 16 bp downstream from the proximal-most transcription initiation site, resulting in a replacement of the majority of exon 1. Homologous recombination between the targeting construct and the cubilin locus results in deletion of exon 1 coding sequence and all of exons 2–6. C, Southern analysis of HindIII/HhaI and BglII digested DNA from representative wild-type (WT) and targeted (-/+) ES cell clones using 5' and 3' probes (left and right panels, respectively). The wild-type allele yields a 7 kb band and the knockout allele yields a 5.5 kb band when genomic DNA is digested with HindIII and HhaI and hybridized with the 5' probe. Additionally, the wild-type allele yields an 11.3 kb band and the knockout allele yields a 7 kb band when genomic DNA is digested with BglII and hybridized with the 3' probe. The results show that the correct recombinant allele is present in the selected ES cell clone. D, Southern analysis of BglII digested tail DNA from a wild-type (WT) and a correctly targeted heterozygous (-/+) mouse.
The cubilin exon 1–6 deletion leads to embryonic lethality
Genotypic analysis was performed on 220 offspring from heterozygous intercross matings. As a result, 100 wild-type, 120 heterozygous and no homozygous offspring were detected (Table 1). These non-Mendelian ratios indicate that lethality occurs in embryos homozygous for the targeted cubilin gene deletion. Furthermore, since the frequency of heterozygotes at 4 weeks was much lower than expected (1:1 versus the expected 2:1 heterozygous:wild-type ratio), embryonic lethality appeared to be occurring in some heterozygotes. To substantiate this possibility we performed genotypic analysis of 181 4-wk offspring from wild-type × heterozygote matings. As a result, 109 wild-type and 72 heterozygous offspring were detected. Based on a chi-square test of these data and a resulting p value = 0.0074, the data are consistent with the hypothesis that haploinsufficiency of cubilin causes embryonic lethality, although with incomplete penetrance.
Table I
Genotype frequency of progeny from heterozygote (cubilin exon1–6+/-) matings
Asterisk indicates that the chi-square test was based on a 1:2 ratio of wild-type to heterozygote offspring.
Retrograde genotypic analysis was performed on embryos from heterozygous intercross matings (Table 1). At 7.5 dpc homozygous embryos were obtained at a frequency consistent with a normal Mendelian ratio. From 8.5–10.5 dpc, homozygous embryos were detected, but not at a frequency in accordance with Mendelian expectations (Table 1). After 13.5 dpc, no homozygous embryos were detected. Together, these findings indicate that homozygous embryos die over a range of developmental stages between 7.5 to 13.5 dpc. Consistent with this conclusion was the relatively high number of embryo resorptions observed after 7.5 dpc (Table 1).
To establish that embryos homozygous for the targeted cubilin gene deletion of exons 1–6 were indeed cubilin null, RT-PCR analysis was performed using two primer pairs designed to amplify separate 5' and 3' regions of the cubilin transcript. No cubilin mRNA was detectable by either primer pair using RNA isolated from homozygous 8.5 dpc embryos (Fig. 2). Furthermore, immunohistological analysis confirmed that there was no cubilin detectable in paraffin embedded sections of homozygous 8.5 dpc embryos (data not shown).
Figure 2
Embryos homozygous for targeted deletion of cubilin do not express cubilin transcripts. Shown are RT-PCR analyses of the expression of cubilin transcripts in RNA from heterozygous (+/-) and null (-/-) 8.5 dpc embryos. In A, a primer pair corresponding to a 5' region of the cubilin transcript was used to assess cubilin transcript expression. In B, a primer pair corresponding to a 3' region of the cubilin transcript was used to assess cubilin transcript expression.
Developmental retardation and mesodermal defects of homozygous embryos
Morphological analysis of 8–8.5 dpc embryos revealed that homozygous embryos were considerably smaller than wild-type or heterozygous littermates (Fig. 3). Mutant 8–8.5 dpc embryos were morphologically similar to wild-type 7.5 dpc embryos. However, rather than exhibiting a normal cylindrical shape, mutant 8–8.5 embryos appeared acorn shaped owing to a relatively smaller embryonic component as compared to extraembryonic component.
Figure 3
Cubilin deficient 8–8.5 dpc embryos display growth retardation with formation of some mesodermal structures, but not somites. Shown are wild-type (A), heterozygous (B) and homozygous 8–8.5 dpc embryos (C-E), derived from a heterozygous intercross mating. Embryos in A and B have been dissected to lie in a planar configuration where as embryos in C-D are intact. Shown in F and G are H&E stained sections of 8.5 dpc wild-type (F) and mutant (G) embryos. Mutant 8.5 dpc embryos possess an allantois and amnion, but lack somites. A, allantois; Am, amnion; S, somite. Bars in A-G = 100 μm.
Histological examination of null 8–8.5 dpc embryos revealed that all mutant embryos underwent gastrulation, but did not form somites (Fig. 3F and 3G). In contrast, 8–8.5 dpc wild-type littermates had formed 3–9 somites. Other mesodermal structures did form in the 8–8.5 dpc mutants, albeit with some variability. For example, an allantois was present in 5 of 7 mutant 8–8.5 dpc embryos examined. Most mutant embryos (5 of 7) also had an amnion and chorion, tissues having mesoderm components. While blood islands formed in the yolk sacs of all 8–8.5 dpc mutants they appeared unusually large, extending into the exocoelomic cavity to a greater extent than wild-type blood islands (Fig. 4). Such blood islands appeared to have a larger than normal number of mesodermal cells. Additionally, blood island hematopoietic development appeared to be retarded as evidenced by the failure of the cells to exhibit the characteristic rounded hematopoietic cell morphology (Fig. 4).
Figure 4
Enlargement of blood islands in cubilin mutants. Shown are H&E stained sections of an 8–8.5 dpc wild-type (A) and mutant (B) embryo. C and D show high magnification views of blood islands from embryos shown in A and B. Embryos in panels A and B are oriented such that anterior is to the right and posterior is to the left. BI, blood island. Bars in A and B = 100 μm. Bars in C and D = 25 μm.
By 9.5 dpc, surviving homozygous embryos had not undergone axial rotation but displayed an overall morphological appearance of wild-type 8–8.5-dpc embryos (Fig. 5). Mutant 9.5 dpc embryos had formed a primitive heart and neural head folds (4 of 4 nulls examined) (Fig. 5) and paired dorsal aortae (data not shown), but lacked somites. Homozygous embryos surviving beyond 9.5 dpc (i.e., 11.5 dpc) were also grossly similar to wild-type 8.0 dpc embryos, but still lacked somites (data not shown). Together these findings indicate that lack of cubilin results in developmental retardation that initiates prior to 8.0 dpc, and a failure to achieve developmental milestones appropriate for a wild-type 8.0 dpc embryo, most notably, failure to form somites. The complete lack of somites at 9.5 dpc suggests that the basis is due to some defect of the paraxial mesoderm rather than merely being a consequence of retarded development.
Figure 5
Cubilin-deficient 9.5 dpc embryos are developmentally retarded and lack somites. A shows a wild-type 9.5 dpc embryo and B a mutant littermate. NF, neural folds; A, allantois; H, heart; YS, yolk sac. Bars in A and B = 200 μm.
It is important to note that embryos confirmed by genotyping to be heterozygous were indistinguishable morphologically from wild-type littermates over a range of stages examined extending from 7–11.5 dpc (Fig. 3B). This indicates that the lethality of heterozygous embryos, apparent from genotypic analysis, is occurring beyond 11.5 dpc.
Failure of yolk sac blood vessels to undergo remodeling
To assess the effects on blood vessel formation, embryos from heterozygous matings were immunolabeled with anti-PECAM-1. As shown in Figure 6A, yolk sac blood vessel formation in 9.5 dpc mutants appeared retarded as compared to wild-type littermates (Fig. 6B). Normally by 9.5 dpc the yolk sac vessels of the wild-type embryo have undergone remodeling to form larger diameter vessels (Fig. 6B, inset). By contrast, the pattern of yolk sac vascular development in 9.5 dpc mutants appeared as an interconnected network of small diameter blood vessels (Fig. 6A), a pattern similar to that seen in an 8.5 dpc wild-type embryo yolk sac. In some areas of the extraembryonic yolk sac, PECAM-1 labeling often showed enlarged sinusoidal-like vessels (data not shown), which was consistent with the enlarged blood island-like structures observed in H&E stained sections (Fig. 4).
Figure 6
Yolk sac blood vessels of 9.5 dpc cubilin mutants fail to undergo remodeling. Anti-PECAM-1-labeled vasculature in a 9.5 dpc homozygous embryo (A), a 9.5 dpc wild-type (WT) embryo (littermate to that shown in A) (B), 8.5 dpc wild-type embryo (C) and 11.5 dpc homozygous embryo (D). The inset panel in B shows a higher magnification view of the remodeled yolk sac vasculature in the boxed region of the 9.5 dpc wild-type yolk sac. YS, yolk sac; A, allantois; NF, neural folds; H, heart. Bars in A and B = 500 μm. Bars in C and D = 200 μm.
When the PECAM-1-labeled yolk sac vasculature of 11.5 dpc mutants was compared to that of wild-type embryos it was again evident that the blood vessels of homozygous embryos had not progressed beyond that of 8.5 dpc wild-type embryos. As shown in Figure 6D, yolk sac blood vessels of an 11.5 dpc mutant had not undergone the remodeling that normally occurs after 8.5 dpc.
In addition to the yolk sac vascular remodeling anomalies, the allantoic vasculature of 11.5 dpc mutants was aberrant. Instead of a branched central blood vessel normally present in an 8.5 dpc embryo [17], the allantoic blood vessels of 11.5 dpc mutants were bulbous and lacked branching (Fig. 6D). Since this abnormality was not observed in 9.5 dpc mutants, it can be concluded that dysmorphogenesis of the allantoic vasculature occurs after 9.5 dpc and perhaps contributes to a chorioallantoic placental defect.
Abnormalities of embryonic and visceral endoderm in cubilin mutants
Considering that cubilin is normally present on the apical surface of the columnar VE (from 6.0 dpc to at least 9.5 dpc) and on a population of the squamous cells within the embryonic endoderm (EE) (from 7.3–8.0 dpc) [13], we next evaluated the impact of cubilin deficiency on these embryonic endodermal tissues. Examination of homozygous 8–8.5 dpc embryos revealed a number of endodermal anomalies (Fig. 7). In place of a normal squamous definitive endoderm (Fig. 7C), the mutant embryos had both cuboidal and columnar cells arranged in a stratified cuboidal epithelium (Fig. 7D, arrow) as well as simple columnar epithelium (not shown). Additionally, mutant cells in these epithelia had apical processes that may be either microvilli or cilia. Such luminal surface structures are normally not found on EE cells. Based on the findings, the absence of cubilin inhibits the formation of an epithelium morphologically comparable to that of a normal definitive endoderm at this stage, however, it remains to be determined whether this is a result of a defect in the specification and differentiation of the definitive endoderm.
Figure 7
Cubilin mutants display anomalies in the epithelial morphology of embryonic endoderm. Shown are H&E stained sections of wild-type (A, C and E) and mutant (B, D and F) 8–8.5 dpc embryos. Embryos in panels A and B are oriented such that anterior is to the right and posterior is to the left. BI, blood island; S, somite; EE, definitive embryonic endoderm; VE, yolk sac visceral endoderm, Asterisk points to aberrant epithelium in place of the normal EE. Bars in A-F = 100 μm.
Examination of the VE of homozygous 8–8.5 dpc embryos revealed it to also be morphologically abnormal. In contrast to the simple columnar VE of normal embryos, the VE of cubilin mutants was a stratified epithelium, comprised of cuboidal and columnar cells. Furthermore, mutant VE cells also lacked the large apical vacuoles that are characteristic of VE cells (Fig. 7E and 7F). Given the morphological similarities between the cells of mutant embryonic endoderm there was not a discernable demarcation between the normally distinct EE and VE epithelia. However, a clear demarcation between the VE and an epithelium that would normally be EE was evident upon immunohistological analysis of GFP reporter expression (from the EGFP cassette inserted into the cubilin gene) and megalin expression in mutant 8.0 dpc embryos (Fig. 8). This indicated that although being morphologically similar, the VE and presumed EE of mutants were distinct at the molecular level. Furthermore, the fact that megalin was appropriately expressed on the apical surfaces of the mutant VE indicated that cubilin is not required to direct cell surface trafficking of megalin.
Figure 8
Green fluorescent protein and megalin expression in yolk sac visceral endoderm of mutant embryos. The cubilin gene targeting strategy (see Fig. 1) created a GFP reporter for cubilin locus expression. Shown in A is an H&E stained section of a homozygous 8.0 dpc embryo. Shown in B is anti-GFP fluorescence of a serial section to the one shown in A. Note that cells within the blood islands are expressing the GFP reporter. Shown in C is an anti-megalin stained 8.0 dpc mutant embryo. Embryos in panels A-B are oriented such that anterior is to the left and posterior is to the right. The embryo in C is oriented with anterior to the right and posterior to the left. VE, yolk sac visceral endoderm; BI, blood island; Am, amnion; Asterisk, points to the boundary of the VE and presumed EE.
The visceral endoderm of homozygous embryos does not mediate uptake of maternal-derived HDL
A normal function of VE is to mediate uptake of maternal-derived HDL [8]. We therefore evaluated the capacity of cubilin-deficient VE to mediate uptake of maternal-derived HDL. As shown in Figure 9, the VE of heterozygous 8.0 dpc embryos from DiI-HDL-infused mothers showed strong DiI labeling within VE cells. DiI label in these embryos was present exclusively in VE which is the only yolk sac endodermal component expressing cubilin as evidenced by expression of the cubilin gene reporter, EGFP. By contrast, there was no detectable DiI labeling of the VE of homozygous embryos (Fig. 9D–F). These findings indicate that VE uptake of HDL is a cubilin dependent process.
Figure 9
Maternally derived HDL is not taken up by the yolk sac visceral endoderm of cubilin-deficient embryos. Shown are confocal micrographs of a heterozygous 8.0 dpc embryo (A-C) and a homozygous littermate (D-F) isolated 1 hour after DiI-HDL was infused into the mother. The distal portion of the VE was removed from all embryos shown. VE, visceral endoderm; EE, embryonic endoderm.
Discussion
Considering that cubilin forms a complex with AMN (forming the cubam complex), that AMN and cubilin are co-expressed in absorptive epithelial including the VE [11], and that AMN is required to mediate key aspects of cubilin function (e.g., membrane association, apical sorting) [11,12], it is not surprising that the phenotype of AMN-deficient mutants [18] closely resembles that of the cubilin nulls described. Like the cubilin null embryos, AMN-deficient embryos display arrested development, with most mutants not advancing morphologically beyond a normal 8–8.5 dpc embryo. On a mixed 129Sv/C57BL/6 genetic background, both mutants form an amnion. General mesoderm formation in the cubilin and AMN mutant embryos appears normal in that both mutants form an allantois, blood islands, yolk sac blood vessels and a heart. However, both mutants fail to form somites, indicating that somitogenesis is in some manner defective (e.g., insufficient paraxial mesoderm or defective condensation and/or segmentation). Both mutants also display endoderm defects as evidenced by the inability of cubilin-deficient VE to facilitate uptake of maternal HDL and the inability of AMN-deficient VE and EE to support wild-type epiblast development in chimeric embryos [18]. The morphological abnormalities of VE and definitive endoderm observed in the cubilin mutants also highlight the importance of cubilin in formation and/or maintenance of these epithelia. Such morphological abnormalities were not reported in the phenotype of AMN-deficient mutants. It remains to be established whether cubilin and AMN are playing roles in the specification and differentiation of the definitive endoderm, which forms during gastrulation by the recruitment of epiblast cells through the primitive streak [19].
The close similarities between the phenotypes of mouse amn and cubilin mutants also suggest that both genes act co-operatively in the VE to support embryonic growth and to control the formation of paraxial mesoderm. How AMN and cubilin mechanistically mediate these processes remains to be resolved. The endocytic function of the cubam complex and its apical expression in VE supports the possibility that it functions in the transport of vital maternal nutrients/factors. The spectrum of maternally derived nutrients/factors transported by VE cubam is potentially quite broad considering the multi-ligand binding capacity of cubilin which includes albumin, transferrin, immunoglobulin light chains, vitamin D-binding protein, myoglobin, galectin-3, Clara cell secretory protein, apoA-I and HDL [6]. Furthermore, at least one cubilin ligand, HDL, is a complex of multiple factors including apolipoproteins, phospholipids, cholesterol and lipid soluble vitamins such as retinol, which is converted to retinoic acid through the action of retinaldehyde dehydrogenase-2 (Raldh2). Indeed, genetic deficiency of mouse Raldh2 leads to early embryo lethality (~10.5 dpc), absence or reduction in somites, yolk sac vascular defects, enlarged heart and a failure of embryos to undergo axial rotation [20,21], all similar to cubilin and amn mutants.
By contrast to the developmental anomalies and early lethality of the cubilin and amnionless nulls, megalin-deficient mice die perinatally and display abnormal morphogenesis of the forebrain (e.g., holoprosencephaly), lung and kidney [22]. Thus, despite the evidence that megalin functions together with cubilin and that it is co-expressed with cubilin in the VE [13], any joint function that these two proteins may have in the early embryo is evidently not vital. Interestingly, mice deficient in disabled-2 (dab2), an adaptor of megalin, display several of the abnormalities observed in cubilin nulls including disorganization of the VE and loss of apical vesicles [23]. However, dab2-/- mutants arrest earlier in development than do cubilin nulls and do not undergo gastrulation. Dab2 has also been shown to be required for cubilin endocytosis in VE [24]. The earlier lethality of dab2 null embryos may be an indication that numerous endocytic receptors expressed in the VE including megalin, cubilin and perhaps amnionless are dependent on dab2.
The importance of maternal derived lipoproteins as a source of cholesterol for normal development is well established [25]. Through the use of cubilin antagonists, the ability of cubilin to mediate VE uptake of holoparticle HDL, HDL-associated cholesterol and apoA-I has been demonstrated in mouse embryos cultured in vitro [8,14]. Our findings provide in vivo evidence that early (8.0 dpc) embryonic uptake of HDL from the maternal circulation is a cubilin-dependent process of the VE. Given the fact that the VE expresses apoA-I and the major LDL constituent, apoB [26], it is likely that cholesterol and other maternal HDL-derived constituents taken up by VE cells via cubilin are incorporated into new HDL and LDL particles and subsequently delivered to the embryo. In support of this, there is evidence that nascent lipoprotein particles have been localized to the rough endoplasmic reticulum and secretory vesicles of the 9.5 dpc mouse yolk sac and seen in the pericellular space underlying the endoderm [27].
Despite the fact that cubilin and AMN deficiency in mice leads to embryonic lethality, mutations in human and canine AMN that cause impaired apical targeting [28] and function (i.e., malabsorption of vitamin B12) of the cubam complex, nevertheless have no effect on embryonic development. One explanation for this apparent paradox may relate to the fact that there are species differences in yolk sac function in rodents, dogs and humans. Another explanation comes from the studies of Tanner et al. [29] who have shown that mutations affecting exons 1–4 of human AMN (OMIM 261100, megaloblastic anemia, MGA1) do not prevent the production variant transcripts and polypeptides resulting from alternative transcriptional start sites and alternative translation initiation sites. Indeed, the two AMN mutants that cause premature termination of translation (14ΔG and 208-2A→G) also express an alternative AMN polypeptide of 40 kDa and several minor species of 44, 42 and 38 kDa [29]. All of the alternative AMN polypeptides would contain the chordin-like module, transmembrane domain and cytoplasmic domain, but would lack various lengths of the amino terminal portion of the full-length protein. While these alternative polypeptides apparently are missing regions required for vitamin B12 uptake, they evidently confer enough normal function to sustain embryonic development. How these alternative polypeptides function, particularly with respect to cubilin, remains to be established. One possibility is that they retain the ability to mediate trafficking of cubilin to the apical surfaces of epithelial cells. However, mutations of the canine AMN gene that disrupt cubilin intracellular trafficking have no apparent effect on embryonic development [28]. It is not known whether the canine AMN mutants express alternative AMN polypeptides similar to those observed in humans. If so, it is possible that the polypeptides possess some activity that overcomes the need for apical expression of the cubam complex to mediate normal development.
Conclusion
The present study reveals an indispensable role for cubilin in mouse embryogenesis particularly in the formation of endoderm and somites. The findings also highlight the importance of cubilin in the process of maternal-embryo transport of HDL by the VE.
Methods
Targeting vector design and generation of mutant mice
Degenerate deoxyoligonucleotide primers 5'-CTICACCARCCICGIATG-3' and 5'-CCRTTRRATYTCRCAYTC-3' were synthesized based on cDNA sequences conserved in the 5' region of both rat cubilin (GI: 24475743) and human cubilin (GI: 3929528). Preparation of template cDNA from adult mouse kidney total RNA and PCR amplification were done using methods described previously [8]. Cycling parameters for PCR amplification were: 94°C for 5 min and then 30 cycles of 94°C for 1 min, 46°C for 1 min and 72°C for 1.5 min. The expected ~820 bp product was sequenced and found to have 93% identity with a 5' portion of the rat cDNA sequence.
The mouse cubilin cDNA was used as a probe to screen a 129-strain mouse genomic library (Genome Systems, Inc.) and a BAC clone (24267) was isolated. DNA sequencing of ~20 kb of the BAC clone was performed to characterize the intron-exon organization of the 5' portion of the cubilin gene, including the region containing exons 1–10. A search of GenBank showed that the sequence of BAC clone 24267 was contained within the mouse chromosome 2 genomic contig GI:82796355.
The 3' arm of cubilin replacement-type targeting vector was constructed from a 5.4-kb EcoRI fragment containing exons 7 and 8 of the mouse cubilin gene. The 5' arm was constructed from a 4.3 kb ApaI-NaeI fragment containing cubilin exon 1 disrupted by insertion of a enhanced green fluorescent protein (EGFP)-N1 (Clontech, Mountain View, CA), neoR (loxP floxed) cassette 33 nucleotides 5' to the first ATG (see Additional file 1). A tk negative selection cassette was placed at the 5' end of this construct. The resulting targeting vector was linearized by NotI digestion and electroporated into murine 129/Sv-strain embryonic stem (ES) cells. After electroporation of the construct and G418 positive selection and 1-(2-deoxy-2-fluoro-8-d-arabinofuranosyl)-5-iodouracil (FIAU) negative selection, clones having the desired homologous recombination were identified by Southern analysis (Fig. 1C). One of the targeted ES clones was injected into C57BL/6J blastocysts that were then transferred to foster mothers to obtain chimeric mice. Two male chimeras were obtained that were germ line competent. Southern analysis confirmed that offspring from these mice contained the properly targeted cubilin allele. Mice used in this study were maintained on a mixed 129Sv/C57BL/6 genetic background.
Genotypes of progeny from heterozygote intercrosses were determined by PCR. For embryos between 7.5 and 8.5 days postcoitum (dpc) DNA was isolated from the whole embryo and used in PCR. For embryos between 9.5 and 14.5 dpc, DNA for genotyping was isolated from the yolk sac. Two primer pairs were used for PCR-based genotyping. To detect the wild-type cubilin allele, PCR was performed with tail clip genomic DNA preparations using a cubilin sense strand primer, 5'-GCCAAGTAGACCAGGCTGAC-3' (residues 10422223–10422242 in GI: 82796355, and an antisense strand primer, 5'-GCTTCTGAGCCCAGTGAAAC-3' (residues 10422576–10422595 in GI: 82796355). Cycling parameters for PCR amplification were: 98°C for 5 min and then 40 cycles of 98°C for 0.5 min, 55°C for 1 min and 72°C for 1 min. The expected amplicon size is 373 bp. To detect the targeted cubilin allele, PCR reactions were performed with an EGFP sense strand primer, 5'-CCTGAAGTTCATCTGCACCA-3' (residues 810–829 in GI:1377911), and EGFP antisense strand primer 5'-TGCTCAGGTAGTGGTTGTCG-3' (residues 1288–1269 in GI:1377911). Cycling parameters for PCR amplification were: 98°C for 5 min and then 40 cycles of 98°C for 0.5 min, 55°C for 1 min and 72°C for 1 min. The expected amplicon size is 478 bp.
For embryos from intercross heterozygous matings that were paraffin embedded within maternal tissue, the assignment of 'mutant' was based on two criteria: 1) by the absence of immunologically detectable cubilin (anti-cubilin T-16 from Santa Cruz Biotechnology, Santa Cruz, CA) in sections of the paraffin embedded embryos; and 2) the distinct mutant morphological phenotype corresponding to that of embryos confirmed by genotypic analysis to be homozygous nulls.
Immunohistochemistry and histology
Embryos were harvested from pregnant females following timed cubilin+/- intercross matings, and fixed for 40 min in 4% paraformaldehyde/PBS. Hematoxylin and eosin (H&E) staining of paraffin embedded embryo sections (7 μm) was performed using standard techniques. Whole-mount immunohistochemistry for the PECAM1/CD31 was performed as described previously [30]. Antibodies to PECAM (clone Mec-13.3) were purchased from BD PharMingen (San Diego, CA). For GFP and megalin immunohistological staining, sections were stained with rabbit antibodies to GFP purchased from Abcam (Cambridge, MA) or rabbit megalin cytoplasmic domain antibodies described previously [13]. Fluorescently conjugated secondary antibodies were purchased from Jackson ImmunoResearch Labs (West Grove, PA). Stained tissue sections were analyzed using a Leica DMR research-grade microscope equipped with Leica objectives and a SPOT-RT camera (Diagnostic Instruments, Sterling Heights, MI).
RT-PCR analysis
Total RNA was isolated from 8.5 dpc embryos from heterozygous matings using Trizol (Invitrogen). Template cDNA was prepared from 1 mg RNA using Superscript II Reverse Transcriptase (Invitrogen). To assess cubilin transcript levels two sets of PCR reactions were performed using a primer pair targeting the 5' end of the transcript and one targeting the 3' end. The 5' primer pair consisted of the forward primer 5'-ATGATGATGACCTTGGCGAATG-3' (residues 275–296 (exon 2) in GI: 94365995) and the reverse primer 5'-GCAGCCAAAGGGTGTTCCAG-3' (residues 587–606 (exon 6) in GI: 94365995). Cycling parameters for PCR amplification were: 98°C for 5 min and then 40 cycles of 98°C for 0.5 min, 58°C for 1 min and 72°C for 1 min. The expected amplicon size is 332 bp. The 3' primer pair consisted of the forward primer 5'-TCTCATACACCAACTACCCC-3' (residues 9221–9240 (exon 58) in GI: 94365995) and the reverse primer 5'-AGCAGTCTTGTGAGGGCAGC-3' (residues 10041–10060 (exon 62) in GI: 94365995). Cycling parameters for PCR amplification were: 98°C for 5 min and then 40 cycles of 98°C for 0.5 min, 58°C for 1 min and 72°C for 1 min. The expected amplicon size is 840 bp. To assess GAPDH transcript levels PCR was performed using a forward primer 5'-CGGTGTGAACGGATTTGGC-3' (residues 70–88 in GI: 47607489) and the reverse primer 5'-GCAGTGATGGCATGGACTGT-3' (residues 581–600 in GI: 47607489). Cycling parameters for PCR amplification were: 98°C for 5 min and then 40 cycles of 98°C for 0.5 min, 54°C for 1 min and 72°C for 1 min. The expected amplicon size is 531 bp. To assess b-actin transcript levels a forward primer 5'-CGGGACCTGACAGACTACCTC-3' (residues 627–647 in GI: 6671508) and the reverse primer 5'-AACCGCTCGTTGCCAATA-3' (residues 827–844 in GI: 6671508) were used. Cycling parameters for PCR amplification were: 98°C for 5 min and then 40 cycles of 98°C for 0.5 min, 55°C for 1 min and 72°C for 1 min. The expected amplicon size is 218 bp. To assess EGFP transcript levels a forward primer 5'-ACGTAAACGGCCACAAGTTC-3' (residues 743–762 in GI: 1377911) and the reverse primer 5'-AAGTCGTGCTGCTTCATGTG-3' (residues 910–929 in GI: 1377911) were used. Cycling parameters for PCR amplification were: 98°C for 5 min and then 40 cycles of 98°C for 0.5 min, 58°C for 1 min and 72°C for 1 min. The expected amplicon size is 187 bp.
Analysis of embryonic uptake of maternal derived HDL
Human DiI-labeled HDL (DiI-HDL) was purchased from Biomedical Technologies (Stoughton, MA). DiI-HDL was depleted of apoE-HDL and other heparin-binding particles according to Oram [31], dialyzed against 150 mM NaCl, 50 mM Tris pH 7.4 (TBS) containing 0.3 mM EDTA and filter-sterilized. Lipoprotein concentration was determined by BCA protein assay (Pierce, Rockford, IL). DiI-HDL was diluted into PBS to a final concentration of 0.265 mg protein/ml and 75–125 μl was infused into the saphenous vein of pregnant heterozygous mice at 8.0 dpc according to the procedure of Hem et al. [32]. One hour after infusion, embryos were isolated free of parietal endoderm and analyzed by confocal microscopy.
Abbreviations
VE, visceral endoderm; EE, embryonic endoderm; Am, amnion; HDL, high density lipoprotein; DiI-HDL, DiI-labeled HDL; apoA-I, apolipoprotein A-I; AMN, amnionless; Cbl, cobalamin; dpc, days postcoitum; H&E, Hematoxylin and eosin; EGFP, enhanced green fluorescent protein.
Authors' contributions
B. T. S. generated the targeting vector, carried out analyses of knockout mice, contributed to experimental design and helped draft the manuscript. J. C. M. carried out morphological and genotypic analyses of the null embryo phenotype, maternal HDL transport experiments and writing of the manuscript. P. A. F. assisted with embryo isolation, immunohistological analysis and embryo imaging. J. L. B. performed degenerate amplification of mouse cubilin 5' cDNA sequences, isolated and characterized the BAC used to generate the targeting vector and directed cubilin 5' RACE analysis. M. A. C. assisted with the design of the targeting vector. D. D. S. performed vector electroporation, selection of targeted ES cells, blastocyst injection and implantation. C. J. D. assisted with the phenotypic characterization of null embryos and writing of the manuscript. W. S. A. conceived of the study, contributed to experimental design and interpretation of results, and coordinated the project and writing of the manuscript.
Supplementary Material
Additional File 1
Mapping of cubilin transcription initiation sites. The file contains findings from 5' RACE experiments used to position the EGFP reporter sequence within the 5' UTR of exon 1 of the reporter knock-in/KO targeting construct.
Click here for file
Acknowledgements
This work was supported by NIH grants HL061873 and DE14347 (WSA), HL57375 (CJD) and CA109958 (DDS) as well as DOD contract 4400122478 (DDS). Brian Smith was a recipient of support from NIH postdoctoral training grant T32 HL007710. Jason Mussell was a recipient of support from NIH postdoctoral training grant T32 HL07260. We also acknowledge the MUSC Gene Targeting and Knockout Mouse Core for assistance with the development of the cubilin knockout mouse. | [
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16800892 | Conditional gene expression in the mouse using a Sleeping Beauty gene-trap transposon
Abstract
Background
Insertional mutagenesis techniques with transposable elements have been popular among geneticists studying model organisms from E. coli to Drosophila and, more recently, the mouse. One such element is the Sleeping Beauty (SB) transposon that has been shown in several studies to be an effective insertional mutagen in the mouse germline. SB transposon vector studies have employed different functional elements and reporter molecules to disrupt and report the expression of endogenous mouse genes. We sought to generate a transposon system that would be capable of reporting the expression pattern of a mouse gene while allowing for conditional expression of a gene of interest in a tissue- or temporal-specific pattern.
Results
Here we report the systematic development and testing of a transposon-based gene-trap system incorporating the doxycycline-repressible Tet-Off (tTA) system that is capable of activating the expression of genes under control of a Tet response element (TRE) promoter. We demonstrate that the gene trap system is fully functional in vitro by introducing the "gene-trap tTA" vector into human cells by transposition and identifying clones that activate expression of a TRE-luciferase transgene in a doxycycline-dependent manner. In transgenic mice, we mobilize gene-trap tTA vectors, discover parameters that can affect germline mobilization rates, and identify candidate gene insertions to demonstrate the in vivo functionality of the vector system. We further demonstrate that the gene-trap can act as a reporter of endogenous gene expression and it can be coupled with bioluminescent imaging to identify genes with tissue-specific expression patterns.
Conclusion
Akin to the GAL4/UAS system used in the fly, we have made progress developing a tool for mutating and revealing the expression of mouse genes by generating the tTA transactivator in the presence of a secondary TRE-regulated reporter molecule. A vector like the gene-trap tTA could provide a means for both annotating mouse genes and creating a resource of mice that express a regulable transcription factor in temporally- and tissue-specific patterns for conditional gene expression studies. These mice would be a valuable resource to the mouse genetics community for purpose of dissecting mammalian gene function.
Background
Derived from ancient salmonid fish sequences [1], the Sleeping Beauty (SB) transposon is a member of the Tc1/mariner superfamily of cut-and-paste transposable elements [2] and has been developed as a vertebrate transformation tool [3] and germline insertional mutagen [4,5]. We and others have shown that the SB transposon is highly active in the mouse germline and can generate heritable loss-of-function mutations that lead to detectable phenotypes [4-7].
Data accumulated from spontaneous and engineered mouse mutations suggests that a significant percentage of mouse genes are essential for early development. This has necessitated the creation of various genetic tools for conditional loss- or gain-of-function genetic studies. In Drosophila, the ability to regulate genes in a tissue- or temporally-specific manner has become one method to study the function of these genes. Likewise, the ability to control the expression of an essential mouse gene is one way to discover its function in tissues that are formed after lethal phenotypes are manifest. Temporal and spatial control of gene expression in mutant backgrounds can be used to determine when a gene product is required during a developmental process [8-10]. With this in mind, and taking lessons from the Drosophila literature on GAL4/UAS-based P-element transposon systems [11], we designed a SB transposon-based gene trap system that allows us to annotate the expression pattern and function of mouse genes.
Previous studies using the SB transposon system for generating insertional mutations in the germline of mice used various gene- and polyA-trap transposon vectors [4,5]. These vectors were designed to truncate an endogenous mRNA, incorporate coding sequences for a visible reporter like β-Galactosidase or green fluorescent protein (GFP), and report the expression patterns of trapped genes. If a transcriptional transactivator, similar to GAL4, were used in place of a visible reporter molecule, a small amount of the molecule could lead to high levels of a secondary reporter transgene under control of a responsive promoter. Analogous to the GAL4/UAS system, the tetracycline-controlled transactivator (tTA) and Tet-response element promoter (TRE) were developed for mammalian systems [12].
Here we report the creation of a mutagenic SB-based, gene-trap transposon vector that can insert into a mouse gene and express the Tet-off transcriptional transactivator (tTA) in its tissue- and temporally-regulated manner. The "gene-trap tTA" can mutate genes and record their expression patterns for functional annotation as well as create a mechanism for driving the expression of transgenes in a regulable manner in vivo. We also report several general parameters that impact the application of the Sleeping Beauty transposon system to mouse functional genomics including analysis of transposon mobilization rates in several different transgenic lines and the rescue of a mutant phenotype by remobilization.
Results
Creation of the gene-trap tTA system
A SB transposon-based gene-trap tTA vector was first designed for and tested in zebrafish embryos [13]. The gene-trap tTA vector was shown function in somatic tissues of zebrafish and activate a TRE-regulated GFP transgene. T2/GT2/tTA and T2/GT3/tTA (Fig. 1A) are similar to the earlier gene-trap tTA (GT/tTA) and encode a carp β-actin splice acceptor, the encephalomyocarditis virus (EMCV) internal ribosomal entry site (IRES), the Tet-Off® (tTA) coding sequence (Clontech, Palo Alto, CA), and the SV40 late polyadenylation signal. Modifications are described in the Materials and Methods and include an upgrade to the 'T2' versions of the SB inverted terminal repeat sequences (ITR), demonstrated to increase transposition activity in vitro [14], and the addition of stop codon sequences in each reading frame just upstream of the IRES. The T2/GT3/tTA transposon vector (Fig. 1A) was additionally modified to include a splice acceptor from the mouse hypoxanthine phosphoribosyltransferase gene (HPRT) and a V5-epitope tag on the opposite strand of the transposon. This allows the gene trap to disrupt gene expression upon insertion into a mouse gene in either orientation. If open reading frame one were conserved after splicing from the endogenous exon, the V5-epitope would be added to the truncated peptide.
Fig. 1B shows the method by which the gene-trap tTA can disrupt a mouse gene. When the gene-trap transposon lands in the intron of a gene, it will intercept splicing from an upstream exon and incorporate the IRES and tTA elements to form a bicistronic messenger RNA. Translation of the endogenous gene peptide is truncated at one of the stop codons incorporated just upstream of the IRES. The IRES element allows cap-independent translation initiation from the second open reading frame to produce tTA from the same mRNA. For the T2/GT2/tTA vector, an insertion into a gene in the opposite orientation is not predicted to mutate the gene, however, the T2/GT3/tTA would interrupt splicing with the HPRT splice acceptor, but will not express tTA.
Before making transgenic mice, we validated the functionality of the gene-trap tTA as a faithful activator of reporter gene expression in vitro. First, a G418-resistance cassette was cloned into the T2/GT2/tTA transposon vector to generate T2/GT2/tTA/SVNeo (Fig. 1C). We then generated a HeLa cell line harboring a TRE-regulated luciferase transgene. Upon co-transfection of the pT2/GT2/tTA/SVNeo transposon vector plasmid and a source of SB transposase, pCMV-SB [1], the gene-trap tTA transposes at random into the HeLa cell genome. The SVNeo cassette confers G418 resistance to cells that integrate and express the transposon vector, but the gene-trap tTA cassette can intercept splicing and express tTA only if the transposon lands in an expressed gene in the correct orientation. Fig. 1D shows the luciferase activation in sixty-two individual G418-resistant clone extracts. As a control, a transposon that contained the same G418 resistance cassette, but not the splice acceptor-IRES-tTA cassette, did not activate luciferase expression (Fig. 1D, white bars). As expected, only a subset of T2/GT2/tTA/SVNeo-transgenic clones, 17 out of 50, showed activation of TRE-luciferase above background levels.
The activity of tTA is regulable by administering the drug tetracycline or its analog doxycycline [15]. We tested this sensitivity by administering doxycycline to three of the TRE-luciferase activated clones from Fig. 1D. Fig. 1E shows the doxycycline-dependent repression of TRE-luciferase activation in each clone. Luciferase activity was reduced approximately two orders of magnitude in clones, 43 and 44, and about one order of magnitude in clone 39. Finally, to assure that gene trap tTA activation was dependent on splicing from an upstream exon of an expressed gene, 5' rapid amplification of cDNA ends (RACE) was performed with primers specific to the gene trap on mRNA extracts prepared from several clones. Additional file 1 shows the sequence tags, chromosomes and gene identification obtained for seven clones. In each case, an upstream exon of the gene is spliced into the gene-trap tTA at the predicted position (underlined). These data demonstrate that the tTA molecule can be expressed from the gene-trap tTA vector in the context of a trapped gene to activate a TRE-regulated transgene and that this activation is repressible by the addition of doxycycline. With the prospect of generating mouse strains with regulated gene-trap tTA activation, we set out to test the system in vivo.
Parameters affecting germline mobilization rates
Transgenic mice were generated with the T2/GT2/tTA and T2/GT3/tTA transposon vectors by standard pronuclear injection (see Materials and Methods). A typical result of this protocol is the head-to-tail concatenation of transgene units before integrating into the mouse genome to create a multi-copy array of transgenes in a mouse chromosome. Each transposon transgene concatemer, thus, serves as a donor site containing multiple substrates for mobilization by the SB transposase. To determine the number of gene-trap tTA transposons in each transgenic line, we performed Southern blot analysis using a restriction digest scheme that resolves the concatemerized transposons to a single band and estimated the number of copies based on its intensity (see Materials and Methods). Table 1 shows ten independent transgenic founder lines with transposon copy numbers ranging from approximately three to sixty copies. To test for mobilization of the transposons in the germline, each transgenic line was crossed to the CAGGS-SB10 transposase-expressing strain [16]. The resulting doubly transgenic "seed" mice were outcrossed to wild type animals to observe gene trap mobilization in the germline. Southern blot analysis on tail biopsy DNA from offspring of seed mice was used, as previously described [4,16], to observe the number of new transposon insertions per gamete. We averaged the insertion data from seed mice from each line to calculate the likelihood (rate per copy number) that a transposon could be mobilized from their respective concatemers (Table 1). The data demonstrates that lines with fewer copies of the transposon substrate show the lowest mobilization rates. This, however, is not a strict correlation. This is evident in comparing lines 6632 and 6657, where the former has fewer copies, but shows a higher per-copy-transposon mobilization rate of the latter.
The inability to mobilize transposons in transgenic mice with few transposons in the concatemer is consistent with our efforts to re-mobilize a single copy insertion in the genome. We analyzed the mobilization of two independently generated single copy insertions. Insertion 01-0032 was generated and described in an earlier report [4] and 02A-0016 is a gene-trap tTA insertion identified in an offspring of a T2/GT2/tTA seed male (line 4583) as described below. When SB transposons are excised from a chromosome locus during gametogenesis, host DNA repair machinery typically creates a "footprint" of a few or several base pairs while repairing the double-strand-break gap [17]. We mobilized the insertions 01-0032 and 02A-0016 by intercrossing doubly transgenic mice that harbor a single-copy transposon insertion and the CAGGS-SB10 transposase transgene (Fig. 2A). We detected transposition in about one out of every 100 offspring for either insertion by PCR amplifying the sequences that flank the donor site and sequencing to detect a footprint (Fig. 2B). Notably, remobilization of insertion 01-0032, which causes an early recessive embryonic lethal phenotype as a result of disruption of the mouse Slc25a22 gene [4], restores activity of the gene and rescues the recessive lethality associated with this insertion as heterozygous 01-0032/footprint mice are phenotypically normal mice. The inability to efficiently remobilize a single copy insertion has been reported elsewhere [6]. For this reason, we incorporated loxP sites into the T2/GT3/tTA vector (Fig. 1A), flanking the mutagenic portion of the gene-trap vector. Cre-mediated recombination in tissues and in the germline have been demonstrated and are significantly higher than the 1% mobilization rate we can achieve by remobilizing single copy insertions [18-20]. Thus, a T2/GT3/tTA gene trap disruption of a mouse gene might be rescued by expression of Cre recombinase in a tissue- or germline-specific manner.
Table 1 also demonstrates that germline mobilization rates are lower in the female germline when compared to their male siblings (dashed boxes). Finally, the solid box shows the difference in transposition rates between two sibling male seed mice, one hemizygous and one homozygous for the transposase transgene. While the SB10+/+ seed male did not show increased transposition over other independently generated seed mice for the 4563 line, there was an increase over its sibling SB10+/- seed male, suggesting that we may be able to increase transposition rates by increasing the transposase dosage in future studies. The analysis of several transposon-transgenic lines has thus revealed parameters that affect transposition rates, and suggest careful line selection will be required for mutagenic programs.
Sleeping Beauty-mediated gene insertion in vivo
T2/GT2/tTA line 4563 and T2/GT3/tTA line 6660 were useful to identify new transposon insertions in the offspring of seed mice. Techniques for cloning transposon-genomic DNA junction sequences have been described [1,4,21]. The chromosomal positions of thirty transposition events in offspring of seed mice from line 4563 were identified by querying the ENSEMBL online mouse genome database using the BLAST function. The T2/GT3/tTA line 6660 was used in a study of mutations induced specifically on mouse chromosome 11 and the insertion data from this line will be reported elsewhere (A.M.G., manuscript in preparation). Additional file 2 and additional file 3 show the data accumulated from the thirty T2/GT2/tTA events. Consistent with previous reports, the T2/GT2/tTA transposons exhibit local hopping, where a significant percentage of new insertions cluster in a particular region of the genome, and the insertions can land in genes [4,5]. Three-primer PCR [3] can readily be used to genotype carriers for any specific transposon insertion. Because we were interested in testing the function of the gene-trap tTA vector in the context of an endogenous mouse gene, carriers of insertions 02A-0001 and 02A-0002, both in mouse genes, (Fig. 3A) were propagated for further study.
Molecular characterization of gene-trap tTA function
Insertion number 02A-0001 is in the first intron of the carbonic anhydrase-12 gene (Car12), while 02A-0002 is inserted into intron 8 of the predicted novel gene ENSMUSG00000066992 (Fig. 3A). The top panel in Fig. 3B shows the expression pattern of mouse Car12 by reverse-transcriptase PCR (RT-PCR) on RNA extracts from several wild type tissues using primers in exons 1 and 5. If the gene trap is splicing properly, the IRES and tTA sequences should be fused to exon 1 of the Car12 mRNA. The second panel shows RT-PCR detection of endogenous Car12 and Car12-IRES-tTA bicistronic transcripts in RNA samples taken from a heterozygous carrier of insertion 02A-0001. Nested PCR was necessary to detect the fusion transcript in some tissues. Comparing the wild type expression pattern of Car12 mRNA and the fusion transcript, it is apparent that expression of the trapped mRNA is tissue specific as predicted. The fusion transcript can be detected in the heart of the carrier, but not wildtype heart RNA extracts. We attribute this to the extra sensitivity of the nested PCR and suspect that endogenous Car12 is expressed in the heart at a low level.
If the carp β-actin splice acceptor is efficient, it should truncate the endogenous gene mRNA after the nearest upstream exon. Total RNA extracts from wild type, and heterozygous or homozygous carriers of both insertions 02A-0001 and 02A-0002 were probed for expression of endogenous Car12 or ENSMUSG00000066992 transcripts respectively. Fig. 3C shows complete ablation of Car12 mRNA in the kidney and brain of a homozygous carrier of insertion 02A-0001. Expression of ENSMUSG00000066992, detectable by RT-PCR exclusively in mouse brain (data not shown), was not ablated in homozygous carriers of insertion 02-0002, however, suggesting that the carp β-actin splice acceptor does not work efficiently in each case.
Interestingly, Car12 mutant mice demonstrate reduced fitness, as an intercross of 02A-0001 carriers results in non-mendelian inheritance of the homozygous class (data not shown). The expression of the carbonic anhydrase (CA-XII) has been previously localized to the membranes of mouse kidney, colon, and testes [22]. The CA-XII peptide has been previously reported to be 46 kDA [23]. Fig. 3C shows the reduction of CA-XII expression in the heterozygous kidney and absence in the homozygous kidney. Antibody cross-reaction with a ~ 30-kDa peptide is consistent with cross-reactions previously seen in kidney extracts with this antibody [23], and serves as a loading control (arrowhead, Fig. 3C).
Tissue-specific activation of a TRE promoter-regulated transgene in vivo
The final in vivo test for the gene-trap tTA system is to determine whether the vectors can activate a TRE promoter-regulated transgene in a tissue-specific manner. We obtained the Tg(tetL)1Bjd/J strain of mice, transgenic for a TRE-regulated luciferase transgene, from Jackson Laboratories. These mice have been previously shown to respond to tissue-specific tTA expression in a doxycycline-dependent manner [24]. Luciferase activity assays from individual organ extracts from a 02A-0001; Tg(tetL)1Bjd/J doubly transgenic animals showed activation of luciferase in all tissues tested when compared to singly transgenic controls (data not shown). This result was surprising because the trapped gene, Car12, is not expressed in the liver (among other tissues) by RT-PCR (Fig. 3B). Insertion 02A-0001 is the result of local transposition, and thus is linked to the original concatemer of transposons. We reasoned that these linked T2/GT2/tTA transgenes present in carriers of this insertion might express tTA, therefore resulting ubiquitous luciferase expression when crossed to the Tg(tetL)1Bjd/J strain. To circumvent this issue, we focused on insertions that were not the result of local hopping and would segregate from other transposable elements. Insertions 02A-0002, 03A-0184, 03A-0217, and 03A-0241 (Figs. 3A and 4A) are the result of non-local transposition of T2/GT2/tTA or T2/GT3/tTA transposons in their respective transposon transgenic strains. Carriers of each of these gene insertions were crossed to the Tg(tetL)1Bjd/J strain of mice with the hope of visualizing tissue-specific, tTA-dependent transactivation of luciferase. As predicted, Fig. 4B shows weak activation of luciferase in the brain of a 02A-0002; Tg(tetL)1Bjd/J doubly transgenic mice, but not other tissues. This pattern was reproduced in three out of four mice. As shown above, this insertion does not completely disrupt the brain expression of ENSMUSG00000066992 (Fig. 3C), but some splicing into the T2/GT2/tTA splice acceptor must occur to allow tissue-specific expression of the tTA.
Hasan, et al. (2001) demonstrated that non-invasive imaging of luciferase expression of a similar tTA-activated strain using an intensified charge-coupled device camera system after subcutaneous injection of luciferin substrate [25]. We used the Xenogen IVIS™100 Imaging System to see if tissue-specific patterns could be detected in gene-trap tTA; Tg(tetL)1Bjd/J doubly transgenic mice. Light emissions are often observed in the extremities, nose, ears, and tails of control Tg(tetL)1Bjd/J in a tTA-independent manner in our hands (Fig. 5A). No additional tissue-specific signals were initially detected in 02A-0001; Tg(tetL)1Bjd/J or 02A-0002; Tg(tetL)1Bjd/J doubly transgenic mice, suggesting that expression of luciferase levels detected in the organ extracts were not sufficient to be detected by the imager. Likewise, when crossed to the Tg(tetL)1Bjd/J strain, insertions 03A-0184 and 03A-0217 (Fig. 4A) showed no activation of luciferase by imaging or by luminometer readings of organ extracts (data not shown). While the expression pattern of Membrane-spanning 4-domains, subfamily A, member 6C (Ms4a6c) is not characterized, Integrin beta 3 (Itgb3) has been localized to the developing mouse heart in early embryonic stages [26]. It is possible that we missed the expression of these gene-trap tTA insertions because these genes are not expressed in adult tissues.
In contrast, a striking pattern of luciferase expression was captured by imaging 03A-0241; Tg(tetL)1Bjd/J doubly transgenic mice (mouse 936) when compared to singly 03A-0241 transgenic (mouse 933) or Tg(tetL)1Bjd/J controls (Fig. 5A). Light emission patterns were detected from anatomical origins corresponding to the sternum, spleen, femurs and vertebrae, presumably reflecting luciferase activation in a component of the bone marrow and hematopoietic cells. The MacF1 gene has more than 100 exons and has been reported to be expressed in multiple isoforms in the mouse with high levels produced in the lung [27]. Using primers in exons that flank the 03A-0241 insertion to amplify MacF1 transcripts, Fig. 5B demonstrates that at least two splice variants of different sizes are variably expressed in several tissues, but MacF1 is not normally expressed in the bone marrow of a wild-type mouse. This suggests that the 03A-0241 insertion into MacF1 may cause abnormal expression of the gene in the bone marrow and hematopoietic cells. However, multiple attempts to amplify the chimeric MacF1-IRES-tTA transcript from bone marrow extracts from carrier mice by RT-PCR were unsuccessful in identifying this spliced transcript.
To determine whether this pattern was linked only to insertion 03A-0241 and the mouse MacF1 gene, Southern blots and genotyping PCR were performed on mouse 936 (and siblings 931–935) and his offspring (mice 949–976) (Fig. 5C). Surprisingly, mice that inherited insertion 03A-0241 by PCR showed linkage to the concatemer donor site by Southern blot (Fig. 5A, arrowhead). This was unexpected because these mice were generated from T2/GT3/tTA line 6660 (Table 1), which has a transposon donor site on mouse chromosome 11. MacF1 is located on mouse chromosome 4 at 57.4 cM, thus the Southern data suggests that a large portion of the chromosome-11 concatemer had translocated by some mechanism to chromosome-4. Thus, the MacF1 insertion on chromosome 4 is still genetically linked to a translocated donor concatemer of multiple T2/GT3/tTA transposons in addition to other linked single-copy insertions (Fig. 5C, arrows). Repeated attempts to identify these other linked chromosome-4 insertions by our cloning techniques and 5'RACE for trapped sequences in bone marrow extracts were unsuccessful in identifying the origin of this tissue-specific gene-trap tTA expression. Nevertheless, the pattern seen in 03A-0241; Tg(tetL)1Bjd/J mice is unique to these mice, as it is not seen in carriers of other T2/GT2/tTA or T2/GT3/tTA insertions (with or without associated concatemers) when crossed to the Tg(tetL)1Bjd/J strain (above and data not shown).
The tissue-specific luciferase pattern in carriers of this insertion could be reproducibly transmitted through the germline to offspring (mice 962–964 and 971–973, Fig. 5D). The upper range of luciferase activation from each offsrpring, measured in photons/second/cm2, ranged approximately a thousand-fold (scale bars). We attribute these differences to differences in the levels of cap-independent translation of the tTA molecule from the IRES in each mouse. Finally, other groups have demonstrated in vivo sensitivity of tTA-dependent activation of gene expression to tetracycline, or its analog doxycycline [24,28,29]. We determined whether we could control gene-trap tTA activation of luciferase by administering doxycycline to the drinking water of the 03A-0241; Tg(tetL)1Bjd/J mice. Fig. 5E demonstrates absence of luciferase signal in the same animals from Fig. 5D, after six days of treatment, suggesting complete repression of the tTA-dependent luciferase activation.
Discussion
We examined the utility of a SB transposon-base gene trap vector that expresses the Tet-off transcriptional transactivator in the patterns of trapped genes in vitro and in vivo. The gene-trap tTA demonstrated full utility in cultured cells, having the ability to insert in genes, trap the splicing of those genes to express the tTA transcription factor which, in turn, could activate a TRE-regulated transgene in a doxycycline-dependent manner. Likewise, analysis of gene-trap tTA transposon insertions into mouse genes revealed that the vector system can completely disrupt the expression of a mouse gene in some cases and can tissue-specifically activate expression of a TRE promoter-regulated transgene in a doxycycline-dependent manner in vivo. While ultimately this vector system can function as designed, several observations suggest we need a more optimal approach for creating a resource of mice that express tTA in developmentally-controlled patterns.
Germline SB transposon mobilization rates in the mouse germline are variable and ranged from less than one in twenty to nearly three events per gamete. An evaluation of several different transposon-transgenic lines has given us insight into the potential the SB system to perform regional mutagenesis throughout the genome. The genomic location of the transposon donor site, along with copy number, impacts the germline mobilization rate from any concatemer (see Table 1). This suggests that not all parts of the genome are equally accessible to SB transposition. Understanding the mechanisms behind this "mobilization position effect" will likely require an increased understanding about molecular regulators of SB transposition, but may include a well known director of position effect, heterochromatinization [30].
As demonstrated in previous studies and here, SB transposon-based gene trap vectors can function in vivo to disrupt the expression of mouse genes. The T2/GT2/tTA vector can completely ablate the expression of a mouse gene as demonstrated by the null mutation that was generated in the mouse Car12 gene. Not all gene insertions completely disrupt gene expression however, as in the case of insertion 02A-0002 into ENSMUSG00000066992. While hypomorphic mutations may be valuable to discover the function of an essential mouse gene, this has lead to ideas on how to improve the efficacy of future vectors. Splice acceptors derived from different genes, termination sequences (polyadenylation signals) and other functional sequences have been used in several ES cell plasmid- and retroviral-based gene-trapping studies with varying efficiencies [31-33]. Drawing data from other groups will lead to improved vectors for trapping mouse genes and perhaps lead to vectors to trap certain classes of genes [34].
The activation of luciferase by the tetracycline-controlled transactivator varied rougly 1000-fold (Fig. 5D) when expressed from the IRES in the gene-trap tTA. Once again, improvements to the functional components of the gene trap vector may lead to more reliable and greater expression of the transactivator. Alternative IRES elements like the 9-nucleotide IRES element from the murine Gtx homeodomain gene [35], or eliminating the IRES entirely, may allow for more consistent tTA expression.
The brilliant pattern of in vivo luciferase expression seen if Fig. 5 was initially thought to originate from insertion in the MacF1 gene. Published reports suggested that neither the bone marrow, nor hematopoietic cells are in the normal expression domain of this gene. Upon further molecular characterization, we discovered that this gene was not spliced into the gene-trap tTA, and by Southern blot, found it was tightly linked to other insertions as well as the concatemer. It seems more likely that the pattern of tTA-dependent luciferase activation seen in carriers of the 03A-0241 insertion comes from one of these linked transposons. Efforts to find the source of this expression by 5'RACE were unfortunately unsuccessful. We propose that in the case of insertion 03A-0241, the concatemer translocated to chromosome 4 first, then a single copy of T2/GT3/tTA hopped locally into MacF1. We have observed similar and other types of genomic rearrangements associated with transposition from the chromosome-11 donor site in other mice (A.M.G., manuscript in preparation). These observations would suggest that gene-trap tTA mobilization in the germline may not be the most efficient means of generating a resource of mice with tissue-specific tTA expression patterns. It is not clear, however, that these large donor site insertions are not rare events that could be tolerated. Independent insertions, unlinked to the concatemer or other transposons, are frequently recovered. By pre-selecting mice that do not inherit the concatemer by Southern blot or PCR analysis, a screen would largely avoid the complications of linked transposons.
An alternative and relatively facile approach would be to inject a linearized plasmid harboring the gene-trap tTA transposon, along with in vitro-transcribed transposase messenger RNA, into one-cell mouse embryos. In the one-cell embryo, the transposase mRNA is translated and transposase catalyzes integration into the mouse genome at rates that are two to three-fold higher than standard pronuclear injection of naked DNA [3]. This method produces transgenic mice with insertions distributed randomly across the genome, and multiple linkage groups are frequently obtained from a single injection [3]. This would be another method to avoid the complications of linked insertions and concatemers.
Finally, using the GAL4/UAS enhancer trap systems used in the fly, expression-based screening has been useful for identifying genes with similar or overlapping expression patterns [36] and for tissue-specific overexpression studies [37]. As we have shown here, a transgenic mouse expressing firefly luciferase under the control of the tet-response enhancer/promoter (TRE) can be coupled with the gene-trap tTA system to identify in vivo patterns with an intensified CCD camera system. In a screen, this type of imaging could be used to identify gene trap insertions that lead to tTA expression in a particular tissue or developmental stage, even in utero [38,39]. Alternatively, mouse strains for other TRE-regulated transgenes have been created, allowing for tTA-dependent expression of β-galactosidase [40] or GFP [41]. If the efficiency of the gene-trap tTA transposon can be improved, these strains could be used by any lab to identify tissue-specific patterns.
Conclusion
As a tool for the mouse community, introducing a gene-trap tTA vector into mouse genes with different spatial and temporal expression patterns would create a valuable tool for the mouse genetic toolbox. Tissue-specific promoters can control the expression of transactivators like tTA and rtTA (reverse tetracycline-controlled transactivator) in transgenic animals [42-44]. Having tissue-specific and temporal control of gene expression in any mouse tissue at any developmental stage could allow for the dissection of biological gene functions that were previously masked by a phenotype such as lethality. An SB-based transposon vector is well suited to accomplish this task by introducing precisely integrated functional reporter units into the mouse genome. In addition, an improved gene-trap tTA could provide more reliable expression of the transactivator than a transgene construct since transcription is regulated in the context of an endogenous gene and is thus less likely to be subject to the position effect variegation that can hinder the expression of transgenes.
Methods
Cloning gene-trap tTA transposon vectors
All primer/oligo sequences for PCR and cloning are provided in Additional file 4. All gene-trap tTA vectors are descendent of pGT/tTA, originally described in Clark et al . 2004 [13]. An artificial exon (based on carp β-actin exon 2), with stop codons in each reading frame, was created by annealing four oligos (AMG049-AMG052), filling in the gaps with Klenow DNA polymerase, ligation with T4 DNA ligase, PCR amplifying with AMG049 and AMG051, and subcloning the AgeI fragment into the partially-digested SmaI-AgeI fragment of pGT/tTA to create pGT2/tTA. Subcloning the exonuclease-treated EcoRV-PstI fragment of pGT2/tTA into the Klenow-treated blunt HindIII fragment of pT2/HindIII [14] generated pT2/GT2/tTA. pT2/GT2/SVNeo was created by subcloning the Klenow-treated HindIII fragment of pT2/SVneo [14] into the EcoRV fragment of pT2/GT2/tTA.
The construction of pT2/GT3/tTA was a multi-step process to add additional features to the GT2 version. First, an NheI site was added to pT2/GT2/tTA by PCR amplifying the entire plasmid with primers AG023 and AG024, containing a 12-bp overlap containing the NheI site sequence, and transformation. E. coli repairs the plasmid by homologous recombination in the overlapped region, resulting in the plasmid pT2/GT2/tTA/NheI. A loxP site was made by annealing two oligos (AG025 and AG026) containing the 34-bp loxP sequence with overhanging 4-base cohesive ends to NheI and cloned into the NheI site of pT2/GT2/tTA/NheI, destroying the NheI site on one side, and adding the loxP sequence to make pT2/GT2/tTA/loxP. The NheI-SpeI fragment of pT2/GT2/tTA/loxP was subcloned into the XbaI fragment of the transposon vector pT2/HB, which has a multiple cloning site, to create pT2/GT2/tTA/loxP-2. 70-mer oligos (SA/V5-1 and SA/V5-2) containing the HPRT splice acceptor and V5 epitope tag with EagI compatible ends were annealed and cloned into the EagI site of pT2/GT2/tTA/loxP-2 to make pT2/GT2/tTA/loxp/SA-V5. In the process, one side of the EagI site was destroyed, leaving a single EagI site. A 32-bp transcriptional termination sequence from the human GASTRIN gene was created by overlapping oligos (Termin-1 and Termin-2) and cloned into this EagI site of pT2/GT2/tTA/loxp/SA-V5 to make pT2/GT2/tTA/loxp/SA-V5/Termin. This sequence was shown to enhance transcriptional termination in plasmid vectors when downstream of polyadenylation signals [45], and was included to increase the chances of mRNA truncation, and thus mutation, in the gene trap. Finally, a second loxP site was made by annealing the same oligos above and ligation into the SpeI site of pT2/GT2/tTA/loxp/SA-V5/Termin to make the final product, pT2/GT3/tTA.
Generation of Tre-Luciferase/Puro cell line and in vitro gene trap testing
pTRE-Luc was kindly provided by Dr. Perry Hackett's lab and was co-transfected with the plasmid pKO Select-Puro (Stratagene, La Jolla, CA) into HeLa cells using Mirus TransIT-LT1 reagent (Promega, Madison, WI). After puromycin resistant clones were isolated, ten clones were tested for inducibility by pTet-off (Clontech, Palo Alto, CA). One clone TLP-10 showed a four-log increase in luciferase expression over background in the presence of transfected pTet-off (data not shown), and was subsequently used for the in vitro studies presented here.
pT2/GT2/tTA/SVNeo was transfected into the TLP-10 cell line along with pCMV-SB [1], and clones were selected in 800 μg/mL G418. pT2/SVNeo [14] was used as a control. Fifty of these clones and twelve control clones were grown to confluency on 60 mm plates and cells harvested with Promega's Cell Culture Lysis Reagent for luciferase assays. Luciferase assays were performed on 20 μL of lysis extract using 100 μL of Promega's Luciferase Assay Substrate and a Lumat LB 9507 luminometer (Berthold, Bundoora, Australia) with a 15 second measuring time. Relative light unit (RLU) measurements were normalized to total protein concentration as determined by Bradford assay. Doxycycline (Sigma cat. #D-9891) was added to a final concentration of 2 mM to repress tTA-induction of TRE-Luciferase in clones 39, 43, and 44.
RT-PCR and 5' rapid amplification of cDNA ends (RACE)
All primer/oligo sequences for PCR are provided in Additional file 4. Total RNA from cultured cells or mouse tissues was extracted with Trizol® (Invitrogen, Carlsbad, CA). RT-PCR was performed using the one-step RobusT™ I RT-PCR Kit (Finnzymes, Espoo, Finland) according to the manufacturer's protocol. A template-switching reaction with Superscript III (Invitrogen) was used to make template for 5'RACE on cultured cell or mouse tissue RNA extract. 250 ng total RNA was mixed with 1 μM each of primers GT2-RT3 and RACE-1, along with 0.01 M dithiothreitol, 500 μM dNTPs, and 1 unit of Superscript III. The mixture was cycled six times at 52°C for 10', 50°C for 15', and then gradually cooled from 50°C to 37°C over two minutes before a final extension for 90" at 37°C. Template was diluted ten-fold in water and 2 μL was used for primary 5'RACE PCR. Briefly, the template was amplified in a 50 μL PCR reaction supplemented with primers GT2-RT3 (0.4 μM), KJC-002 (0.04 μM) and KJC-003 (0.1 μM), 200 μM dNTPS, 2 mM MgCl2, and 1 unit of Taq DNA polymerase (CLP, San Diego, CA). The PCR machine was programmed for touchdown PCR at 95°C for 3', 10 cycles of 95°C for 30", 65°C for 30" (-0.5°C per cycle), 70°C for 2', and then 25 cycles of 95°C for 30", 60°C for 30", and 70°C for 2'. The primary RACE reaction was diluted 1:50 and 2 μL used in a nested PCR under the same exact conditions, except supplemented with 0.5 μM of each primer GT2-RT2 and KJC-004.
Generation of T2/GT2/tTA and T2/GT3/tTA mice by pronuclear injection, fluorescent in situ hybridization, and Southern blotting
The 3379-bp FspI -SapI fragment of pT2/GT2/tTA or the 3738-bp FspI -SapI fragment of pT2/GT3/tTA were gel-purified, ethanol precipitated twice, and resuspended in 5 mM Tris-Cl (pH 7.5), 0.1 mM EDTA for pronuclear injection into the FVB/N strain of mice (Charles River Laboratories, Wilmington, MA). The pT2/GT3/tTA fragment was co-injected with the similarly-prepared HinP1I fragment of the plasmid pTYBS [Overbeek, 1991 #211], a tyrosinase minigene construct kindly provided by Dr. Paul Overbeek, in an attempt to coat-color mark animals that inherit the transposon/tyrosinase minigene transgenic insertion site. Expression of the tyrosinase, for unknown reasons, was not evident in any of the founders (data not shown). FISH on T2/GT3/tTA transgenic mouse lines was performed on splenic lymphocytes using standard techniques, and performing nick translation to label the pT2/GT3/tTA plasmid or whole BAC DNA from the Wellcome Trust Sanger Institute [46] were used as a probe. Standard Southern blotting techniques were used to analyze BamHI-digested genomic DNA from founders and subsequent generations using the 842-bp NcoI -SphI fragment of the tTA open reading frame as a probe. BamHI restricts each copy in any T2/GT2/tTA or T2/GT3/tTA concatemer to a single band at 2124-bp or 2236-bp respectively (see Fig. 1A). Transposons mobilized from the concatemer to a genomic site are detected as bands of variable size which are larger than the concatemer band, their size determined by the nearest genomic BamHI site (see Fig. 5C).
Western blotting for CA-XII expression
Whole kidney extracts were probed with a polyclonal CA-XII antibody kindly provided by the laboratory of Dr. William Sly. 50 μg total protein was run on a 10% SDS-PAGE (Invitrogen) and transferred to PVDF and probed with a 1:1000 dilution of anti-CAXII primary antibody followed by a 1:10000 dilution of horseradish peroxidase-conjugated Anti-rabbit IgG secondary (Amersham Biosciences) in TBST (10 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.1 % Tween 20) supplemented with 1% dry milk.
In vivo luciferase imaging and analysis of luciferase expression, doxycycline treatment
Transgenic mice were imaged as described in Wilber, et al . (2005) [47], using luciferin from Xenogen Corporation (Alameda, CA). For luciferase assays on organ extracts, mice were sacrificed, and whole organs dissected and homogenized in Promega's Cell Culture Lysis Reagent. Extracts were analyzed as described above for the in vitro work. Drinking water was supplemented with 5 mg/mL Doxycycline and 0.1% sodium saccharine for 6 days to suppress tTA-induced activation.
Authors' contributions
AMG performed the updated vector construction, in vitro testing of the gene-trap tTA, preparation of transgenes, breeding and characterization of transgenic mice, 5'RACE, RT-PCR, western blotting, Southern blotting, and aided the in vivo imaging studies. AW performed the in vivo imaging for luciferase expression. CMC bred and characterized 'rescue by remobilization' for insertion 01-0032. PDL performed PCR-based methods to identify gene-trap tTA insertions in transgenic mice. KJC originally designed the gene-trap tTA system and was instrumental to upgrading its design and functionality. PBH, RSM, and DAL are not only the mentors of the above authors, but were involved in the design and execution of this manuscript.
Supplementary Material
Additional File 1
T2/GT2/tTA/SVNeo clone 5'RACE sequence and gene identification Splicing events of genes into the gene-trap tTA were identified by 5'RACE PCR and sequencing. For seven clones, the partial sequence tag demonstrates splicing into the proper branch point of the carp β-actin splice acceptor, allowing identification of the trapped gene by the sequence of the adjoined endogenous exon sequence. These genes and their functions are described according to the ENSEMBL May 17, 2005 freeze of the NCBI m34 build.
Click here for file
Additional File 2
Germline T2/GT2/tTA insertions The genomic positions of thirty T2/GT2/tTA insertions identified in the offspring of seed mice from line 4563 (Table 1) are reported. Shown are the chromosome and position, along with the identification and description of known or predicted genes potentially disrupted for each insertion. This data is based on the ENSEMBL May17, 2005 freeze of the NCBI m34 build.
Click here for file
Additional File 3
Distribution of thirty T2/GT2/tTA insertions Cloning insertions from offspring of seed mice from transgenic line 4563 (Table 1) reveals two local hopping intervals, one on mouse chromosome 1 near 45.8 Mb, and a second on mouse chromosome-9 around 66.5 Mb. By Southern blot, it was later determined that the 4563 line of mice originally obtained and segregated two independent concatemer integrations during the initial transgenesis (data not shown)
Click here for file
Additional File 4
Primer sequences
Click here for file
Acknowledgements
The authors would like to thank Dr. Abdul Waheed (Saint Louis University) for providing the anti-CAXII antibody in addition to all members of the Arnold and Mabel Beckman Center for Transposon Research for continuous dedication to stimulating discussion. The authors were supported by N.I.D.A. grant R01-DA014764 (AMG, DAL), N.I.H. grant T32 HD007480 (AMG), and N.I.G.M.S. grant T32 GM08347 (AW).
Figures and Tables
Figure 1
Gene-trap tTA vector design and in vitro testing A). The T2/GT2/tTA and T2/GT3/tTA vectors. The 'GT2' version is capable of mutating genes in one orientation while the 'GT3' version can mutate genes in both orientations upon insertion into a gene. (B- BamHI sites ) B) Compared to normal expression (left), when the T2/GT2/tTA SB transposon-based gene-trap vector inserts into a gene in the direction of transcription (right), endogenous splicing incorporates the IRES and tTA sequences and the bicistronic mRNA is prematurely truncated at the SV40 late polyadenylation site. The bicistronic mRNA allows cap-independent translation of the tTA molecule in addition to any peptide encoded by upstream exons. C) A stable, TRE-regulated luciferase cell line was created to test the T2/GT2/tTA/SVNeo transposon vector. After co-transfection with a plasmid source of transposase, G418R clones were individually expanded for analysis. D) Individual luciferase expression levels of G418R clone cell extracts from twelve control (pT2/SVNeo) and fifty gene-trap tTA clones from C. E) Incubation of three clones from C in media supplemented with 2 mM doxycycline results in 10- to 100-fold reduction of luciferase expression.
Figure 2
Phenotypic rescue by transposon mobilization A). Breeding scheme to remobilize a transposon insertion, 01-0032, out of the mouse Slc25a22 gene as previously reported by Carlson, et al . (2003). Animals for both the single-copy transposon insertion 01-0032 and the CAGGS-SB10 transposase were intercrossed. B) Germline single-copy remobilization rates of the independently generated insertions 01-0032 and 02A-0016 (Additional file 2) were detected by analyzing the donor site for evidence of a transposon footprint by sequencing (data not shown).
Figure 3
Molecular characterization of gene-trap tTAfunction A). Insertions 02A-0001 in the mouse Car12 gene and insertion 02A-0002 in the mouse ENSMUSG00000066992 gene. Genes are shown in the orientation of transcription from left to right, vertical lines are exons, along with the position and orientation of the T2/GT2/tTA insertion (arrows). B) RT-PCR detection of Car12 (top) and Car12-IRES-tTA chimeric (bottom) transcription in wild type and 02A-0001 carrier tissues. Kd-kidney, Br-brain, Lv-liver, Te-testes, Ad-adipose, Co-colon,SI-small intestine, Ov-ovary, Mu-muscle, Ht-heart C) RT-PCR analysis of gene disruption in tissues from wild type (+/+), heterozygous (+/ins), and homozygous (ins/ins) carriers of insertions 02A-0001 and 02A-0002. C) Western analysis of CA-XII expression in a wild type, heterozygous and homozygous carrier of insertion 02A-0001. Cross-reaction of the CA-XII antibody with other isoforms serves as a loading control (arrowhead).
Figure 4
T2/GT3/tTA gene disruptions and tissue-specific activation of a tet-responsive transgene in vivo A). T2/GT3/tTA insertions into mouse genes not linked to the original donor site. The genes are shown in the orientation of transcription with vertical lines as exons (from left to right) with the position and orientation of the IRES-tTA trapping cassette (arrow). B) Luciferase assays performed on individual tissues from four independent 02A-0002; Tg(tetL)1Bjd/J doubly transgenic and control Tg(tetL)1Bjd/J mice.
Figure 5
In vivo imaging of gene-trap tTAactivation of a tet-responsive transgene A). Luciferase emission pattern seen in a 03A-0241; Tg(tetL)1Bjd/J doubly transgenic mouse (936) as imaged by an intensified charge-coupled device (CCD) camera. Ventral and dorsal aspects are shown. The intensity of luciferase expression is compared in photons/second/cm2. B) RT-PCR analysis of MacF1 expression in multiple adult tissues, Gapdh was used as a control for sample quality. ES-embryonic stem cells, Lv-liver, Kd-kidney, Br-brain, Lu-lung, Ht-heart, Sp-spleen, St-stomach, BM-bone marrow, SI-small intestine, Co-colon, Ad-adipose, Mu-muscle, Ov-ovary, b-testes, Th-thymus C) Southern analysis of mice from A and B (see Materials and Methods). The donor site concatemer appears as an intense BamHI fragment at 2236 base pairs (arrowhead). The detection of insertion 03A-0241 by three-primer PCR is shown below each lane. Three bands marked by arrows segregate with insertion 03A-0241 genotyping and the concatemer.D) Pattern inheritance by offspring of mouse 936. Mice were imaged and scaled to different ranges of intensity that ranged from 1x104 to 1×108 photons/second/cm2 (scale bars). E) Repression of in vivo luciferase activation after six days of treatment with doxycycline in the same mice from D.
Table 1
Mobilization of gene-trap tTA transposons in the germlines of seed mice.
a Transposon copy number for each transgenic line was determined by Southern blot (see Materials and Methods)
b The number of single-copy insertions as determined by the presence of new bands on a Southern blot after mobilization by transposase in the germline of seed mice
c Reported is the average hits/gemete divided by the copy number | [
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16968134 | Inhibition of Gap Junction Communication at Ectopic Eph/ephrin Boundaries Underlies Craniofrontonasal Syndrome
Abstract
Mutations in X-linked ephrin-B1 in humans cause craniofrontonasal syndrome (CFNS), a disease that affects female patients more severely than males. Sorting of ephrin-B1–positive and –negative cells following X-inactivation has been observed in ephrin-B1+/− mice; however, the mechanisms by which mosaic ephrin-B1 expression leads to cell sorting and phenotypic defects remain unknown. Here we show that ephrin-B1+/− mice exhibit calvarial defects, a phenotype autonomous to neural crest cells that correlates with cell sorting. We have traced the causes of calvarial defects to impaired differentiation of osteogenic precursors. We show that gap junction communication (GJC) is inhibited at ectopic ephrin boundaries and that ephrin-B1 interacts with connexin43 and regulates its distribution. Moreover, we provide genetic evidence that GJC is implicated in the calvarial defects observed in ephrin-B1+/− embryos. Our results uncover a novel role for Eph/ephrins in regulating GJC in vivo and suggest that the pleiotropic defects seen in CFNS patients are due to improper regulation of GJC in affected tissues.
Introduction
Physical segregation, or sorting, of different cell populations during development is essential for the proper spatial organization of the animal body. Eph receptor tyrosine kinases and ephrins regulate many developmental processes [1–3], and play an important role in tissue patterning by restricting cell intermingling and establishing developmental boundaries [4–6].
A dramatic example of the role of Eph and ephrins in cell sorting, as well as the importance of proper cell sorting during development, was recently provided by the analysis of the phenotypes exhibited by ephrin-B1 heterozygous female mice [7,8] and by the identification of mutations in the ephrin-B1 gene in human craniofrontonasal syndrome (CFNS) patients [9,10]. As a result of random X-inactivation, X-linked ephrin-B1 expression is mosaic in ephrin-B1+/− mice and ephrin-B1–positive and ephrin-B1–negative cells segregate from one another. This correlates with a polydactyly phenotype that is never observed in ephrin-B1 null animals [7,8]. Similarly, CFNS is an X-linked developmental disorder characterized by a number of craniofacial defects including abnormal development of the cranial and nasal bones, and craniosynostosis (premature fusion of the coronal sutures), as well as extracranial anomalies (including polydactyly and syndactyly), affecting mainly female patients [11].
Despite years of intensive studies, the molecular mechanisms by which Eph receptors and ephrins influence cell sorting are still poorly understood. Eph receptors and ephrins function as an unusual receptor/ligand pair in which both receptor and ligand are capable of activating a signaling cascade. One prominent outcome of Eph/ephrin interactions is the regulation of cell–substrate adhesion and reorganization of the actin cytoskeleton. It has also been reported that Eph receptors and ephrins regulate gap junction communication (GJC) [6]. Indeed, studies in zebrafish have shown that expression of Eph receptors and ephrins in animal cap cells was sufficient to block GJC at the boundary between both cell populations. Gap junctions are intercellular membrane channels that mediate cell coupling by allowing the passage of small molecules directly from cell to cell. During vertebrate development, regions of GJC coincide with developmental compartments [12,13]. For instance, GJC is reduced at inter-rhombomeric boundaries, as compared to GJC in the rhombomeres themselves. GJC is involved in various developmental processes and mutations in connexins, the structural proteins forming gap junctional pores, have been linked to a number of human diseases [14]. Notably, GJC plays an important role in skeletal development [15] and mutations in connexin43 (Cx43) lead to cranial and skeletal defects both in humans and mice [16,17].
In this report, we have investigated the underlying cause of the phenotypes observed in ephrin-B1 heterozygous mice. We show that cell sorting in ephrin-B1+/− females induces calvarial defects due to the impaired differentiation of neural crest cells (NCCs). We provide evidence that GJC is inhibited at ectopic ephrin boundaries and that ephrin-B1 physically interacts with Cx43 and influences its distribution. In addition, we report that overexpression of Cx43 partially rescues the calvarial defects observed in ephrin-B1 heterozygotes. Finally, we show that regulation of GJC correlates with cell sorting in response to Eph/ephrin interaction. From these results we conclude that mosaic loss of ephrin-B1 exerts a dominant effect during development involving perturbation of GJC at ectopic Eph/ephrin boundaries and leading to defective tissue differentiation. These observations extend our understanding of the mechanisms underlying CFNS in humans and of the role of Eph receptors and ephrins in vivo.
Results
Craniofacial Phenotypes in ephrin-B1 Heterozygous Females
We and others have shown previously that ephrin-B1+/− females exhibit a polydactyly phenotype that is never seen in ephrin-B1Y/− males or ephrin-B1−/− females [7,8]. Because CFNS human female patients exhibit numerous craniofacial defects, we undertook a closer analysis of ephrin-B1 heterozygous female mice that revealed several defects in NCC-derived tissues that were not seen in hemizygous males or homozygous females. At P1, ephrin-B1+/− heterozygous females (n = 17) presented an opening (foramen) between the frontal bones and jagged bone fronts (Figure 1Ab), indicative of abnormal development of the frontal bones. (The parietal bone was affected at lower penetrance). Ephrin-B1−/− homozygous null females (n = 3) and ephrin-B1Y/− hemizygous males (n = 13) exhibited a normal development of these bones (Figure 1Aa and unpublished data). The frontal bone phenotype was recapitulated in heterozygous females carrying a deletion of ephrin-B1 specifically in NCCs (n = 3) (Figure 1Ac), consistent with the NCC origin of frontal bones [18]. Ephrin-B1+/− females also exhibited abnormal alignment of the vibrissae buds which was recapitulated by conditional deletion of ephrin-B1 in NCCs (ephrin-B1+/lox;Wnt1Cre+), indicating that the defect is autonomous to this lineage (Figure S1).
It has been proposed by others that the mild manifestations of CFNS in male carriers might be due to compensatory mechanisms by other ephrins [10]. To test this hypothesis, we asked whether further alteration in the dosage of ephrin signaling would lead to calvarial phenotypes in ephrin-B1 null mutants. We chose to focus on ephrin-B2 since loss of this gene has been shown previously to affect migration of a sub-population of cranial NCCs [19] and ephrin-B2 is expressed in the craniofacial area (Figure 2) We generated a mouse line harboring a null mutation in the ephrin-B2 gene by placing a cDNA coding for a fusion protein between histone 2B (H2B) and the green fluorescent protein (GFP) under the control of the ephrin-B2 promoter (A. Davy, P. Soriano, unpublished data). ephrin-B2 and ephrin-B1 heterozygous animals were mated to generate ephrin-B1/ephrin-B2 double heterozygous animals. Skeleton preparations showed that removal of one copy of ephrin-B2 in an ephrin-B1+/− background worsened the calvarial phenotype (Figure 1B). Indeed, in addition to a larger gap between the frontal bones (Figure S2A), ephrin-B1+/−/ephrin-B2+/GFP embryos (n = 5) exhibited a coronal suture defect that was not observed in ephrin-B1 single mutants, males or females. Bone fronts at coronal sutures in ephrin-B1+/−/ephrin-B2+/GFP embryos did not overlap, and ectopic bone was frequently observed in the suture mesenchyme (Figure 1Bc). Examination of skeleton preparations from E15.5 embryos indicated that although bone formation in ephrin-B1+/−/ephrin-B2+/GFP embryos proceeded comparably to ephrin-B1+/− embryos, bone fronts seemed unable to extend toward each other at the coronal suture (Figure 1Bf). Importantly, ephrin-B1Y/−/ephrin-B2+/GFP double hemi/heterozygous males (n = 3) exhibited normal calvarial development (Figure 1Bd and unpublished data). Altogether, these results show that mosaic loss of ephrin-B1 exerts a dominant effect on the development of calvarial bones. Removal of one copy of ephrin-B2 in an ephrin-B1+/− background was sufficient to uncover additional defects at the coronal suture; however, it did not lead to calvarial phenotypes in ephrin-B1 null embryos, suggesting that the lack of calvarial phenotype in these embryos is not due to compensation by ephrin-B2.
Defects in NCC-Derived Structures Correlate with Cell-Sorting in ephrin-B1+/− Embryos
To better understand the basis for the calvarial phenotypes observed in ephrin-B1 heterozygotes, we analyzed the expression pattern of ephrin-B1 and ephrin-B2, and their cognate receptors EphB2 and EphB3 at various stages of development. These receptors have been implicated both in the polydactyly phenotype and in the formation of the palate [20], a NCC-derived structure that also requires ephrin-B1. At E14.5, expression of ephrin-B1, EphB2, and EphB3 can be detected in the developing frontal bones (Figure 2A). Ephrin-B1 and EphB3 are expressed throughout the bone whereas EphB2 expression appears to be restricted ventrally. Interestingly, ephrin-B1 is also strongly expressed in the meningeal layer which derives from neural crest cells [18]. At E12.5, a stage that corresponds to the early stages of calvarial bone differentiation, ephrin-B1 is expressed throughout the head mesenchyme as well as in the vibrissae buds in wild-type embryos (Figure 2Ba and Figure S1). In ephrin-B1+/− embryos, expression of ephrin-B1 is patchy throughout the craniofacial mesenchyme and in the telencephalon, and highlights the defective formation of vibrissae buds (Figure 2Bb). Both ephrin-B2 and EphB2 exhibit expression patterns that are similar to ephrin-B1 (Figure 2Bc and 2Be). Expression of ephrin-B2, however, is unchanged in ephrin-B1+/− (Figure 2Bd) whereas EphB2 expression appears patchy (Figure 2Bf). It has been reported previously that patchy expression of ephrin-B1 in ephrin-B1+/− limb buds reflects sorting between ephrin-B1–positive and –negative cell populations that are generated in the ephrin-B1 heterozygous females via random X-inactivation and that this abnormal expression of ephrin-B1 in the limb bud correlates with a polydactyly phenotype that is observed in ephrin-B1+/− females [7,8]. Our data demonstrate that the calvarial phenotypes observed in ephrin-B1 heterozygous females correlate with an abnormal expression of ephrin-B1 and EphB2 in the presumptive frontal bone, likely due to cell sorting between ephrin-B1–positive and ephrin-B1–negative cells in the craniofacial mesenchyme.
The Frontal Bone Defect Is Caused by Abnormal Osteogenic Differentiation
To uncover the nature of the dominant effect of mosaic loss of ephrin-B1, we reasoned that understanding why sorting-out between ephrin-B1–positive and ephrin-B1–negative cells has such consequences for the development of this tissue would shed light on the dominant function of ephrin-B1. NCCs are the source of the frontal bone osteoprogenitor population, and both ephrin-B1 and ephrin-B2 control migration of these cells, raising the possibility that improper migration of NCC progenitors could be responsible for the frontal bone phenotype. Using a combination of Wnt1Cre/R26R alleles to specifically label NCCs, we found no difference in the size of the progenitor pool between ephrin-B1 heterozygous females and wild-type animals (Figure 3A), indicating that defective migration is not the likely cause for the frontal bone phenotype. In addition, no proliferation or cell survival defects were detected on sections of mutant frontal bones (unpublished data).
To test whether the defects in calvarial bone development in ephrin-B1+/− embryos correlated with perturbation of osteoblastic differentiation, we used alkaline phosphatase (AP) activity as a marker of early osteoblastic differentiation. At E16.5, AP staining of frontal bones showed delayed differentiation in ephrin-B1+/−/ephrin-B2+/GFP and ephrin-B1+/− (Figure S2B and unpublished data). At E12.5, similar levels of AP activity could be detected in wild-type, ephrin-B1Y/−, and ephrin-B1+/− embryos (Figure 3B), indicating that defective bone growth was not due to delayed onset of differentiation in heterozygous mutants. However, unlike wild-type and ephrin-B1Y/− embryos which showed continuous AP activity, AP staining of the presumptive frontal bone appeared irregular in ephrin-B1 heterozygous embryos (Figure 3Bc). These observations indicate that the calvarial defects observed in ephrin-B1+/− mutants are not due to abnormal migration or survival of NCCs, but instead might be due to a defective differentiation of the presumptive osteogenic mesenchyme.
To confirm the differentiation defects observed in ephrin-B1 heterozygotes, we isolated presumptive osteogenic mesenchymal cells from E14.5 wild-type and ephrin-B1+/− embryos, and evaluated their ability to differentiate in vitro. Using AP activity as a marker for osteogenic differentiation, we observed that cells isolated from ephrin-B1+/− embryos were consistently less prone to differentiate in vitro (Figure 3C), indicating that these defects are autonomous to the osteoprogenitor cells. In these primary cultures, expression of ephrin-B1 was detected in a punctate pattern in both AP-positive as well as AP-negative cells (Figure S3A), indicating that expression of ephrin-B1 and AP do not strictly correlate, and suggesting that ephrin-B1 does not regulate AP activity directly.
Defective Osteogenic Differentiation in ephrin-B1+/− Embryos Correlates with Abnormal Cx43 Distribution
Using this in vitro system, we tested a number of markers that have been shown to regulate osteogenic differentiation, including N-cadherin and Cx43 expression, as well as activation of MAPK. Among these, only Cx43 distribution showed prevalent changes between cultures from wild-type and ephrin-B1 heterozygous embryos (Figure 4A and unpublished data). Cx43 is a structural protein that forms gap junctional pores (connexons). Whereas cytoplasmic Cx43 shows a diffuse staining by immunofluorescence, cell surface connexons appear as bright dots because they aggregate to form functional gap junctional plaques at cell–cell interfaces. In cultures of mesenchymal cells isolated from wild-type embryos, the differentiating cells expressed a high level of Cx43, and gap junctional plaques were readily visible between these cells (Figure 4Aa and 4Ab). On the contrary, in cultures of mesenchymal cells isolated from ephrin-B1 heterozygous embryos, the distribution of Cx43 was altered and gap junctional plaques were not detected (Figure 4Ac and 4Ad). Western blot analysis indicated that the overall level of Cx43 was unchanged in primary cultures isolated from ephrin-B1 heterozygous embryos, indicating that the distribution but not the expression of Cx43 was altered in primary cultures from ephrin-B1+/− embryos (Figure S3B).
We next tested whether distribution of Cx43 was also altered in ephrin-B1 mosaic embryos. For this purpose, we co-stained paraffin sections from control and heterozygous mutant E12.5 embryos with AP and Cx43. However, even though AP staining of the frontal bone was very irregular in the ephrin-B1+/− embryo, compared to the control frontal bone (Figure S3Ca and S3Cc), we were unable to detect significant differences in Cx43 staining on these sections (Figure S3Cb and S3Cd). Since ephrin-B1 null embryos do not show calvarial phenotype or abnormal osteogenic differentiation, we reasoned that ephrin-B1 itself might not be required for proper localization of Cx43, but rather that abnormal distribution of Cx43 might be seen only at the boundary between ephrin-B1–positive and –negative cells in the mosaic embryos. To be able to detect boundaries between ephrin-B1–positive and ephrin-B1–negative cells, we generated chimeric embryos by injecting ephrin-B1 null embryonic stem (ES) cells in wild-type ROSA26 blastocysts which express β-galactosidase constitutively. Paraffin sections of X-gal–stained E11.5 chimeric embryos were processed for immunofluorescence using the Cx43 antibody. Gap junctional Cx43 was readily detected between wild-type cells and between ephrin-B1 null cells (Figure 4B). However, gap junctional Cx43 was almost never observed between ephrin-B1–positive and –negative cells. We concluded that the number of junctional pores is diminished at ephrin-B1–positive/ephrin-B1–negative boundaries in vivo.
Ephrin-B1 Associates with Cx43 and Regulates GJC
Compagni et al. have shown that the levels of EphB2 receptor are up-regulated in ephrin-B1–negative domains in ephrin-B1+/− embryos, thus creating ectopic Eph/ephrin boundaries [7]. We therefore reasoned that decreased junctional Cx43 at ephrin-B1–positive/ephrin-B1–negative boundaries in vivo could in fact indicate an inhibition of GJC by Eph/ephrin signaling. To test whether Eph/ephrin signaling could regulate GJC, we used calcein-AM as a marker of GJC in vitro. We found that interaction between ephrin-B1 and Eph-B2 resulted in inhibition of GJC in vitro (Figure 5). NIH 3T3 cells expressing ephrin-B1 showed reduced transfer of calcein-AM when plated on cells expressing low levels of Eph-B2 (Figure 5Ab), as compared to control cells (Figure 5Aa). More dramatically, plating of ephrin-B1–expressing cells over cells expressing high levels of Eph-B2 resulted in a complete inhibition of GJC (Figure 5Ac). Similar results were obtained using primary NCCs that express both ephrin-B1 and Eph-B2 at low levels. Although primary NCCs were able to establish strong GJC with control fibroblasts (Figure 5Ba and 5Bd), plating onto fibroblasts expressing ephrin-B1 markedly decreased dye transfer, even though the majority of the cells were able to spread normally (Figure 5Bb and 5Be). The fact that some cells spread normally, but were unable to transfer the dye (Figure 5Bf), indicates that inhibition of GJC is not due to cell repulsion. Plating of primary NCCs onto fibroblasts expressing Eph-B2 resulted in a moderate reduction in GJC (Figure 5Bc). We quantified GJC by two means: first, we evaluated the number of donor cells that had spread and were able to transfer the dye (Figure 5Ca). Second, we counted the number of receiving cells for each donor cell that had spread and transferred the dye (Figure 5Cb). Both measurements indicate that GJC is diminished when primary NCCs are plated on ephrin-B1– (and to a lower extent EphB2–) expressing cells. These results demonstrate that interaction between EphB2 and ephrin-B1 impairs establishment of GJC.
To better understand the molecular mechanisms by which Eph/ephrins might impinge on Cx43 distribution and regulate GJC, we analyzed the distribution of both ephrin-B1 and Cx43 in cell culture. Co-immunofluorescence studies showed that Cx43 and ephrin-B1 partially co-localize in primary mesenchymal cells (Figure 6A). We then analyzed the effect of Eph/ephrin engagement on Cx43 distribution in cell lines in which ephrin-B1 was transiently transfected. Expression of ephrin-B1 and Cx43 was partially overlapping in untreated NIH 3T3 cells, especially at interfaces between cells expressing ephrin-B1, which exhibited strong Cx43 staining (Figure 6Ba–c). Following engagement by EphB2-Fc (a soluble form of EphB2 receptor), ephrin-B1 was found in clusters that partially overlapped with the Cx43 punctate staining (Figure 6Bd–f). Similar results were obtained using MDCK cells. These results demonstrate that ephrin-B1 and Cx43 partially co-localize at the subcellular level both in absence and presence of Eph/ephrin interaction. To test whether ephrin-B1 and Cx43 physically interact, we performed a pull-down assay in NIH 3T3 cells expressing ephrin-B1. We used a recombinant protein consisting of the extracellular domain of Eph-B2 receptor fused to the Fc fragment of human IgG (EphB2-Fc) to pull down ephrin-B1. Cx43 was detected in the pull down, indicating that it interacts with ephrin-B1 (Figure 6Ca). In a converse experiment, ephrin-B1 was co-immunoprecipitated with an anti-Cx43 antibody (Figure 6Cb). These results indicate that ephrin-B1 and Cx43 interact with each other.
Regulation of GJC Underlies ephrin-Induced Cell Sorting
To identify the domain of ephrin-B1 required for the interaction with Cx43, we performed a pull down using a recombinant protein consisting of the extracellular domain of ephrin-B1 fused to the Fc fragment of human IgG (ephrinB1-Fc). Cx43 was not detected in the pull down indicating that the intracellular domain of ephrin-B1 is required for the interaction with Cx43 (Figure 6Cb). We next asked whether the PDZ-binding domain of ephrin-B1 was necessary for the interaction with Cx43, since Cx43 has been shown to interact with PDZ-containing molecules, and the data presented above suggested that Cx43 interacts with the intracellular domain of ephrin-B1. We generated a mutant form of ephrin-B1 that shows reduced binding to PDZ-containing proteins (ephrin-B1ΔPDZ [8]). The ability of this mutant form of ephrin-B1 to interact with Cx43 was tested using NIH 3T3 cells that were transiently transfected with either wild-type ephrin-B1 or ephrin-B1ΔPDZ. Western-blot analysis of whole cell lysates indicated that both proteins were expressed, albeit at different levels, and that expression of ephrin-B1 did not influence the phosphorylation status of Cx43 (detected by differences in mobility on a SDS-PAGE), which is known to regulate GJC (Figure 7A). Cx43 could be detected in the pull downs from cells expressing either ephrin-B1 wild type or ephrin-B1ΔPDZ, however, the relative abundance of phosphorylated versus unphosphorylated band was changed. More phosphorylated Cx43 (slower mobility) was observed in the ephrin-B1 wild-type pull down, whereas more unphosphorylated Cx43 was detected in the ephrin-B1ΔPDZ pull down (Figure 7A). These results indicate that the PDZ binding domain of ephrin-B1 is not required for its interaction with Cx43; however, ephrin-B1ΔPDZ and wild-type ephrin-B1 interact preferentially with different forms of Cx43.
Because phosphorylated Cx43 is thought to represent junctional Cx43 whereas unphosphorylated Cx43 represents the inactive, cytoplasmic pool, we tested whether ephrin-B1ΔPDZ co-localized with Cx43 following engagement by EphB2-Fc. Although ephrin-B1ΔPDZ was present at the cell surface and localized in clusters, we did not observe co-localization of ephrin-B1ΔPDZ and Cx43 following engagement by EphB2-Fc (Figure 7B). To test the effect of this mutation on calvarial development, we generated ephrin-B1ΔPDZ chimeric embryos. Examination of skeletal preparations of E18.5 ephrin-B1ΔPDZ chimeras revealed that these embryos did not exhibit defects in the calvarial bones, even in chimeras exhibiting a high degree of contribution of mutant ES cells (Figure S4), nor did they present polydactyly (unpublished data). However, subtle but consistent defects in sternum development, a phenotype that is associated with complete loss of ephrin-B1 [7,8], were observed in almost all of the chimeras (Figure S4), indicating that the lack of calvarial and polydactyly phenotypes in the chimeric embryos is not due to an ineffective contribution of ephrin-B1ΔPDZ ES cells to bone. To test the effect of the ephrin-B1ΔPDZ mutation on the distribution of Cx43 in vivo, we generated chimeric embryos by injecting mutant ES cells carrying the ephrin-B1ΔPDZ allele (or ephrin-B1 null ES cells as a control) in wild-type ROSA 26 blastocysts. Unexpectedly, paraffin sections of X-gal–stained E11.5 chimeric embryos revealed that unlike ephrin-B1 null cells, cells expressing ephrin-B1ΔPDZ do not sort-out from wild-type cells (Figure 7C). These results indicate that mosaic loss of reverse signaling through the PDZ binding domain is not sufficient to drive cell sorting and does not lead to defective calvarial bone development.
The inability of ephrin-B1ΔPDZ to co-localize with Cx43 upon engagement and to drive cell sorting suggested that regulation of GJC itself may play a role in the sorting-out process between ephrin-B1–positive and ephrin-B1–negative cells. To test this hypothesis, we transiently transfected HEK293T cells (that have very low levels of endogenous Cx43) with Cx43 and allowed them to sort out following trypsinization. Unlike control transfected cells, Cx43-overexpressing cells segregated from untransfected cells and were consistently found in clusters (Figure 7Da), indicating that the establishment of GJC does indeed promote cell sorting.
Overexpression of Cx43 Partially Rescues the Calvarial Phenotype
Because the lack of cell sorting in the experiments described above precluded the establishment of a functional link between the calvarial phenotype and the regulation of GJC in ephrin-B1 heterozygote embryos, we performed a genetic rescue experiment. Mice carrying a CMV-Cx43 transgene that allows for the generalized overexpression of Cx43 [21] were bred to ephrin-B1−/− mice. Western-blot analysis showed a modest increase in the level of Cx43 in presumptive frontal bones of embryos carrying the transgene (unpublished data). Skeleton preparations of P1 offspring indicated that all of the skeletal defects previously observed in the ephrin-B1 mutants, including the phenotype specific to ephrin-B1 heterozygotes, were also found in this mixed genetic background (unpublished data). To assess whether overexpression of Cx43 had an effect on the calvarial phenotype observed in ephrin-B1+/− mutants, we quantified the foramen area between the frontal bones in newborn pups from all genotypes (see the Materials and Methods section). Although overexpression of Cx43 had no significant impact on the development of the frontal bones in male pups, the foramen area was decreased in the ephrin-B1+/− pups carrying the CMV-Cx43 transgene compared to ephrin-B1+/− pups without the transgene (Figure 8). Whereas the difference in foramen area between the ephrin-B1+/− pups with and without the CMV-Cx43 transgene was significant, the degree of rescue of frontal bone development in presence of the transgene was variable from sample to sample (unpublished data). Importantly, overexpression of Cx43 did not change the skeletal defects that are independent of cell sorting (i.e., sternum defects) (unpublished data). These results establish a functional link between ephrin-B1 and GJC, and suggest that improper regulation of GJC is implicated in the calvarial defects observed in ephrin-B1 heterozygous females.
Discussion
ephrin-B1+/− Mice as a Model for CFNS
In this study we have shown that, analogous to humans, heterozygous loss of ephrin-B1 in mice results in defective development of the skull vault. Human CFNS patients carrying mutations in the ephrin-B1 gene exhibit a range of craniofacial defects including ocular hypertelorism, malformation of the face (in particular the forehead and the nose), cranium bifidum occultum, and craniosynostosis [22]. Our study shows that at birth, ephrin-B1+/− mice exhibit a delay in the ossification of calvarial bones leading to a frontal foramen similar to cranium bifidum occultum, and in many instances, to a bifid nose, mimicking the human disease. Removing one copy of ephrin-B2 resulted in increased severity of the frontal foramen, and an additional coronal suture defect in an ephrin-B1+/− background, but had no effect in ephrin-B1 null embryos, suggesting that the lack of phenotype in ephrin-B1 hemizygous males is not due to functional compensation by ephrin-B2. The effect of removing both copies of ephrin-B2 on calvarial development could not be analyzed due to the early embryonic lethality of ephrin-B2 null embryos.
Interestingly, our mutant mice did not exhibit craniosynostosis of the coronal sutures seen in humans. The processes leading to calvarial foramina and craniosynostosis are genetically linked, as evidenced by loss of function and gain of function mutations in the transcription factor Msx2, respectively. Moreover, heterozygosity for the transcription factor Twist can lead to craniosynostosis, as well as to foramina, and both transcription factors regulate differentiation of frontal bone osteoprogenitors (Ishii et al., 2003). Ectopic bone growth observed within the coronal suture of ephrin-B1+/−/ephrin-B2+/GFP embryos might be prescient of a premature fusion of the suture, but this could not be analyzed at later stages, since 100% of these animals die at birth (A. Davy, P. Soriano, unpublished data). The genetic interaction suggests, however, that the discrepancy in coronal suture phenotype between ephrin-B1 heterozygous mice and humans could be due to genetic modifiers. Our expression analysis does not support a model in which wild-type expression of ephrin-B1 controls suture formation by establishing boundaries in craniofacial mesenchyme. In fact, the observation that ephrin-B1 null animals have no defects in calvarial bones argues that the function of ephrin-B1 is not normally required for either calvarial bone development or proper suture formation. On the other hand, the fact that ephrin-B1 is expressed in the meningeal layer could account for both the suture as well as the low-penetrance parietal bone phenotype observed in ephrin-B1+/− embryos because this layer is involved in suture and parietal bone formation [18,23].
Most of the mutations in ephrin-B1 that have been identified in CFNS patients are located in the 5′ end of the gene and are consistent with loss-of-function mutations, either by introducing a premature stop codon, or by presumably interfering with the binding of ephrin-B1 to Eph receptors [9,10]. However, it was reported more recently that some patients harbored mutations in the 3′ end of the gene, which might affect specifically the reverse signaling activity of ephrin-B1 [24]. Our data using chimeric embryos demonstrate that a mutation in the PDZ binding domain of ephrin-B1, which is known to phenocopy some of the phenotypes observed in ephrin-B1 null embryos including cleft palate [8], does not induce calvarial defects or polydactyly. These results indicate that the mutations found in CFNS patients might impinge on protein stability, protein localization, or binding of effector molecules independent of the PDZ domain.
Defective Osteogenic Differentiation and Inhibition of GJC
Our results are consistent with a model in which inhibition of GJC at ectopic Eph/ephrin boundaries in ephrin-B1+/− females results in an abnormal differentiation of osteoprogenitors, thereby leading to the defective development of frontal bones and also, presumably, to polydactyly. Consistent with a previous report [6], we found that Eph/ephrin interaction inhibits GJC. In addition, junctional Cx43 was decreased at ectopic Eph/ephrin boundaries in vivo, and reduced junctional Cx43 correlated with decreased osteogenic differentiation of primary cells isolated from ephrin-B1 heterozygous embryos. Finally, we found that overexpression of Cx43 partially rescued the calvarial phenotype observed in ephrin-B1 heterozygote females. These results are consistent with other reports documenting the role of GJC and Cx43 in osteogenic differentiation [25–27]. In addition, several genetic studies have shown that defective regulation of GJC affects craniofacial and digit development, indicating that the structures affected in ephrin-B1 heterozygous females are highly sensitive to alteration in GJC. Mice deficient for Cx43 exhibit delayed ossification of the calvarial bones and craniofacial abnormalities [28], but more importantly, mutations in GJA1 (the gene coding for Cx43) in humans are responsible for oculodentodigital dysplasia (ODDD), a syndrome that is characterized by defective craniofacial development and digit formation [16]. An ENU screen in mice recently uncovered a dominant mouse mutation that exhibits many of the classic features of ODDD, and positional cloning revealed that these mice carry a mutation in GJA1 that acts in a dominant-negative fashion to disrupt gap junction assembly and function [17]. On the basis of our results, we can not rule out the possibility that Eph/ephrin signaling might also impinge on osteogenic differentiation directly by activating signal transduction cascades (that would not involve the PDZ domain of ephrin-B1). However, the cytoplasmic kinases that are known targets of Eph/ephrin signaling (Src family kinases and MAPKs) have been shown to be positive regulators of osteogenic differentiation, which is not consistent with the inhibition of differentiation that we observe in vivo and in vitro. Moreover, we have not been able to detect a change in the level of MAPK activation in ephrin-B1+/− cultures (unpublished data).
We found that ephrin-B1 and Cx43 form a complex and that following engagement by Eph receptors, ephrin-B1 and Cx43 co-localize in clusters that could be indicative of endocytosis. Recent reports have shown that Eph/ephrins complexes are internalized following interaction [29–31], and endocytosis is also a well-known mode of regulation of connexons at the cell surface [32]. In both cases, pieces of the plasma membrane from neighboring cells are internalized. It is therefore conceivable that Cx43 might be co-internalized with Eph/ephrin complexes. However, it is also possible that endocytosis of connexons is regulated via a signal transduction cascade, since it has been shown recently that Eph/ephrin signaling regulates clathrin-mediated endocytosis by tyrosine phosphorylation of Synaptojanin 1 [33]. Tyrosine phosphorylation of Cx43 is also a mechanism by which GJC is regulated. In our pull-down assay, wild-type ephrin-B1 interacted preferentially with phosphorylated Cx43 whereas ephrin-B1ΔPDZ interacted preferentially with unphosphorylated Cx43, suggesting that the interaction between ephrin-B1 and Cx43 might not be direct, and that these proteins might interact differently when at the cell surface or in the cytoplasm.
Regulation of GJC and Cell Sorting
We observed that junctional Cx43 was enriched at interfaces between ephrin-B1–positive cells, suggesting that while GJC is inhibited at Eph/ephrin interfaces, it might be promoted at ephrin/ephrin interfaces. This observation, as well as the fact that overexpression of Cx43 leads to cell sorting, supports the idea that regulation of GJC contributes to Eph/ephrin-induced cell sorting. Although GJC has not been formally linked to cell sorting previously, there is ample evidence in the literature that regulation of GJC influences cell–cell contacts. Indeed, it has been shown in various experimental settings that inhibiting GJC, either through the use of blocking antibodies or dominant negative constructs, induces loss of adhesion [34–37]. In Xenopus, both overexpression of ephrin-B1 and dominant-negative connexin result in de-adhesion of the blastomeres [36,38]. Sorting of cells overexpressing Cx43 has also been reported previously in PC12 cells [39]. We therefore propose that regulation of GJC contributes to cell sorting downstream of Eph/ephrin interaction (Figure 9A).
A question that remains unanswered is whether or not the cell sorting observed in ephrin-B1 heterozygotes is fully dependent on Eph receptors. Interestingly, an Eph-independent role for ephrin-B1 in regulating tight junctions has recently been reported [40], and it was recently shown that EphA4 can induce cell sorting independently of ephrins [41,42]. However, the fact that some CFNS patients harbor point mutations in the extracellular domain of ephrin-B1 that presumably have an effect on Eph/ephrin interaction argues for the involvement of Eph receptors in the sorting process.
On the basis of our data, we propose a model that explains the dominant effect of mosaic loss of ephrin-B1 in ephrin-B1 heterozygous females, and that revisits the function of Eph/ephrin in embryo patterning (Figure 9B). In wild-type mice and within a developmental compartment, ephrin-expressing cells establish GJC which stabilizes cell–cell interactions and creates a communication compartment. In ephrin-B1 heterozygotes as well as at a developmental boundary, GJC is inhibited between ephrin-positive and Eph-positive cells, possibly via endocytosis of Cx43, which prevents stable cell–cell interactions and leads to the formation of distinct compartments. Inhibition of GJC at developmental boundaries has been shown in a variety of models, including inter-rhombomeric boundaries [12,43], which also happen to be Eph/ephrin boundaries. In addition, overexpression of Cx43 has recently been shown to rescue a central nervous system boundary defect in mice [44].
In conclusion, our work demonstrates that in addition to their prominent role in regulating the actin cytoskeleton, Eph receptors and ephrins also play an important role in regulating gap junctional communication and suggests that improper regulation of GJC leads to the phenotypes observed in ephrin-B1 heterozygous individuals. Together, these functions make Eph receptors and ephrins potent regulators of boundary formation and tissue patterning during embryonic development.
Materials and Methods
Mice.
Ephrin-B1 mutant mice and CMV-Cx43 transgenic mice have been described elsewhere [8,22]. The ephrin-B2GFP allele was generated by inserting the H2BGFP cDNA cassette at the MluI site in the first exon of ephrin-B2. The MluI-XbaI fragment encompassing the start codon of the ephrin-B2 gene was replaced with H2BGFP. The mutation was introduced in ES cells by homologous recombination. Mice were maintained in a 129S4/C57Bl6J mixed background. Genotyping was done by PCR using the following sets of primers: GFP-F: 5′-GCAAGAAGGCGGTGACTAAGGCGC-3′; GFP-R: 5′-GGCCGCCGCCAGTGCTTGAGGTCG-3′. Mice were housed in microisolator racks in a facility accredited by the Association for the Assessment and Accreditation of Laboratory Animal Care, and experimentation was reviewed by the Hutchinson Center Institutional Review Committee. The ES cells carrying the ephrin-B1ΔPDZ mutation have been described previously [8]. The sequence of the primer specific to the mutant allele was: 5′-GCCATGCTGGGCCTTCACT-3′. To obtain ephrin-B1 null ES cells, one clone of targeted ES cells carrying the conditional allele of ephrin-B1 (ephrin-B1lox) was electroporated with a PGK-Cre expression vector. ES cell clones were isolated and screened by Southern-blot for the recombined ephrin-B1 locus. Targeted ES cells were injected into blastocysts obtained from crosses between F1 (129S4/C57Bl6J) ROSA26-βgal homozygous males and either wild-type C57BL6J/CBAJ or MF1 females.
X-gal staining and skeletal preparations.
Procedures used for X-gal staining and skeletal preparations have been described in detail elsewhere [8]. For the experiment with mice carrying the CMV-Cx43 transgene, quantification of the foramen area between frontal bones was performed using Photoshop (Adobe, San Jose, California, United States) to record the number of pixels corresponding to the foramen. The mean values for females with (97,901 pixels) and without (117,391 pixels) the transgene were then normalized to the mean value obtained for males (91,824 pixels) (no significant difference was found between males with or without the transgene, n = 10). Statistical significance was calculated using an unpaired t-test with Welch correction.
In situ hybridization.
Section in situ hybridization experiments were performed on frontal sections of E14.5 embryos as described previously [45]. Whole-mount in situ hybridization was performed according to a protocol described elsewhere [46]. Probe sequences used for ephrin-B1, EphB2, and EphB3 are available upon request.
Primary cultures.
Presumptive frontal bone mesenchyme was dissected from E14.5 embryos and incubated with 1% trypsin/0.5% DNAse in PBS for 10 min at 37 °C. Trypsin was inactivated by addition of complete medium (DMEM containing 15% FCS) and cells were dissociated by trituration with a glass Pasteur pipette. Cells were pelleted by centrifugation and washed twice in complete medium. Cells were plated at high density in a 24-well plate and kept in culture in complete medium. After 3 d, cells were fixed in 2% PFA and rinsed three times in NTMT. AP activity was detected by incubating fixed cells with NBT/BCIP (Roche, Basel, Switzerland). Some cultures were further processed for immunofluorescence as described below. Quantification of AP activity was performed on digital images using the Image J software (http://rsb.info.nih.gov/ij). The data presented are representative of five independent experiments.
Primary NCCs were isolated from dissected branchial arches of E9.5 embryos as described above, except cells were cultured in F12 medium supplemented with 10% FCS.
GJC assays.
NIH 3T3 cells were stably transfected with expression vectors for ephrin-B1, Eph-B2 receptor, or the pcDNA3 control vector. Recipient cells were plated at high density. Donor cells (NIH 3T3 expressing ephrin-B1 or primary NCCs) were incubated with calcein-AM (Molecular Probes, Eugene, Oregon, United States) for 20 min at 37 °C. Calcein-AM loaded cells (donors) were extensively washed in PBS, trypsinized, and dropped onto confluent monolayers of NIH 3T3 cells transfected with either pcDNA3 or expressing ephrin-B1 or various levels of Eph-B2. Transfer of calcein-AM through gap junctions was assessed after 3 h incubation at 37 °C. GJC establishment was quantified by two means for each conditions: (1) the percentage of cells that were spread and had transferred the dye versus cells that were spread but did not transfer the dye; and (2) the number of cells receiving the dye for each donor. These data were acquired by visual assessment either directly on the inverted microscope or from digital images. Statistical significance was calculated using a Student t-test.
Western blot analysis.
Cells were scraped in 1% NP40 lysis buffer (50 mM Hepes [pH 7.5], 150 mM NaCl, 10% glycerol, 1.5 mM MgCl2, 1mM EGTA, 100 mM NaF), except for the experiment presented in Figure 7A (100 mM Tris [pH 7.4], 150 mM NaCl, 1mM EDTA). Protein lysates were incubated with either 5-μg EphB2-Fc or 4-μg Cx43 monoclonal antibody (generous gift from P. Lampe) for 4 h at 4 °C and subsequently with 20-μl ProteinA-Sepharose. Affinity complexes were analyzed by SDS-PAGE using the following antibodies: ephrin-B1 (A20 or C18, Santa Cruz Biotechnology, Santa Cruz, California, United States), Cx43 (Sigma, St. Louis, Missouri, United States). We have noted that the binding of Cx43 to ephrin-B1 is sensitive to the lysis buffer used: the interaction is lost in RIPA buffer.
Immunofluorescence.
NIH 3T3 cells were plated on glass coverslips and transiently transfected with an expression vector for ephrin-B1. Forty hours after transfection, cells were either fixed in 2% PFA or incubated with 4-μg/ml EphB2-Fc for 30 min at 37 °C and then fixed in PFA. Cells were permeabilized with 0.1% Tx-100 for 3 min, rinsed in PBS, and incubated with a mix of EphB2-Fc (4 μg/ml) and either Cx43 monoclonal antibody or N-cadherin monoclonal antibody (Zymed, Carlsbad, California, United States). In primary cells, ephrin-B1 was detected using the 25H11 rat monoclonal antibody [47]. FITC- and Cy3-conjugated secondary antibodies were from Jackson ImmunoResearch (West Grove, Pennsylvania, United States). For the sorting experiments, HEK293T cells were transiently transfected with either DsRed or a Cx43 expression construct. Twenty-four hours after transfection, cells were trypsinized into a single-cell suspension and replated onto glass coverslips. Immunofluorescence was performed at 48 h post-transfection using a Cx43 antibody (Sigma).
For immunofluorescence on sections, paraffin sections were rehydrated and subjected to a citrate boil to reveal antigens. Tissue sections were blocked in Blocking solution (PBS/5% horse serum) 1 h at room temperature and incubated overnight at 4 °C in Cx43 antibody (1/100 in Blocking solution [Sigma]). Incubation with the Cy3-conjugated secondary antibody was for 1 h at room temperature (1/250 in Blocking solution). The number of Cx43 dots was counted on digital images acquired on a Delta Vision Deconvolution microscope (Applied Precision Inc., Issaquah, Washington, United States). The number of Cx43 dots was normalized to the number of junctions: for each section, we evaluated the number of wild-type/knock-out (KO) junctions (15–20) and counted a similar number of wild-type/wild-type and KO/KO junctions on each side of the wild-type/KO boundary.
Supporting Information
Figure S1
Patterning of the Vibrissae Buds Is Altered in ephrin-B1+/− Embryos
(A) Whole E14.5 embryos were stained with H&E to highlight whiskers buds of ephrin-B1Y/− hemizygous males (a) and ephrin-B1+/− heterozygous females (b). X-gal staining of E14.5 ephrin-B1+/lox embryo carrying the R26R and Wnt1-Cre alleles (c). The patterning of vibrissae buds is abnormal in heterozygote but not homozygote mutants. This defect is autonomous to the NCC lineage since eliminating ephrin-B1 specifically in this lineage is sufficient to phenocopy the vibrissae bud defect (c).
(B) In situ hybridization on frontal sections of E14.5 wild-type embryo using probes for ephrin-B1 (a) and EphB2 (b) showing expression of these genes in the mesenchyme around vibrissae buds (arrowheads).
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Figure S2
Calvarial Defects and Decreased Osteogenic Differentiation
(A) Skeleton preparations of whole heads from E17.5 littermate embryos, ephrin-B2+/GFP single heterozygous mutants (a), ephrin-B1+/− single heterozygous females (b), and ephrin-B1+/−/ephrin-B2+/GFP double heterozygous females (c). Ectopic bone can be seen in the suture mesenchyme of ephrin-B1/ephrin-B2 double heterozygous females (arrowhead in c) and the foramen between the frontal bones is larger than in ephrin-B1+/− embryos. An outline of the bones is presented for better visualization (d–f).
(B) AP staining on cryosections of E15.5 embryos. A wild-type (a), an ephrin-B1 null male (b), and an ephrin-B1+/−/ephrin-B2+/GFP double heterozygous female (c) are shown. Bone front in the control embryo (a) and (b) is closer to the midline than bone front in the heterozygous littermate (c). In addition, the thickness of the bone is decreased in the ephrin-B1+/−/ephrin-B2+/GFP double heterozygous female. Both observations show that differentiation of the frontal bone is decreased in the heterozygote.
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Figure S3
Ephrin-B1 and Cx43 Distribution in Primary Cultures and Differentiating Frontal Bones of ephrin-B1+/− Embryos
(A) Ephrin-B1 was detected by immunofluorescence in cultures of primary mesenchymal cells (a) and (c) that were also stained for AP activity (b) and (d). Expression of ephrin-B1 was readily detected as a punctate staining on these primary cells, both in AP-positive (a) and (b) and in AP-negative cells (c) and (d).
(B) Western blot analysis of Cx43 levels in primary cultures of mesenchymal cells isolated from littermates of various genotypes. No difference in the overall Cx43 levels was detected.
(C) Paraffin sections from E12.5 control embryos (ephrin-B1Y/−) (a) and (b) and ephrin-B1+/− embryos (c) and (d) were stained for AP activity (a) and (c) and subsequently processed for Cx43 immunostaining (b) and (d). AP staining in the developing frontal bone of ephrin-B1+/− embryo is more irregular than in the control embryo, consistent with the results presented in Figure 3. However, no significant difference can be seen in the overall staining for Cx43 in this tissue.
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Figure S4
ephrin-B1ΔPDZ Chimeric Embryos Do Not Exhibit Calvarial Defects
(A) Skeletal preparations of whole heads from E18.5 ephrin-B1ΔPDZ chimeric embryos showing low (a) and high (b) contribution of mutant ES cells, assessed by PCR on tail DNA (c).
(B) Skeletal preparations of E18.5 wild-type (a), ephrin-B1−/− (b), and ephrin-B1ΔPDZ chimeras (c) and (d) show that chimeric embryos exhibit sternum defects reminiscent of ephrin-B1 null embryos.
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Acknowledgements
We thank Cecilia Lo for sharing the CMV-Cx43 mice; Mark Henkemeyer for providing the Eph-B2 expression construct and the in situ probe for EphB2; Paul Lampe for sharing his Cx43 antibody and the Cx43 expression construct; Wieland Huttner for the 25H11 rat monoclonal antibody [47]; Philip Corrin, Jason Frazier, and Marc Grenley for excellent technical assistance; and our laboratory colleagues for critical reading of the manuscript.
Abbreviations
AP - alkaline phosphatase
CFNS - craniofrontonasal syndrome
Cx43 - connexin43
ES - embryonic stem
GFP - green fluorescent protein
GJC - gap junction communication
KO - knock-out
NCC - neural crest cell
Figures and Tables
Figure 1
ephrin-B1+/− Embryos Exhibit Defects in NCC Derivatives
(A) Skeleton preparations of whole heads from E18.5 mutant embryos (a–c). Bones are stained with Alizarin Red while cartilage is stained with Alcian Blue. An opening (foramen) between frontal bones is observed in ephrin-B1+/− heterozygous females (b) but not in ephrin-B1−/− homozygous females (a). Mutant embryos in which ephrin-B1 is specifically deleted in NCC also exhibit defects of the frontal bones (c). Arrowheads show the bone front of the frontal bones. Schematic drawings of the frontal bones (d–f).
(B) Skeleton preparations of whole heads from E18.5 embryos (a–c) or E15.5 embryos (d–f) show that ephrin-B1+/−/ephrin-B2+/GFP embryos present an additional coronal suture defect (c) and (f) as compared to ephrin-B1+/− single heterozygous females (b) and (e). At E18.5, bone fronts never overlap at coronal sutures of ephrin-B1+/−/ephrin-B2+/GFP embryos, and ectopic bone forms in the suture mesenchyme (arrow in [c]). At E15.5, parietal and frontal bone fronts forming the coronal suture (arrow) of ephrin-B1+/−/ephrin-B2+/GFP embryos (f) are further apart than in control littermates (d) and (e). Coronal suture and frontal bones are normal in single ephrin-B2+/GFP heterozygous mutants (a) and in ephrin-B1Y/−/ephrin-B2+/GFP males (d).
fb, frontal bone; pb, parietal bone.
Figure 2
Expression of ephrin-B1 and EphB2 Is Abnormal in ephrin-B1+/− Embryos
(A) In situ hybridization on frontal sections of E14.5 wild-type embryo using probes for ephrin-B1 (a), EphB2 (b), or EphB3 (c). Ephrin-B1 is expressed throughout the developing frontal bone (marked by dotted lines) as well as in the meningeal layer (arrowheads). Expression of EphB2 is restricted ventrally (arrowheads) whereas EphB3 is expressed throughout the developing frontal bone.
(B) In situ hybridization of wild-type (a), (c), and (e) or ephrin-B1+/− embryos (b), (d), and (f) using a probe for ephrin-B1 (a) and (b), ephrin-B2 (c) and (d) and EphB2 (e) and (f). Both ephrin-B1 and EphB2 show sorting in the telencephalon and the craniofacial mesenchyme of ephrin-B1+/− embryos (arrowheads) while expression of ephrin-B2 is unaffected.
Figure 3
Calvarial Foramen Correlates with Impaired Osteogenic Differentiation
(A) E11.5 embryos carrying the R26R and Wnt1-Cre alleles were processed for X-gal staining to label NCCs. No difference in the osteogenic precursor population (arrow) was detected in ephrin-B1+/− females (b) as compared to wild type (a).
(B) E12.5 wild-type (a), ephrin-B1Y/− (b), and ephrin-B1+/− (c) embryos were stained for AP activity. The onset of osteogenic differentiation does not seem affected in the mutant embryos, but AP staining pattern is irregular in the heterozygote mutant as compared to wild-type and homozygote mutant, with areas devoid of staining (arrows).
(C) Primary mesenchymal cells were isolated from presumptive calvaria of E14.5 ephrin-B1Y/− hemizygous male (a) or ephrin-B1+/− heterozygous female (b) and plated at high density. AP activity was evaluated after 3 d in culture. Digital quantification of AP activity (c).
Figure 4
Decreased Osteogenic Differentiation Correlates with Abnormal Distribution of Cx43
(A) Primary mesenchymal cells isolated from presumptive calvaria of wild-type (a) and (b) or ephrin-B1+/− embryos (c) and (d) were stained for AP activity (a) and (c) and subsequently processed for Cx43 immunofluorescence (b) and (d). Junctional Cx43 evidenced by bright dots is readily detected in cultures from wild-type embryos (WT), but not in cultures from ephrin-B1+/− embryos.
(B) Detection of Cx43 by immunofluorescence in limb bud sections of an X-gal–stained chimeric embryo obtained by injecting ephrin-B1 null cells into a ROSA26-βgal blastocyst. Gap junctional Cx43 appears as bright dots (a). The boundary between wild-type cells (dark cells in [c]) and ephrin-B1 null cells (white cells in [c]) is indicated by red arrows. A schematic representation of the results is shown (b). Grey cells are wild type and white cells are ephrin-B1 null. Cx43 dots are in red, whereas yellow in (b) marks the only Cx43 dot that might be between a wild-type and a null cell. Quantification of Cx43 dots in multiple sections show a reduction in the number of Cx43 dots between wild-type and null cells (WT-KO) (d).
Figure 5
Eph/ephrin Interaction Inhibits Gap Junction Communication
(A) NIH 3T3 cells stably expressing ephrin-B1 were loaded with calcein-AM and dropped onto monolayers of NIH 3T3 cells either not expressing (a) or expressing variable levels (b) and (c) of Eph-B2 receptor. Transfer of dye to neighboring cells (arrowheads) was evaluated by fluorescence after 3 h. No dye transfer (arrows) was observed when high levels of Eph-B2 were expressed.
(B) Primary NCCs were loaded with Calcein-AM and dropped onto NIH 3T3 cells that were transfected either with a control plasmid (pcDNA3 [a]), or an expression construct for ephrin-B1 (b) or Eph-B2 (c). In the control situation, the majority of cells transferred the dye (arrowheads). The boxed area is shown at higher magnification in (d). Transfer of dye is visualized by faint staining of cells surrounding the very bright donor cell. Following Eph/ephrin interaction, many of the donor cells did not transfer the dye (arrows). Boxed areas in (b) are shown at higher magnification in (e) and (f). Cells transferring the dye (f) and cells not transferring the dye (e) are able to spread.
(C) The percentage of cells that had spread and showed transfer of dye was evaluated after 3 hours (a). In addition, the number of receiving cells for each donor cell was counted (b).
*p < 0.05 compared to control.
Figure 6
Ephrin-B1 and Cx43 Co-localize and Interact with Each Other
(A) Primary mesenchymal were co-stained for Cx43 ([a], red in [c]) and ephrin-B1 ([b], green in [c]).
(B) NIH 3T3 cells were transiently transfected with ephrin-B1 and co-immunofluorescence studies were performed to detect Cx43 (a) and (d) and ephrin-B1 (b) and (e) either without (a–c) or with engagement of ephrin-B1 (d–f). In untreated cells, Cx43 is enriched at interfaces between ephrin expressing cells and co-localizes with ephrin-B1 (c). Engagement by EphB2-Fc results in clustering of ephrin-B1 (e) and Cx43 partially co-localizes with ephrin-B1 clusters (f). Insets present higher magnification views.
(C) Ephrin-B1 was affinity precipitated from NIH 3T3 cells using EphB2-Fc. The presence of Cx43 in the affinity complex was assessed by Western-blot (a). Protein lysates from NIH 3T3 cells expressing ephrin-B1 were incubated with ephrinB1-Fc, and the presence of Cx43 in the affinity complexes was assessed by Western blot (b) Cx43 was immunoprecipitated from NIH 3T3 cells expressing ephrin-B1 and the presence of ephrin-B1 in the immunocomplexes was detected by Western blot (right panel).
IP, immunoprecipitation; Wcl, whole cell lysate.
Figure 7
Regulation of GJC Correlates with Cell Sorting
(A) NIH 3T3 cells were transiently transfected with either wild-type ephrin-B1 (B1), ephrin-B1ΔPDZ (ΔPDZ), or a control plasmid (−). Protein lysates were incubated with EphB2-Fc, and the presence of Cx43 in the affinity complex was assessed by Western blot. ProtA indicates a sample that was incubated with Protein A in absence of EphB2-Fc. Whole cell lysates (a) and affinity precipitations (b) were analyzed by Western blot.
np, non phosphorylated Cx43; p1, p2, two different forms of phosphorylated Cx43; wcl, whole cell lysate.
(B) NIH 3T3 cells were transiently transfected with ephrin-B1ΔPDZ and treated with EphB2-Fc. Subcellular localization of Cx43 (a) and ephrin-B1ΔPDZ (b) was analyzed by immunofluorescence. No co-localization was observed between these two proteins (c).
(C) Paraffin sections of limb buds from E11.5 chimeric embryos obtained from injection of either ephrin-B1 null ES cells (K/O) (a) or ephrin-B1ΔPDZ ES cells (ΔPDZ) (b). Chimeric embryos were X-gal stained, processed for histology, and paraffin sections were counter-stained with Nuclear Fast Red. The wild-type cells are blue whereas the mutant cells are red. Cells expressing ephrin-B1ΔPDZ do not sort from wild-type cells.
(D) HEK293T cells overexpressing Cx43 were detected by immunofluorescence as a tightly packed cluster of cells (a) whereas cells expressing DsRed are scattered among non-expressing cells (b). (c) and (d) show the visible image for the same field of cells.
Figure 8
Partial Rescue of the Calvarial Defect in ephrin-B1+/− Mice Overexpressing Cx43
Skeleton preparations from an ephrin-B1Y/− male (a), an ephrin-B1+/− female (b) and an ephrin-B1+/− female carrying the CMV-Cx43 transgene (c). Quantification (d) of the foramen area between the frontal bones of an ephrin-B1+/− female (B1+/−) and an ephrin-B1+/− female carrying the CMV-Cx43 transgene (CMV-Cx43;B1+/−). Mean values for the females with and without the transgene were normalized to the value obtained for males. The number of embryos for each genotype is indicated. Error bars represent standard error of the mean values.
*p = 0.0224.
Figure 9
Mosaic Loss of ephrin-B1 and Establishment of Developmental Boundaries
(A) Potential cross-talk between gap junctions and adherens junctions. Ephrin-B1 co-localizes with Cx43 in junctional plaques. Co-regulation of gap junctions and adherens junctions has been extensively studied.
(B) Establishment of communication compartments for embryo patterning. Within a compartment and in the developing calvarial bones, all cells express ephrin-B1 and are coupled via GJC, exchanging second messengers (black dots). At a developmental boundary and at ectopic ephrin boundaries in ephrin-B1+/− embryos, Eph/ephrin interaction leads to inhibition of GJC, possibly through endocytosis of Cx43, concomitant with a loss of stable cell–cell interactions between the two cell types. Sorting between these cells and inhibition of GJC concur to establish distinct developmental compartments.
Footnotes
Competing interests. The authors have declared that no competing interests exist.
Author contributions. AD and PS conceived and designed the experiments. AD and JOB performed the experiments. AD, JOB, and PS analyzed the data. AD and PS wrote the paper.
Funding. This work was supported by grants HD24875 and HD25326 from the National Institute of Child Health and Human Development to PS. AD is a Canadian Institute of Health Research postdoctoral fellow. JOB is the recipient of a F32 postdoctoral fellowship from the National Institute of Dental and Craniofacial Research (DE17506). | [
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17029558 | Mutation at the Evi1 Locus in Junbo Mice Causes Susceptibility to Otitis Media
Abstract
Otitis media (OM), inflammation of the middle ear, remains the most common cause of hearing impairment in children. It is also the most common cause of surgery in children in the developed world. There is evidence from studies of the human population and mouse models that there is a significant genetic component predisposing to OM, yet nothing is known about the underlying genetic pathways involved in humans. We identified an N-ethyl-N-nitrosourea-induced dominant mouse mutant Junbo with hearing loss due to chronic suppurative OM and otorrhea. This develops from acute OM that arises spontaneously in the postnatal period, with the age of onset and early severity dependent on the microbiological status of the mice and their air quality. We have identified the causal mutation, a missense change in the C-terminal zinc finger region of the transcription factor Evi1. This protein is expressed in middle ear basal epithelial cells, fibroblasts, and neutrophil leukocytes at postnatal day 13 and 21 when inflammatory changes are underway. The identification and characterization of the Junbo mutant elaborates a novel role for Evi1 in mammalian disease and implicates a new pathway in genetic predisposition to OM.
Synopsis
Otitis media (OM), inflammation of the middle ear, is the most common cause of deafness in children. Although acute episodes of OM in children are associated with middle ear infections, in a substantial portion of cases recurrent episodes of OM, or a chronic suppurative OM, will develop. There is evidence from genetic studies of human families that there is a significant genetic component contributing to the development of recurrent and chronic forms of OM. However, the genes involved have not been identified. The authors have identified and characterized mouse mutants that demonstrate chronic OM as a route to identifying genes involved with OM. This study describes one mutant, Junbo, which shares many features with human OM. Junbo develops an acute OM following birth that subsequently develops into a chronic suppurative form of OM. Junbo carries a mutation in the transcription factor gene, Evi1. Evi1 is expressed in a variety of cell types in the middle ear lining when inflammatory changes are underway. The identification of the Junbo mutation implicates a new gene involved in predisposition to OM.
Introduction
Otitis media (OM), inflammation of the middle ear, remains the most common cause of hearing impairment in children [1,2]. Acute episodes of OM in infants and children are most often associated with middle ear infections involving the pathogens Streptococcus pneumoniae and Haemophilus influenzae [3]. Prolonged stimulation of the inflammatory response, along with poor mucociliary clearance, can also lead to the persistence of middle ear fluid giving rise to the clinical presentation of otitis media with effusion [2]. In a substantial portion of children, recurrent episodes of OM or a chronic suppurative OM will develop. The high prevalence of the disease, coupled with its recurrent and chronic nature, accounts for the large number of tympanostomies undertaken in affected children. OM is still the most common cause of surgery in children in the developed world. There is still considerable debate over the etiology of OM and the underlying pathological mechanisms [2]. However, risk factors for OM include craniofacial abnormalities, impaired mucocilliary function, and the presence of an inflammatory stimulus, such as bacteria. There is evidence from studies of the human population and mouse models that there is a significant genetic component predisposing to recurrent or chronic OM [4–7], yet little is known about the underlying genetic pathways involved. While several inbred strains are predisposed to the development of OM, their genetic analysis and utility is compounded by the complex genetic bases and the low penetrance of the phenotype [6]. In addition, there are several mouse mutants that demonstrate an OM phenotype, but the OM develops as part of a complex syndrome with a wide spectrum of phenotypes [6]. It will be important to identify and characterize the genes underlying highly penetrant mouse mutants that develop OM in the absence of other diverse pathology and represent appropriate models for OM in the human population.
Large-scale phenotype-driven mouse ENU (N-ethyl-N-nitrosourea) mutagenesis programs provide a rich source of novel mutant phenotypes that are the basis for systematic efforts to identify the genetic basis for diverse disease states [8–10]. One such screen at MRC Harwell, recovered a large number of mutant phenotypes representing ENU-induced mutations at a number of novel loci in the mammalian genome [9]. Mouse models have and continue to play an important role in studying the genetic causes of hearing impairment. A number of mutations have been cloned and have provided us with several profound insights into the critical proteins involved with the development and function of the auditory apparatus at the level of both the middle and inner ears [11,12]. Nevertheless, it is clear that we do not possess mouse mutants for all loci and pathways potentially involved in hearing impairment.
We report the characterization of a new mutant, Junbo, identified in the Harwell mutagenesis program as having a deafness phenotype. Junbo is a model of OM that shares many features with the human condition. Acute OM arises spontaneously in the postnatal period and develops into chronic suppurative OM with otorrhea. The underlying molecular basis of this phenotype has been identified as a mutation in the Evi1 transcription factor causing a nonconservative Asn763Ile change in the second of the two zinc-finger domains in this protein. The Junbo mouse highlights a new role for the transcription factor Evi1 and provides the first evidence for the genetic pathways involved in the etiology of this complex childhood disease.
Results
Identification, Mapping, and Cloning of the Junbo Mutation
An ENU mutagenesis screen [9] identified a new dominant mutant, Junbo (Jbo), with hearing loss. Preliminary phenotyping using a click-box test (see Materials and Methods) of an age-matched cohort derived from the founder mouse indicated that mice demonstrated a hearing loss at ~40 d after birth (DAB). We used a pooling strategy employing DNA from affected mice and genome-wide fluorescent simple sequence length polymorphism-based screening [9,13] to provide an initial map position for the Jbo mutation on Chromosome 3. This map position was further refined using additional affected animals to an approximately 1.5-Mb region delineated by the Eif5a2 locus and microsatellite marker D3Mit178. Direct sequence analysis of coding regions within this interval identified an A2288T transversion in the Evi1 locus, causing a nonconservative Asn763Ile change in the second of the two zinc-finger domains in this protein (Figure 1A and 1B). No other sequence changes were identified in any other coding sequences within the minimal nonrecombinant region containing the mutation.
Figure 1
The Evi1 Gene Is Mutated in Junbo Mice
(a) Sequence analysis of the Evi1 locus in BALB/c, C3H/HeN, Jbo/+ adult, and Jbo/Jbo embryonic DNA. An A2318T transversion is detected in Jbo/+ and Jbo/Jbo mutants that is not present in either parental substrains.
(b) Schematic of the EVI1 peptide. Ten zinc finger motifs are clustered into two DNA-binding domains, ZF1 and ZF2. EVI1 contains a proline-rich repressor domain between the two sets of zinc fingers and a highly acidic domain at the C-terminus. Expanded peptide sequence across the ninth zinc finger motif shows the high degree of conservation of this region between orthologous proteins from different species: Mus musculus (P14404), Rattus norvegicus (ENSRNOG00000012645), Homo sapien (Q03112), Danio rerio (ENSDARP00000008993), Fugu rubrides (ENSDARP00000008993), Drosophila melanogaster (CG31753), Caenorhabditis elegans (R53.3a). Contact residues are highlighted in yellow, the position of the Junbo mutation is highlighted in red.
(c) Extra digits are seen on the forelimbs of both heterozygote and homozygote mice E18.5 (white arrows). In heterozygotes, (Jbo/+, middle panel) an extra digit is observed on either forelimb. The homozygotes, (Jbo/Jbo, right panel) have extra digits on both forelimbs. The anterior digit is often reduced in size in the homozygote limbs (red arrow). Wild-type mice, left panel.
A knockout allele of Evi1 has been produced, Evi1tm1Mmor [14]. Evi1tm1Mmor mice carry a targeted deletion resulting in an isoform-specific null for the longest Evi1 transcripts, while the Δ324 (shorter) isoform [15,16] remains unaltered. This isoform lacks the final two zinc-finger motifs from the first (N-terminal) DNA-binding domain and consequently is predicted to exhibit altered binding abilities. Characterization of heterozygote mice carrying this targeted mutant failed to uncover any phenotype, while homozygotes show prenatal lethality from presumptive cardiac failure [14]. In addition, these embryos displayed widespread hypocellularity, most markedly of the cortical mesenchyme, retarded development of the first and second branchial arches, and severely reduced vasculature of the yolk sac. To confirm that the A2318T alteration identified in Evi1 is responsible for the Junbo phenotype, we undertook a complementation screen between Jbo/+ and Evi1tm1Mmor/+ animals. 81 live births were produced from crosses of Jbo/+ and Evi1tm1Mmor/+ mice and all were genotyped. None of the progeny coinherited the Evi1tm1Mmor and the Jbo mutations. However, the other expected genotypes were recovered—Jbo/+ (38%), Evi1tm1Mmor/+ (28%), +/+ (33%)—confirming the allelism of the knockout and Junbo mutant. We established intercrosses between Jbo/+ animals using in vitro fertilization and implantation into wild-type females as Jbo/+ females undergo repeated spontaneous abortion. At 10.5 days postcoitum (dpc), 17% of homozygote mice had small hind limbs, a large pericardial sac, and a malformed forebrain, features demonstrated by the knockout [14]. However, many homozygotes were scored as normal in appearance at this and later stages and proceeded to die between E18.5 and birth, though homozygotes were of comparable body size to wild-type littermates. It appears that though some homozygote embryos may die at 10.5 dpc, others continue to develop unhindered when carried by a wild-type mother. The only other phenotypic feature detected in Jbo/+ mice was an extra digit on one forelimb. Jbo/Jbo mice have extra digits on both forelimbs (see Figure 1C).
Pathology Phenotyping of the Junbo Mutant
To determine the cause of the deafness phenotype in Junbo, we performed an initial examination of deaf adult animals and embryonic skeletal preparations (15.5 dpc–18.5 dpc). This revealed no gross morphological defects of the inner ear and ossicular chain or ossification of the temporal bone and tympanic ring in Jbo/+ animals. Dissection of the bullae of seven Jbo/+ mice and four +/+ mice (>130 DAB) revealed that the middle ear cavity (MEC) of mutant mice was filled with exudate, indicative of OM. X-ray analysis revealed that there was no consistent abnormality of bullae shape associated with OM in Jbo/+ mice. Examination of seven Jbo/+ mice (14 bullae, >180 DAB) revealed that 6 Jbo/+ mice bullae had a normal shape and demonstrated radio opacity, indicating the presence of an effusion in the middle ear, whereas in six Jbo/+ mice, bullae were misshapen and also had an effusion present (demonstrated by radio opacity). Although X-ray imaging is a less sensitive means of diagnosing OM than histology, two Jbo/+ mice bullae did not demonstrate a radio opacity reflecting the possibility of resolution of a small number of Junbo mice at advanced age (see below). Control ears (two wild-type mice, four bullae) were clear of effusion and presented with a normal bullae shape. We concluded that the presence of abnormally shaped bullae in the Junbo mutant is not necessary for the development of an effusion. We therefore proceeded to study the pathology of the OM in the postnatal period, at weaning, and in the adult, first in conventionally housed mice of relatively low microbiological status and then in specific pathogen-free (SPF) mice.
In conventionally housed mice we found no evidence of inflammation of the embryonic connective tissue or exudation into the MEC in Jbo/+ or wild-type litter mates at 5 DAB. By 13 DAB the MEC is more fully formed and 100% of the Jbo/+ mice and ~33% wild-type mice had acute OM (Figure 2A–2D). However, Jbo/+ mice had suppurative exudate in both the MECs, while wild-type mice had unilateral OM, or serous exudation in one ear and suppurative exudation in the contralateral ear. Typical acute inflammatory changes in the MEC mucoperiosteum included edematous polyps (Figure 2E, which may represent unresorbed embryonic connective tissue [17]) and/or bulging sub-epithelial bullae filled with serous fluid, stromal edema, widely patent capillaries, and lymphatics, with variable numbers of infiltrating neutrophil leukocytes. The epithelial covering was formed by small basal cells or ciliated columnar cells (Figure 2F and 2G). A second, milder type of OM occurred in a further ~27% of wild-type mice consisting of only focal mucoperiosteal aggregations of neutrophil leucocytes with or without a light neutrophil leukocyte MEC exudation (unpublished data); ~40% wild-type mice had no evidence of OM at this age. Importantly, OM appeared at this stage as part of a more generalized respiratory tract inflammation comprising suppurative rhinitis (Figure 2H) and nasopharyngitis, and multifocal mild alveolitis/interstitial pneumonia with eosinophil leukocytes in perivascular cuffs (Figure 3A and 3B) but without evidence of intralesional bacteria or viral inclusions. There were no significant histological lesions in intestine, liver, pancreas, kidney, heart, thymus, and spleen.
Figure 2
Histology of Middle Ear and Nose in Wild-Type and Junbo Mutant Mice
Images (a–h) are from 13-d-old postnatal mice and are given with their original magnification (a) Jbo/+ dorsal section of MEC partly filled with exudate ×40, (b) +/+ normal middle ear temporal bone covered with thin mucoperiosteum (arrowheads) ×400, (c) +/+ inflamed middle ear with thickened mucoperiosteum with neutrophil leukocyte infiltrates and neutrophil-rich exudates in the MEC ×400, (d) Jbo/+ middle ear with more severe suppurative exudation into the MEC ×400, (e) Jbo/+ inflamed edematous polyp (arrowhead) of un-resorbed embryonic middle ear connective tissue, tympanic membrane ×100, (f) Jbo/+ MEC lined by ciliated columnar cells (arrowhead) ×600 (g) Jbo/+ MEC lined by basal cells (arrowhead) ×600, (h) Jbo/+ suppurative rhinitis: nasal cavity with suppurative exudate, nasal septum with inflamed nasal mucosa ×200.
Images (i–l) are of adult (180-d) Jbo/+ middle ear with chronic suppurative OM; changes include (i) fibrous polyps (arrowheads) ×200, (j) hyperplasia of ciliated epithelial cells (arrowhead) and fibrosis of mucoperiosteum stroma ×400, and (k) fibrous thickening of the tympanic membrane, outer ear canal ×400 compared with (l) normal +/+ tympanic membrane, outer ear canal ×400. E, exudate; MP, mucoperiosteum; NC, nasal cavity; NM, nasal mucosa; NS, nasal septum; OEC, outer ear canal; TB, temporal bone; TM, tympanic membrane
Figure 3
Histology of the Lung and Middle Ear Exudate in Junbo Mice
(a) 13-d postnatal Jbo/+ lung with perivascular and peribronchiolar cuffs containing Sirius red positive eosinophil leukocytes (arrowhead), bronchiole, pulmonary artery ×400, (b) focal eosiniphilic alveolitis (arrowhead) and thickened alveolar septae (open arrowhead) with eosinophil-rich infiltrates. (c) 21-d postnatal Jbo/+ MEC pus with colonies of Gram positive cocci ×600. B, bronchiole; PA, pulmonary artery
Any postnatal OM had resolved in wild-type mice by weaning (0% incidence at 21 DAB) and OM was exceptional in adults (3% incidence). In contrast, OM was present in 100% of Jbo/+ mice at weaning and occurred in 94% of adult Jbo/+ mice 29 DAB to >180 DAB. Suppurative rhinitis was still present in some Jbo/+ mice examined, but in two the eosinophilic pneumonia was absent or minimal. Large numbers of Gram positive cocci were present in OM exudate at day 21 in ~70% of cases (Figure 3C), suggesting nasopharyngeal staphylococcal and streptococcal flora may play a role in progression of OM, if not its initiation in Jbo/+ mice. There was no evidence of significant proliferation of mucous cells or periodic acid-Schiff-positive mucus in the MEC.
Adult Jbo/+ mice >29 DAB develop bilateral chronic suppurative OM. Chronic middle ear effusions contained variable numbers and proportions of viable and necrotic neutrophil leukocytes and foamy macrophages; bacteria were infrequently present and there were small numbers of multinucleate macrophages, cholesterol clefts, and small amounts of birefringent foreign body (perhaps secondary to perforation of the eardrum). The effusions did not contain significant amounts of periodic acid-Schiff-positive mucus. Chronic changes in inflamed thickened mucoperiosteum include formation of multiple polyps (Figure 2I) covered by low cuboidal cells or hyperplastic ciliated columnar cells and scattered mucous cells (Figure 2J). The mucoperiosteal stroma has variable degrees of fibrosis, neutrophil leukocyte infiltration, scattered mast cells, lymphoplasmacytic infiltrates, and occasional lymphoid nodules. The eustachian tube is patent and can contain exudates, and the epithelial lining can have elevated numbers of mucous cells and intra-epithelial leukocytes. Fibrous thickening of the eardrum (tympanosclerosis) is common (Figure 2K and 2L). OM was often associated with perforation of the eardrum, possibly providing drainage to account for apparent resolution of a few cases: (6%) of OM in the oldest age group of Jbo/+ mice (>180 DAB).
The Junbo colony has subsequently been re-derived by embryo transfer into new high-health status facilities (Mary Lyon Centre, Harwell). Mice are housed in individually ventilated cages, and screening shows all FELASA-listed pathogens (see Materials and Methods) have been excluded; 75 cage air changes/h reduce respiratory irritants such as ammonia to <3 ppm. In SPF conditions, OM in Jbo/+ mice is relatively milder at early time points and is not associated with rhinitis. Typically at 13 DAB there are small numbers of inflammatory cells in the mucoperiosteum and sometimes, light MEC effusion. By 20–22 DAB and 28 DAB, 4% Jbo/+ mice had bilateral OM, 54% unilateral OM, and 42% had very mild or no OM. However, by 54 DAB, Jbo/+ mice (100%) had OM, but in 10% of cases this was unilateral. OM does not occur in pre-weaned wild-type mice and in only 3% (a single case in a 22-DAB mouse) of older wild-type mice.
Comprehensive pathology phenotyping failed to show a consistent pattern of significant organ pathology outside of the middle ear in adult Jbo/+ mice. Specifically there is no evidence of opportunistic infections in sites such as skin, lung, urogenital, or gastrointestinal systems that might be a sign of immune deficiency (see below).
In summary, the microbiological status of the mice and/or air quality affect onset of OM, and under “dirty” conventional conditions even wild-type mice can develop OM in the postnatal period as part of upper respiratory tract disease; however, this resolves in wild-type mice by weaning. In conventionally housed and SPF Jbo/+ mice, OM emerges as a chronic condition.
Immunology and FACS Analysis of SPF Wild-Type and Junbo Mice
There were no significant differences in the antibody responses of Jbo/+ and wild-type control mice to immunization with T-dependent (IgG1 and IgG2a to keyhole limpet hemocyanin) and T-independent (IgGM and IgG3 to pneumococcal polysaccharide type 3) antigens (Table S1). FACS analysis of blood neutrophils identified by the cell surface markers Gr-1 and Mac-1 did not reveal significant differences between levels of immature, mature cell forms, and total circulating blood neutrophils in Jbo/+ and wild-type mice at either 20–22 DAB or 49–58 DAB. However, there was a significantly lower (p = 0.003) ratio of immature forms in the circulating neutrophil pool in 20–22 DAB Jbo/+ mice, but not in 54–58 DAB Jbo/+ mice (Table 1).
Table 1
FACS Analysis of the Proportion of Granulocytes in Blood of Wild-Type and Junbo Mice
Expression of Evi1 in Wild-Type and Junbo Mice
We proceeded to investigate the expression of Evi1 in both embryonic whole-body tissues (E9.5 and at eight intervals through to birth) and postnatal middle ear tissue (13 DAB, 21 DAB) in order to relate the underlying mutation to the Junbo phenotype. We examined expression in wild-type and Jbo/+ mice, but at no time point did we identify any significant differences in expression patterns, up to and including birth, from those described [14,18]. However, in postnatal head tissues, we now find that Evi1 is expressed in nuclei of myeloid cells in bone marrow, neutrophil leukocytes, fibroblasts, and basal epithelial cells in the inflamed middle ear lining (Figure 4), and that patterns of expression are similar in Jbo/+ and wild-type mice.
Figure 4
Evi1 Protein Immunostaining in 13-d-Old Jbo/+ and Wild-Type Mice with Acute OM
(a) Jbo/+ positive myeloid cells in temporal bone marrow ×600. Note chondrocytes and osteocytes are also strongly positive. (b) Jbo/+ mucoperiosteum has positive neutrophil leukocytes, fibroblasts, and basal epithelial cell nuclei ×600. (c) +/+ similar pattern of staining in bone marrow ×600 and (d) +/+ inflamed mucoperiosteum ×600. B, basal epithelial cell nuclei; C, chrondrocyte; F, fibroblast; MC, myeloid cell; N, neutrophil leukocyte; O, osteocyte
OM Phenotype of the Evi1tm1Mmor/+ Mice
OM is not a prominent feature in Evi1tm1Mmor/+ mice. Only one of 12 Evi1tm1Mmor/+ mice >68 DAB had OM, and in this single case it was unilateral. OM was not present in 15 wild-type littermates. The absence of OM in Evi1tm1Mmor/+ mice, coupled with the presence of extra digits in Jbo/+ and Jbo/Jbo mice, suggests that the Junbo mutation may have gain-of-function effects. However, the genetic background of Evi1tm1Mmor/+ mice differs from Jbo/+ and Jbo/Jbo mice (see Materials and Methods). In addition, the Evi1tm1Mmor mutation is an isoform- specific null. It is therefore difficult to reach definitive conclusions as to the nature of the Junbo mutation (see Discussion).
Discussion
A mutation in the Evi1 transcription factor underlies the development of a chronic suppurative OM in the Junbo mutant. Genetic mechanisms appear to interact with microbiological status and environmental conditions, such that SPF Jbo/+ mice with improved air quality have milder early OM. Gnotobiotic studies are planned to investigate the role of nasopharyngeal Staphylococcus and Streptococcus spp. commensals in OM pathogenesis. The absence of multi-systemic inflammation and opportunistic infections at other sites, and the normal T-dependent and T-independent immune responses in Jbo/+ mice, argue against an overt immune deficiency being responsible for OM.
The Evi1 locus was initially identified as a common site of retroviral integration underlying susceptibility to myeloid tumors in the AKXD mouse recombinant inbred strain [19,20]. Transcriptional activation of the Evi1 locus by translocations and inversions leads to myeloid leukemias and myelodysplastic syndrome in humans [21]. This locus encodes a 145-kDa nuclear transcription factor with two distinct zinc-finger domains composed of seven and three zinc-finger motifs, respectively [22]. Each zinc-finger domain has been shown to bind specific target consensus DNA sequences, and additional proline-rich and highly acidic domains located within the protein have been demonstrated to be capable of repressing or activating target promoter activities, respectively [21–24]. Evi1 has also been shown to be capable of repressing the TGF-β signaling pathway through direct binding of Smad3 mediated by the first zinc-finger domain, suggesting a functional role in the control of cell development and proliferation [25]. In addition, cell line assays have demonstrated a role for Evi1 in the transcriptional control of c-fos and the AP-1 proliferative pathway through interactions of the second zinc-finger domain [26]. The Junbo mutation results in a nonconservative Asn763Ile change in the second of these three zinc-fingers. Previous in vitro site-directed mutagenesis studies of the contact residues within these three zinc-fingers uncovered a complete loss of DNA-binding ability following any residue change [23]. The Junbo Asn763Ile alteration is within three amino acids of a contact residue, and the mutated amino acid contributes to the putative alpha helix of the Cys2His2 structure. While it seems likely that the Junbo mutation will disturb the function of the second zinc-finger domain, it is unclear whether or not there are effects on the role of the first zinc-finger domain that is involved in TGF-β signaling.
It is interesting to note that OM is not part of the phenotype of Evi1tm1Mmor/+ mice. However, the Evi1tm1Mmor allele results in an isoform-specific allele for the longest Evi1 transcript while the Δ324 shorter isoform is unaffected. In addition, the Evi1tm1Mmor knockout was established in ES-D3 cells with chimaeras subsequently mated to CF-1 mice [14]. Both the presence of the shorter isoform and the dissimilar genetic backgrounds between Evi1tm1Mmor/+ mice and Jbo/+ mice may contribute to the differences in the expressivity of the OM phenotype. Alternatively, the Junbo mutation may lead to gain-of-function effects. However, it is not possible at this stage to distinguish between these possibilities.
A mutation in the Evi1 transcription factor may give rise to OM by more than one mechanism, given that it is expressed in a number of different cell types in the middle ear. One mechanism arises from our observation that Evi1 is expressed in neutrophil leukocytes during OM development. Evi1 has multiple functions relating to hematopoietic differentiation and development of myeloid leukemia [27–30]. One putative target gene for Evi1 in neutrophil leukocytes is the inositol triphosphate type 2 receptor gene (Itpr2) that is required for functional regulation of neutrophil leukocyte maturation via F-met-leu-phe receptor signaling in response to bacterial proteins [31]. Our FACS analysis of circulating neutrophils at two time points, when OM first appears and when chronic suppurative OM is well established, indicates that localized inflammation in the middle ear in Jbo/+ mice does not result in significant neutrophilia. Neutrophils are apparently released from hematopoietic tissues at comparable levels in Jbo/+ and wild-type mice and there was no detectable block in neutrophil development in Jbo/+ mice. The ratio of immature neutrophils in the circulating pool was no higher; indeed, it is reduced in recently weaned mice when ~60% have acute OM. In older Jbo/+ mice with fully penetrant chronic suppurative OM, immature and mature neutrophil ratios are not significantly different from wild-type mice.
A variety of in vitro studies have highlighted the role of the TGFβ/SMAD pathway on mucin expression and thus underlined the potential importance of this signaling pathway in OM [32]. A loss of Evi1 function could affect TGFβ/SMAD signaling pathways. There are two in vitro studies that suggest that Evi1 mutations could affect mucin expression that might underlie predisoposition to OM. Firstly, nontypeable Haemophilus influenzae (NTHi), a known bacterial pathogen involved in human OM, activates Tgfβ receptor-Smad3/4 signaling that together with TLR2-MyD88-TAK1-NIK-IKKβ/γ-IκBα-dependent activation of NF-κB is known to mediate NTHi-induced MUC2 mucin transcription [33]. Evi1 loss-of-function mutations might lead to a de-repression of the TGFβ/SMAD pathway and an upregulation of MUC2 expression leading to an enhancement of effusive processes as a contributor to OM. Alternatively, it has also been shown that NTHi upregulates MUC5AC mucin production via activation of the TLR2-MyD88-dependant p38 pathway [34]. However, the activation of TGFβ/SMAD signaling by NTHi also leads to down-regulation of p38 activity by inducing MAPK phosphatase-1 and suppressing MUC5AC mucin induction. Thus, in this case loss of Evi1 function and de-repression of the TGFβ/SMAD pathway would presumably lead to increased suppression of MUC5AC mucin induction that may reduce mucociliary defense in suppurative OM. Importantly, the identification of the Junbo mutation now provides in vivo evidence to support the role of these signaling pathways in OM. It will be interesting to explore further the molecular phenotype of the Junbo mutant and to examine whether disregulation of mucin expression is a contributing factor in the development of OM.
Additional levels of complexity of Evi1 function relevant to OM pathogenesis may result from the first (N-terminal) zinc-finger domain binding to a number of putative target genes Gadd45g, Gata2, Zfpm2/Fog2, Skil (SnoN), Klf5 (BTEB2), Dcn, and Map3k14 (Nik) [35]. For instance, Evi1 regulation of Nik could act in the TLR2-MyD88-TAK1-NIK-IKKβ/γ-IκBα-dependent activation of NF-κB pathway to mediate NTHi-induced MUC2 mucin transcription as described above; and also in the pro inflammatory cytokine IL-1 signaling pathway IL-1R1-MyD88-IRAK-TRAF6-TAK1-NIK-IKK-NF-κB/IκB [36].
In conclusion, the Junbo mutant provides an important genetic disease model of OM; particularly because it shares important features with the chronic human condition [7]. Inflammatory disease is restricted to the middle ear and is not a consequence of overt immune deficiency. OM arises spontaneously in the postnatal period, develops into chronic suppurative OM with otorrhea, with the early severity and age of onset dependent on microbiological status and/or air quality. We have shown that a mutation at the Evi1 locus underlies the susceptibility and persistence of OM. Our observations underline the role of Evi1 in mucin gene regulation as a possible contributor to OM. In this regard, the Evi1 gene and associated pathway members can be considered important candidates for examining the genetic basis for susceptibility to OM in the human population.
Materials and Methods
Mice and deafness screening.
The founder mouse carrying the Junbo mutation was generated in a large-scale ENU mutagenesis program at Harwell, United Kingdom [9]. Male BALB/c mice were mutagenized and mated to normal C3H/HeN females, and the offspring were screened for a variety of defects, including deafness and vestibular dysfunction. The Jbo founder was identified because of a lack of a Preyer reflex when presented with a calibrated 20 kHz 90 dB SPL tone burst. For analysis of the phenotype, the colony was maintained on a C3H/HeN background in accordance with Home Office regulations.
Microbiological status of the mice.
Junbo mice were originally derived in a conventional facility. Sentinel mice from this colony were seropositive for the FELASA- [37] listed viral agents MHV (judged by histology to be enteropathic strains), Adenovirus II, and TMEV, none of which are primary respiratory pathogens [38]. Intestinal flagellates, pinworms, and the opportunist respiratory pathogen Pasteurella pneumotropica were also common isolates. A number of pathogens known to cause respiratory disease and/or OM in the mouse such as pneumonia virus, Sendai virus, Mycoplasma pulmonis, Streptococcus pneumoniae, and Pseudomonas aeruginosa have not been found over many years in the sentinel screens. Non-FELASA-listed bacteria isolated from the nasopharynx of sentinel mice include Staphylococcus spp., Staphylococcus aureus, Alpha-haemolytic streptococci, and other Streptococcus spp. Junbo mice are now housed in a high- health-status SPF unit in which all FELASA-listed pathogens have been excluded. Non-FELASA nasopharyngeal flora remains the same.
Genetic crosses and mapping.
The founder C3HeNBALB/cENUF1-Jbo/+ mouse was maintained by repeated backcrossing to C3H/HeN. Mutant progeny were identified by the lack of a Preyer reflex. 41 DNAs were initially pooled from affected individuals and a genome scan was carried out as described [9,13]. A high-resolution genetic map was constructed using 242 affected progeny N4 and N5 backcross progeny using published microsatellite markers D3Mit90, D3Mit328, D3Mit92, D3Mit203, D3Mit55, D3Mit178, D3Mit151, D3Mit273, D3Mit180, D3Mit239, D3Mit21, D3Mit224, D3Mit182, D3Mit119, D3Mit241, and D3Mit22. The genetic map across the Jbo locus was constructed by minimizing the number of recombination events across the region.
The Evi1tm1Mmor mutation was generated in D3 ES cells with subsequent matings to CF-1 mice [14]. Evi1tm1Mmor/+ mice were mated to C3H/HeN mice before crossing to Jbo/+ mice for complementation testing. For phenotypic analysis Evi1tm1Mmor/+ mice were also mated to C3H/HeN mice and F1 heterozygous mutant progeny along with wild-type littermate controls aged to appropriate time points.
Pathology, histology, and X-Ray analysis.
The onset and time course of development of the middle ear disease was examined by histology in conventionally housed postnatal mice 4–5 DAB (five Jbo/+, two +/+); postnatal 13 DAB (13 Jbo/+, 16 +/+); at weaning 21 DAB (seven Jbo/+, 17 +/+); and in adult mice 29 DAB (five Jbo/+, 11 +/+), 44 DAB (six Jbo/+, eight +/+), 180 DAB (seven Jbo/+, nine +/+), and 180–360 DAB (six Jbo/+, three +/+). Representative mice from one to three litters were examined at each time point. In a second study to assess the OM phenotype under SPF conditions, Junbo mice were sampled from an embryo re-derived colony that was housed in individually ventilated cages. The cohorts were as follows: 5 DAB (four Jbo/+, nine +/+), 13 DAB (11 Jbo/+, 19 +/+), 20–22 DAB, 28 DAB (24 Jbo/+, 17 +/+), 49–58 DAB (20 Jbo/+, 18 +/+), and 85–115 DAB (19 Jbo/+). 12 conventionally housed 68-DAB Evi1tm1Mmor/+ mice and 15 same-aged wild-type littermates were also examined for OM.
Tissues were fixed 24–48 h in 10% neutral buffered formalin (NBF) and embedded in paraffin wax. Heads and bones were decalcified 24–48 h with Immunocal (Decal Corporation, Tallman, New York, United States). 4-μm dorsal plane sections of decalcified middle ears were stained with Haemotoxylin and Eosin (H & E). In 13- and 21-DAB mice with OM, transverse sections of the nasal turbinates, dorsal plane sections of the oropharynx, lungs, livers, kidneys, heart, spleen, intestines, pancreas, and livers were examined by histology. Sections of middle ears, snouts, and lungs from OM cases were examined for bacteria using Gram stain and Sirius red for eosinophil leukocytes.
To assess the possibility of opportunistic infections at other body sites, a general pathology screen was performed on four 28-DAB and seven 56-DAB SPF Jbo/+ mice. In addition, a more extensive whole-body pathology analysis of 25+ tissues following EMPReSS necropsy SOPs [39] was performed on four female and three male 180-DAB Jbo/+ mice; five female and four male wild-type litter mates were used as controls.
For X-ray analysis, heads were skinned and the brains and mandibles removed, and then the skull was fixed in 10% NBF. After the dorsoventral view was taken, the head was bisected midsagittally and lateromedial views taken. Images were taken on a MX-20 Faxitron X-ray machine (Faxitron X-ray Corporation, Wheeling, Illinois, United States) at 26 kV and 0.3 mA with an exposure time of 3 s.
Immunohistochemistry.
Whole embryos (E9.5, E10.5, E11.5, E12.5, E13.5, E16.5, and E18.5) and adult heads (13 DAB and 21 DAB) from wild-type and Jbo/+ mice were used for immunolabelling. 3-μm wax sections were de-paraffinised in xylene substitute and rehydrated through graded ethanol solutions. Endogenous peroxidase activity was quenched with 3% hydrogen peroxide in isopropanol. The sections were microwaved in 10 mM citrate buffer (pH6.0) and rinsed with phosphate-buffered saline at room temperature. The immunostaining was performed using a Dako (Glostrup, Denmark) autostainer at room temperature. To inhibit the non-specific endogenous biotin staining the Dako Biotin Blocking System was used. A blocking solution of 10% donkey serum (Serotec) was used for 1 h. Goat polyclonal antibody raised against the carboxy terminus of human EVI1 was used in concentration 1:100 (Santa Cruz Biotechnology, Santa Cruz, California, United States) for 1 h. Biotinylated donkey anti-goat antibody (Santa Cruz Biotechnology) and ChemMate Detection Kit (Dako) were used to develop the specific EVI-1 signals. Negative control sections were incubated in donkey serum and processed identically. The slides were counterstained with Haematoxylin.
Skeletal preparations.
Embryos were fixed in 95% ethanol and processed through a standard alcian blue and alizarin red bone/cartilage staining procedure.
Immunology and FACS analysis in SPF mice.
The T-dependent arm of the immune system was assessed by immunization with keyhole limpet hemocycanin and measurement of IgG1 and IgG2a, and the T-independent arm by immunization with pneumococcal polysaccharide type 3 and measurement of IgGM and IgG3 in 42–56-DAB SPF mice (ten Jbo/+, ten +/+) [40]. For FACS analysis whole blood was collected in lithium heparin tubes from the jugular vein after overdosing mice with barbiturate-administered IP. SPF mice were assessed at 20–22 DAB (17 Jbo/+, 20 +/+) and 49–58 DAB (14 Jbo/+, 14 +/+). FACS analysis of granulocytes was performed after labeling cells with R-PE-labeled Gr-1 and FITC-labeled Mac-1 markers. The Wilcoxon sum-of-ranks test was used to test for statistical differences between Jbo/+ and +/+ post-bleed antibody titers and granulocyte parameters.
Supporting Information
Table S1
T-Dependent and T-Independent Responses in Immune- Challenged Wild-Type and Junbo Mice
(35 KB DOC)
Click here for additional data file.
Acknowledgements
We thank Dr. Emma Coghill for advice on using the Amera software and Stuart Townsend for assistance with FACS analysis. We also thank the Histology and Pathology teams at Harwell, United Kingdom for processing of specimens.
Abbreviations
DAB - days after birth
dpc - days postcoitum
ENU - N-ethyl-N-nitrosourea
MEC - middle ear cavity
NTHi - nontypeable Haemophilus influenzae
OM - otitis media
SPF - specific pathogen-free
Footnotes
Competing interests. Part of this work was funded by GlaxoSmith-Kline.
Author contributions. NP, REHH, HT, ND, AJH, MTC, and SDMB conceived and designed the experiments. NP, REHH, HT, HTT, DB, SM, ZL, FM, MF, PG, AMW, SP, IB, TAH, and MTC performed the experiments. NP, REHH, HT, HTT, DB, SM, ZL, FM, ND, and MTC analyzed the data. MF, PG, AMW, and SP contributed reagents/materials/analysis tools. NP, REHH, MTC, and SDMB wrote the paper.
Funding. This work was supported by the Medical Research Council, United Kingdom. Hilda Tateossian is supported by the Eumorphia program (European Commission contract QPG2-CT-2002–00930). | [
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17201918 | SirT1 modulates the estrogen–insulin-like growth factor-1 signaling for postnatal development of mammary gland in mice
Abstract
Introduction
Estrogen and insulin-like growth factor-1 (IGF-1) play important roles in mammary gland development and breast cancer. SirT1 is a highly conserved protein deacetylase that can regulate the insulin/IGF-1 signaling in lower organisms, as well as a growing number of transcription factors, including NF-κB, in mammalian cells. Whether SirT1 regulates the IGF-1 signaling for mammary gland development and function, however, is not clear. In the present study, this role of SirT1 was examined by studying SirT1-deficient mice.
Methods
SirT1-deficient (SirT1ko/ko) mice were generated by crossing a new strain of mice harboring a conditional targeted mutation in the SirT1 gene (SirT1co/co) with CMV-Cre transgenic mice. Whole mount and histology analyses, immunofluorescence staining, immunohistochemistry, and western blotting were used to characterize mammary gland development in virgin and pregnant mice. The effect of exogenous estrogen was also examined by subcutaneous implantation of a slow-releasing pellet in the subscapular region.
Results
Both male and female SirT1ko/ko mice can be fertile despite the growth retardation phenotype. Virgin SirT1ko/ko mice displayed impeded ductal morphogenesis, whereas pregnant SirT1ko/ko mice manifested lactation failure due to an underdeveloped lobuloalveolar network. Estrogen implantation was sufficient to rescue ductal morphogenesis. Exogenous estrogen reversed the increased basal level of IGF-1 binding protein-1 expression in SirT1ko/ko mammary tissues, but not that of IκBα expression, suggesting that increased levels of estrogen enhanced the production of local IGF-1 and rescued ductal morphogenesis. Additionally, TNFα treatment enhanced the level of the newly synthesized IκBα in SirT1ko/ko cells. SirT1 deficiency therefore affects the cellular response to multiple extrinsic signals.
Conclusion
SirT1 modulates the IGF-1 signaling critical for both growth regulation and mammary gland development in mice. SirT1 deficiency deregulates the expression of IGF-1 binding protein-1 and attenuates the effect of IGF-1 signals, including estrogen-stimulated local IGF-1 signaling for the onset of ductal morphogenesis. These findings suggest that the enzymatic activity of SirT1 may influence both normal growth and malignant growth of mammary epithelial cells.
Introduction
Mammalian SirT1 belongs to a family of nicotinamide adenine dinucleotide-dependent histone deacetylases [1,2]. SirT1 is most closely related to yeast Sir2, the founding member of the evolutionarily conserved Sir2 family. Yeast Sir2 is required for silencing transcription at the telomeric region and mating type loci, and for suppression of ribosomal DNA recombination [3,4]. The expression of an extra copy of Sir2 in either yeast mother cells or multicell organisms such as nematodes can significantly extend the lifespan [5,6]. Inactivation of Sir2 enhances stress resistance and extends chronological lifespan of nondividing yeast cells, which is opposite to the requirement for Sir2 function in the reproductive lifespan [7]. Whether SirT1 regulates the reproductive lifespan and/or the chronological lifespan in mammals remains unknown.
Sir2 is an integral part of an evolutionarily conserved insulin/insulin-like growth factor-1 (IGF-1) signaling (IIS) system in worms (Caenorhabditis elegans), fruit flies (Drosophila), mice, and humans [8,9]. The IIS system includes membrane-bound receptors, cytoplasmic kinases, and nuclear transcription factors. To maintain the proper expression of the effector genes for the IIS system, these conserved components form a sophisticated regulatory system, which centers on a family of forkhead transcription factors (forkhead box 'other' proteins (FoxOs)), and operates on two levels. On one level, SirT1-mediated protein deacetylation attenuates the transcriptional activity of nuclear FoxO transcription factors [10-12]. On the second level, the FoxO transcription factors can be sequestered within the cytoplasm when phosphorylated by activated Akt kinases in response to insulin and IGF-1 signals [13]. Conceivably, the IIS system senses the levels of insulin and IGF-1 and negatively regulates the expression of the effector genes. The IIS system is responsible for food storage, stress tolerance, and longevity in lower organisms, such as C. elegans [8,9,14]. In more advanced species, steroid hormones evolved to regulate the IIS system [15]. In mice and humans, the IGF-1 signaling of the IIS system mediates local effects for growth and hormonal regulation for multiple tissues, including mammary glands [16,17].
Mammalian SirT1 has evolved to modify the activity of a growing number of transcription factors, including p53, NF-κB, and PGC-1α, suggesting that SirT1 functions in a wide range of cellular responses to stress, inflammation, and nutrients [18-21]. SirT1-deficient mice display characteristic phenotypes of perinatal death and growth retardation as well as other diverse phenotypes, such as eye defects, with varying severity [22,23]. The underlying causal mechanism for these phenotypes, however, remains unknown. We recently generated SirT1-deficient (SirT1ko/ko) mice and found that both male and female SirT1ko/ko mice can be fertile, which is in contrast to the sterile phenotypes observed in one strain of SirT1-deficient mice [22]. This led to our study of the link between SirT1 and IGF-1 signaling using the mammary gland as a model organ.
The mammary gland is a unique organ because it develops after birth and undergoes dynamic changes throughout the reproductive lifespan of a female. At the onset of puberty, ovarian estrogen stimulates ductal morphogenesis during which mammary epithelial progenitor cells differentiate and proliferate while interacting with adipocytes and stromal cells within mammary fat pad [17,24,25]. Ovarian estrogen, in synergy with pituitary growth hormone (GH), stimulates stromal cells to produce local IGF-1. The local IGF-1, but not liver-produced systemic IGF-1, provides a paracrine signal for commencing ductal morphogenesis [26]. Mice lacking GH, estrogen, IGF-1, GH receptor, or estrogen receptor alpha (ERα) fail to undergo postnatal ductal morphogenesis [17,24-31], indicating that both steroid hormones and IGF-1 are on the common pathway for a critical developmental checkpoint. Once the arborated ductal network is established, cycles of differentiation, proliferation, and death of secretory alveolar epithelium repeat with each pregnancy [17,24,25].
In the present article we report the finding and characterization of impeded ductal morphogenesis in virgin SirT1ko/ko mice and lactation failure in SirT1ko/ko mothers. The characterization of these phenotypes has identified a SirT1-dependent regulatory mechanism by which SirT1 modulates the effectiveness of the estrogen–IGF-1 signaling for mammary gland development. The estrogen–IGF-1 signaling is defined as the ovarian estrogen-regulated, stromal cell-produced local IGF-1 signal for stimulating mammary epithelial cells.
Materials and methods
Mice
A previously described SirT1 targeting construct, KOII [23], was used to generate mice harboring a conditional targeted mutation in the SirT1 gene (SirT1co/co mice) (see Additional file 1). The breeding of SirT1co/co mice and CMV-Cre transgenic mice results in mice harboring a germline-transmitted deletion of exon 4 of the SirT1 gene (SirT1+/ko mice). Both SirT1co/co mice and SirT1+/co mice were used to establish breeding colonies for generating SirT1co/co and SirT1ko/ko mice, respectively. Both SirT1co/co mice and SirT1ko/ko mice were in a mixed 129SvJ/C57B6 background. Mice were housed in a special-pathogen-free facility and all procedures were approved by the University of Washington Animal Care and Use Committee. A PCR-based genotyping method was established to identify the wild-type, co, and ko loci of the SirT1 gene using three primers: 5' co primer, 5'-GGTTGACTTAGGTCTTGTCTG; 5' ko primer, 5'-AGGCGGATTTCTGAGTTCGA; 3' primer, 5'-CGTCCCTTGTAATGTTTCCC. Murine embryonic fibroblasts (MEFs) were isolated from the embryos between embryonic day 12.5 and embryonic day 14.5. The body weight was measured once a week.
Serum insulin-like growth factor-1
A commercial radioimmunoassay (ALPCO Inc., Windham, NH, USA) was used. Samples (300 μl) were prepared via an alcohol extraction followed by a 2-day disequilibrium incubation at 4°C. Samples were measured in duplicate and data were calculated as the samples were counted. Six standards ranging from 10 to 500 ng/ml were used to generate the standard curve. The lower limit of detection was 10 ng.
Survival assay
To determine the sensitivity of embryonic stem cells after ionizing radiation or hydrogen peroxide treatment, 300 embryonic stem cells were seeded onto a 10-cm plate for 24 hours. The plates were either exposed to a 137Cs source at indicated doses or were treated with embryonic stem media containing hydrogen peroxide at indicated concentrations for 30 minutes. Seven days after irradiation or hydrogen peroxide treatment, the embryonic stem colonies stained with crystal violet were counted.
Estrogen implantation
Estrogen pellets in the form of 17β-estradiol, at 0.1 mg, and the 21-day releasing time were obtained from Innovative Research (Sarasota, FL, USA). One pelletwas subcutaneously implanted in the subscapular region ofindividual female mice using a sterilizedmetal trochar (Innovative Research).
Whole mount analysis
The inguinoabdominal mammary fat pads were spread on microscope slides, fixed in Carnoy's fixative overnight, hydrated and stained with carmine alum stain (Sigma, St. Louis, MO, USA) overnight, and were then dehydrated, treated with xylene to remove fat, and mounted with Secure Mount (Fisher Scientific, Pittsburgh, PA, USA) and cover slips.
Histological, immunohistochemical, and immunofluorescence analyses
The mice were given a single dose of bromodeoxyurdine (BrdU) at 100 g/kg body weight via intraperitoneal injection 2 hours before euthanasia. The mammary fat pads, as well as other tissues, were collected and fixed in 10% formalin solution (Fisher Scientific). Paraffin-embedded sections were prepared at 4 μm thickness followed by standard H & E staining for histological analysis and by immunohistochemical staining for BrdU-labeled cells (Sigma) and apoptotic cells (Promega, Madison, WI, USA). To detect the presence of mouse milk proteins, a rabbit anti-mouse milk specific protein antibody (Nordic Immunology, Tillburg, The Netherlands) was used for immunofluorescent staining, followed by Texas Red anti-rabbit secondary antibody (Molecular Probes) and DAPI staining for nuclei (Molecular Probe, Eugene, OR, USA). Microscopic analyses of all histological findings were carried out on an AxioVert 200M microscope with AxioVision 4.5 software (Carl Zeiss, München-Hallbergmoos, Germany).
Protein analysis
Protein extracts were prepared from mammary tissues as well as from MEFs. The following primary antibodies were used: anti-SirT1 (Upstate Biotechology, Lake Placid, NY, USA), anti-IGF-1 binding protein-1 (IGFBP-1) (R&D Systems, Minneapolis, MN, USA), and anti-actin and anti-IκBα (Santa Cruz Biotechnology, Santa Cruz, CA, USA). The corresponding secondary antibodies were horseradish peroxidase-conjugated anti-rabbit (Pierce, Rockford, IL, USA), anti-rat, and anti-goat (Santa Cruz Biotechnology).
Results
Growth retardation in SirT1ko/ko mice
The conditional targeted SirT1 mutant mice (SirT1co/co mice) carry an insertion mutation of the neomycin-resistant gene and lox sequences in the SirT1 gene flanking exon 4 that encodes a conserved Sir2 motif (Figure 1a). The mutation does not affect the expression of SirT1 in SirT1co/co mice (Figure 1b). As expected, SirT1co/co mice are phenotypically indistinguishable from wild-type mice. To convert the SirT1 co allele into the SirT1 ko allele, SirT1co/co mice were crossed with CMV-Cre transgenic mice to generate SirT1 heterozygotes carrying the SirT1+/ko, CMV-Cre+ genotype. The expression of the CMV-Cre transgene catalyzes the deletion of exon 4 in most lineages of cells, including germ cells (Figure 1a). This SirT1 ko allele should be identical to the previously described Δex4 mutation [23]. SirT1+/ko, Cre+ mice were backcrossed with wild-type mice to generate the mice harboring a germline-transmitted deletion mutation (that is, SirT1+/ko mice). The breeding of SirT1+/ko male mice and SirT1+/ko female mice resulted in SirT1ko/ko mice. The cells derived from either SirT1+/ko mice or SirT1ko/ko mice expressed a SirT1 mutant protein due to the inframe deletion of exon 4 (Figure 1b). This SirT1 mutant protein may not be functional, however, since SirT1+/ko mice are phenotypically indistinguishable from wild-type mice and SirT1Δex4/Δex4 mice were phenotypically identical to SirT1 null mice [23].
Similar to other strains of SirT1-deficient mice, nearly two-thirds of SirT1ko/ko newborns die shortly after birth and the majority of surviving SirT1ko/ko mice manifest growth retardation (Table 1 and Figure 1c) [22,23]. Growth retardation may result from a systemic defect in hormonal regulation, DNA double-strand break repair, or other causal mechanisms. We found that SirT1ko/ko mice have reduced levels of serum IGF-1 (Figure 1d). IGF-1 often acts as a local effector for pituitary GH. The serum level of GH in SirT1ko/ko mice, however, appeared to be within the normal range (data not shown). On the other hand, yeast Sir2 and its associated Sir complex, as well as mouse SirT6, have also been implicated in DNA double-strand break repair and the maintenance of chromosome stability [32-34]. We found that SirT1ko/ko embryonic stem cells do not have increased sensitivity to either ionizing radiation or hydrogen peroxide treatment when compared with wild-type cells, or with positive control cells such as Ku70-deficient or Atm-deficient cells (Figure 1e,f). Both Ku70-deficient mice and Atm-deficient mice display growth retardation phenotypes [35,36]. The growth retardation phenotype in SirT1ko/ko mice is therefore probably not due to a defect in DNA damage repair, but rather results from a deficit in IGF-1 signaling.
Lactation failure
Both male and female SirT1ko/ko mice can be fertile, which is in contrast with the reported sterile phenotype in one strain of SirT1-deficient mice [22]. When male SirT1ko/ko mice impregnated female SirT1+/ko mice, the number of surviving SirT1ko/ko offspring was reduced due to the partial perinatal lethal phenotype (Table 1). In a reciprocal approach, male SirT1+/ko mice can also impregnate female SirT1ko/ko mice (Table 1). After parturition, SirT1ko/ko mothers exhibited normal nursing behavior. The pups did not survive for more than 3 days after birth, however, unless they were immediately removed and put under the care of a foster mother. All pups died of dehydration or starvation as a result of lack of milk in their stomachs.
To determine whether the SirT1ko/ko mothers encountered a lactation defect, whole mount and histological analyses were used to characterize the morphological changes in the mammary glands. We found that virgin SirT1ko/ko mice displayed impeded ductal morphogenesis up to 9 months of age, while age-matched virgin wild-type mice displayed extensive ductal elongation and branching (Figure 2a). The absence of ductal morphogenesis persisted in pregnant SirT1ko/ko mice up to day 13 of pregnancy (Figure 2b). Despite the fact that pregnancy can induce ductal morphogenesis at the late stage of pregnancy, underdeveloped mammary glands in SirT1ko/ko female mice cause a severe deficit in milk production, as shown by histological analysis as well as the immunofluorescence analysis using an anti-milk antibody (Figure 2b,c).
Pregnancy-induced mammary gland development
To characterize the developmental defect leading to lactation failure, we analyzed the mammary glands of SirT1ko/ko female mice on lactation day 1. Terminal end buds (TEBs) are club-shaped transitional structures (see Figure 3a, upper panel). In wild-type female mice, TEBs form and precede ductal elongation and branching at the onset of puberty [17,24,25]. When TEBs/ducts reach the edge of mammary fat pads, TEBs regress as shown in the virgin mice in Figure 3a (upper panel). Interestingly, the transitional TEBs can be readily identified in SirT1ko/ko female mice on lactation day 1, manifesting as either newly formed TEBs or as TEBs attached to developing ductal and alveolar structures (Figure 3a, lower panel). Moreover, SirT1ko/ko mice displayed varying degrees of ductal development, ranging from a lack of any ductal structure to a fully developed ductal network, the latter of which is comparable with the morphology and cellularity of wild-type mammary tissues on day 13 of pregnancy (Figure 3a). These observations indicated that pregnancy could rescue impeded ductal morphogenesis in virgin SirT1ko/ko mice.
Normal alveolar morphogenesis takes place under the control of additional hormones and growth factors during pregnancy and lactation [17,24,25]. These coordinated efforts are necessary to support a densely saturated lobuloalveolar system for secreting milk after parturition (see wild-type mice on lactation day 1 in Figures 2b and 3a). In some SirT1ko/ko mice, the alveolar morphogenesis was initiated, as indicated by the presence of a few secretory alveolar epithelial cells within scattered alveolar structures sprouted from a well-established ductal branching network (Figures 2c and 3a, lower panel). The extent of alveolar morphogenesis in SirT1ko/ko mice was inadequate, however, as shown by the severe deficit in milk production when compared with that in wild-type mice (Figure 2c).
Exogenous estrogen-induced ductal morphogenesis
To test whether the increased levels of estrogen in pregnant mice were sufficient to induce ductal morphogenesis in SirT1ko/ko mice, a single estrogen pellet was implanted in the subscapular region of each virgin SirT1ko/ko mouse and the mammary tissues were analyzed 14 and 21 days after the implantation. By day 14, wild-type mice displayed ductal side branching reminiscent of the morphological changes during early pregnancy (Figure 4a, left panel). In contrast, SirT1ko/ko mice showed TEBs and ductal elongation, which are the characteristic features of ductal morphogenesis in pubertal wild-type mice. By day 21, ductal elongation and side branching were restored in SirT1ko/ko mice and the extent of ductal morphogenesis was indistinguishable between wild-type and SirT1ko/ko female mice (Figure 4a, right panel). These observations clearly indicated that exogenous estrogen alone is sufficient to rescue ductal morphogenesis in virgin SirT1ko/ko mice. Furthermore, the presence of transitional TEBs on day 14 and of ductal side branching on day 21 suggested that the characteristic features of mammary gland development at puberty and during early pregnancy could be coupled in response to increasing levels of estrogen. Increased levels of estrogen, either during pregnancy or through implantation, were therefore sufficient to stimulate the differentiation of epithelial progenitor cells and ductal morphogenesis in virgin SirT1ko/ko mice.
Concurrent mammary epithelial cell proliferation and differentiation
The finding of pregnancy-induced mammary gland development prompted us to investigate how mammary epithelial progenitor cells in pregnant SirT1ko/ko mice can differentiate so rapidly. Both BrdU labeling and the TUNEL (Terminal deoxynucleotidyl transferase mediated dUTP Nick End Labeling) assay were used to measure the cell proliferation and apoptosis in the mammary tissues, respectively. We found in SirT1ko/ko mammary tissues that about one-third of the cells in TEBs were BrdU-positive as compared with the number of BrdU-positive cells in the newly differentiated ductal or alveolar epithelial cells (Figure 5a(upper panel), 5b). This BrdU-positive finding differs from that in wild-type mammary tissues, in which less than 3% of ductal epithelial cells and about 25% of alveolar epithelial cells were BrdU-positive. The newly generated ductal epithelial cells in SirT1ko/ko mice formed elongated ducts and side branches, while pushing TEBs towards the edge of mammary fat pad (Figure 3a, lower panel). The newly generated ductal epithelial cells can consequently further differentiate into a few observed alveolar cells in which milk proteins were detected (Figure 2c). These observations suggested that the proliferation and differentiation of epithelial progenitor cells, which is programmed to take place at different times of a female's reproductive life, could be concurrent at a late stage of pregnancy in SirT1ko/ko mice.
Apoptosis is a part of the self-renewal and remodeling processes in normal mammary tissues [17,24,25]. Apoptotic epithelial cells can be readily identified in both ductal and alveolar epithelium of SirT1ko/ko mice on lactation day 1 (Figure 5a, lower panel). The levels of apoptotic cells were significantly higher than those in wild-type mice on lactation day 1, but seemed to be closer to that in wild-type mice in mid-pregnancy (e.g. day 13 of pregnancy) (Figure 5c). The rush of proliferation and differentiation during pregnancy-rescued ductal development in SirT1ko/ko mice manifested multiple features on lactation day 1 that could reassemble some characteristic features of wild-type mice in mid-pregnancy. Successive pregnancies, however, failed to improve the survival rate of either SirT1+/ko pups or SirT1ko/ko pups (three or four pregnancies in 3 months for each of three SirT1ko/ko females tested), indicating that the lactation defect persisted. A SirT1 deficiency therefore affected ductal morphogenesis in virgin mice, and lobuloalveolar proliferation in parous mice.
Deregulated negative feedback signals
The characterization of lactation failure has revealed two developmental defects in SirT1ko/ko female mice. The impeded ductal morphogenesis in virgin SirT1ko/ko mice bore some resemblance to the defective ductal morphogenesis seen in mice lacking GH, estrogen, or IGF-1, as well as in mice harboring mutations in the corresponding receptors, such as the GH receptor and ERα [17,24-31]. These previous studies have established the notion that GH and estrogen synergistically stimulate ductal morphogenesis, which is mediated by stromal cell-produced local IGF-1 in mammary tissues. Impeded ductal morphogenesis in virgin SirT1ko/ko mice is therefore probably due to a deficit in IGF-1 signaling.
In addition to this early defect, pregnancy-induced mammary gland development in SirT1ko/ko mice appeared to be arrested at the beginning of lobuloalveolar development, which is reminiscent of the phenotype in mice lacking IκB kinase alpha (IKKα) activity for NF-κB activation [37]. IKKα-deficient mice exhibit normal ductal morphogenesis at the onset of puberty and exhibit normal ductal side branching during early pregnancy. Owing to the absence of NF-κB-dependent cyclin D1 expression, however, IKKα-deficient mice display defective alveolar epithelial cell proliferation and fail to lactate [37]. While SirT1 negatively regulates the transcription activity of FoxOs and NF-κB in mammalian cells [10,11,20], SirT1ko/ko mice displayed the loss-of-function phenotypes. We hypothesized that SirT1 deficiency deregulates the expression of negative feedback signals and thereby desensitizes the cells to various types of stimulation. Indeed, among the many FoxOs and NF-κB-regulated genes, IGFBP-1 and IκBα encode negative feedback signals for IGF-1 signaling and NF-κB activation, respectively. We found that the basal levels of both IGFBP-1 and IκBα were increased in adipose tissues from virgin SirT1ko/ko mice and in ductal epithelial cells from SirT1ko/ko mice on lactation day 1 (Figure 3b). Meanwhile, the expression pattern of IGFBP-1 and IκBα in wild-type mammary tissues, shown in Figure 3b, was in agreement with the findings of others [16,38,39]. Namely, the expression of IGFBP-1 was downregulated during pregnancy, whereas NF-κB activity was increased at a late stage of pregnancy. The loss of SirT1 therefore increased the basal level of both IGFBP-1 and IκBα in multilineages of cells, and potentially increased thresholds for activating SirT1-modulated signaling pathways.
SirT1 deficiency did not affect the ability of mammary epithelial progenitor cells to differentiate into functional alveolar epithelial cells despite the lactation failure (Figure 2c). SirT1 deficiency may therefore alter the homeostasis of the signals for ductal morphogenesis and/or the cellular response to the signals. Increased expression of both IGFBP-1 and IκBα in adipose tissues and mammary epithelial cells could simply reflect a global deregulation of gene expression in SirT1ko/ko cells in which the activity of both FoxOs and NF-κB is increased. IGFBP-1 is a potent inhibitor of IGF-1 in vivo [40], suggesting that the elevated basal level of IGFBP-1 in adipose tissues in SirT1ko/ko mice could reduce the effect of IGF-1. Moreover, deregulated IGFBP-1 expression, but not deregulated IκBα expression, was reversed in response to increased levels of estrogen in the mammary fat pads after implantation (Figure 4b). The finding is relevant to the fact that the expression of IGFBP-1 is downregulated as the level of estrogen increases in pregnant wild-type mice (Figure 3b). Increased levels of estrogen therefore stimulate the production of stromal cell-derived local IGF-1, which overcomes the IGFBP-1 barrier in SirT1ko/ko mice and, ultimately, reverses the increased levels of IGFBP-1 in mammary tissues.
It was noted that increased levels of estrogen and local IGF-1 did not interfere with the expression of IκBα (Figure 4b). To determine whether deregulated IκBα expression in SirT1ko/ko cells can be reversed, we treated MEFs with TNFα and measured the kinetics of IκBα expression. Three independent lines of SirT1ko/ko MEFs were used, which displayed varying basal levels of increased IκBα expression. Following IκBα degradation induced by TNFα, the levels of newly synthesized IκBα in all three SirT1ko/ko MEFs were always higher than that of wild-type MEFs (Figure 6). This in vitro finding implies that SirT1 deficiency affects NF-κB signaling when the IκB kinase activation is normal.
Our current system might not be suitable for studying the effects of SirT1 deficiency on NF-κB activation in vivo. NF-κB signaling exhibits distinct biological responses because the expression of IκBα depends on NF-κB activation while the other isoforms of IκB (β and ε) are independent of NF-κB feedback [41]. For example, TNFα stimulation induces dampened oscillatory behavior after the first hour, whereas lipopolysacharide treatment leads to stable NF-κB activation [42,43]. The NF-κB signaling in the developing mammary gland is under the temporal control of RANK signaling and IKKα activation [37]. The finding of increased levels of newly synthesized IκBα within the first hour indicated that SirT1 deficiency could disrupt the temporal control of NF-κB signaling to all types of stimulation (Figure 6). It remains unknown, however, whether the reduced IGF-1 signaling would affect the efficacy of RANK signaling and IKKα activation in developing mammary glands. The fact that individual SirT1ko/ko mice manifested varying degrees of ductal morphogenesis (Figure 3a) makes it difficult to dissect the potential compound effect of SirT1 deficiency on both upstream IKKα activation and downstream NF-κB signaling. Other experimental systems, such as mammary epithelial cell-specific SirT1ko/ko female mice, may be used to address this issue.
Discussion
Our study of SirT1ko/ko mice has unveiled a regulatory mechanism by which SirT1 modulates the efficacy of estrogen-stimulated local IGF-1 signaling for ductal morphogenesis. SirT1 deficiency alters the homeostasis of gene expression and attenuates the efficacy of IGF-1 signaling. As a result, SirT1ko/ko mice manifest partial perinatal lethality and lactation failure phenotypes, implicating the role of evolutionarily conserved SirT1 in both postnatal survival and offspring survival of mammals.
SirT1 balances the homeostasis of IGF-1 and IGFBP-1 in vivo
SirT1 is an integral part of the IIS system that checks and balances the efficacy of insulin and IGF-1 signals. The IGF-1 signaling of the IIS system is required for survival after birth. As illustrated in Figure 7a, the function of SirT1 parallels the linear IGF-1/IGF-1 receptor/phosphoinositide-3'-OH kinase/Akt signaling pathway, both of which can negatively regulate the transcription activity of FoxOs. Mice harboring targeted mutations in either IGF-1 or the IGF-1 receptor gene exhibit perinatal lethality and growth retardation [44,45]. In the IGF-1 receptor-expressing cells, the binding of insulin and/or IGF-1 activates the phosphoinositide-3'-OH kinase and Akt kinase cascade for survival [8,9,13]. The phosphoinositide-3'-OH kinase/Akt signaling pathway is also highly conserved from worms to mammals, and functions to modulate energy metabolism and lifespan in lower organisms. The mammalian Akt kinase family has three members, Akt1–Akt3, which provide specificity and versatility. Mice lacking both the Akt1 and Akt2 genes display perinatal lethal and growth retardation phenotypes that are strikingly similar to that of either IGF-1 or IGF-1 receptor-deficient mice [46]. In this regard, SirT1ko/ko mice, as well as other strains of SirT1-deficienct mice, also exhibit perinatal lethal and growth retardation phenotypes (Figure 1c and Table 1) [22,23]. The consistent observation of perinatal lethal and growth retardation phenotypes from our study and studies with mice lacking IGF-1, the IGF-1 receptor, or Akt1/Akt2 suggests that upregulated FoxO activity unleashes the expression of downstream effector genes that lead to the birth-related stress. This birth-related stress may be related to the switch from a maternal level of IGF-1 to a neonatal level of IGF-1 because the surviving SirT1ko/ko mice can live into adulthood and can reproduce despite the growth retardation phenotype (Figure 1c). In this context, at least one of the FoxO-deficient mice strains, FoxO3 (Foxo3a/FKHRL1)-deficient mice, grow normally and develop functional mammary glands [47].
IGFBP-1 is one of six IGF-1 binding proteins, but is the only one that can be negatively regulated by insulin [48]. IGF-1 binding proteins inhibit the action of local IGF-1 in a spatial and temporal manner [40,49]. In mammalian cells, FoxO3 can directly bind to the promoter of the IGFBP-1 gene and activate the transcription [50-52]. SirT1 acts to negatively regulate the activity of nuclear FoxO3 and to decrease the expression of IGFBP-1 when the IGF-1 signal is low (Figure 7a). Increased levels of IGFBP-1 can diminish the action of IGF-1 signals in vivo. In that way, SirT1 balances the homeostasis of IGF-1 and IGFBP-1 for varying levels of IGF-1 signals.
The homeostasis of IGF-1 and IGFBP-1 clearly affects fertility, as shown by the infertility phenotype in both IGF-1-deficient mice and IGFBP-1 transgenic mice [28,40,49]. The serum level of estrogen in IGF-1-deficient female mice appears significantly reduced, whereas in ERα-deficient mice, which are also infertile, there is a threefold increase [53]. In contrast, SirT1ko/ko mice can be fertile despite the fact that the fertility was reduced as compared with that of wild-type females (Table 1). Although our preliminary assessment indicated that the serum level of estrogen in SirT1ko/ko female mice was not overtly altered, it remains possible that a subtle irregularity or an individual variation of the estrogen level or ERα expression might be attributed to the reduced fertility. An alternative explanation for reduced fertility is that SirT1 deficiency deregulates the activity of FoxO3 and manifests an opposite effect of FoxO3 deficiency. FoxO3-deficient mice exhibit early depletion of ovarian follicles, which is reminiscent of premature ovarian failure in women [47,54]. Premature ovarian failure manifests as early cessation of ovarian function and as premature menopause caused by genetic mutations, chemo/radiation therapy, or other unknown factors. It is possible that SirT1 deficiency counterbalances normal release of ovarian follicles and reduces the fertility in mice.
SirT1 modulates the efficacy of the IGF-1 signal for ductal morphogenesis
We hypothesize that, in virgin SirT1ko/ko mice, increased levels of IGFBP-1 in adipose tissues effectively diminish IGF-1 signals, including the estrogen–IGF-1 signaling, and halt the differentiation of mammary epithelial progenitor cells (Figure 7b). Several results of this study provided support for this hypothesis. First, the expression of IGFBP-1 is increased in mammary fat pads from virgin SirT1ko/ko mice, in which there is no evidence of any duct (Figures 2a and 3a). Second, both pregnancy and estrogen implantation can rescue impeded ductal morphogenesis in SirT1ko/ko mice (Figures 3a and 4a), demonstrating that increased levels of estrogen can enhance the production of local IGF-1. Moreover, the varying degrees of rescue among individual mice, as well as individual mammary fat pads of the same mouse, also indicated that local IGF-1 must be halted in SirT1ko/ko mice (Figure 3a, and data not shown). Finally, increased levels of estrogen specifically activate the IGF-1 signaling in mammary tissues, as shown by the reversal of IGFBP-1 expression in SirT1ko/ko mice after estrogen implantation (Figure 4b). These results indicate that a gradient of IGF-1 signals, including both circulating and local IGF-1, may regulate mammary gland development at embryonic, postnatal, and reproductive stages, and that increased expression of IGFBP-1 decreases the efficacy of IGF-1 signals.
Mammary development may undergo estrogen/IGF-1-independent phases and estrogen/IGF-1-dependent phases, which encompasses a late stage of embryonic development to the onset of puberty and involves maternal IGF-1, circulating IGF-1, and estrogen-stimulated local IGF-1. By embryonic day 16.5, mammary mesenchymes are connected to primitive fat pads and begin to proliferate, which results in rudimentary ducts [17,25,55,56]. We propose that this process marks the transition from estrogen/IGF-1-independent mammary stem cells to IGF-1-dependent epithelial progenitor cells, and that maternal IGF-1 is sufficient to stimulate the differentiation of epithelial progenitor cells to estrogen-independent ductal epithelial cells but not estrogen-dependent epithelial cells (Figure 7b). This explains why the development of rudimentary ducts is normal in either IGF-1 or ERα knockout mice and that TEBs form and regress immediately before and after birth, respectively [17,26,27]. During the prepubertal period, the rudimentary ducts grow isometrically under the influence of circulating IGF-1. Like maternal IGF-1, circulating IGF-1 is not sufficient to induce the differentiation of estrogen-independent ductal epithelial cells into estrogen-dependent epithelial cells, which will express ERα and become estrogen dependent (Figure 7b). At the onset of puberty, ovarian estrogen, in synergy with pituitary GH, causes a surge in the production of stromal cell-derived local IGF-1 and stimulates the differentiation of estrogen-independent ductal epithelial cells to estrogen-dependent epithelial cells, which begins postnatal development of mammary gland (Figure 7b). Impeded ductal morphogenesis results in virgin SirT1ko/ko mice because local IGF-1 is apparently not sufficient to overcome increased IGFBP-1 in mammary adipose tissues. SirT1 therefore positively regulates the efficacy of the estrogen–IGF-1 signaling for ductal epithelial cell proliferation and differentiation.
The potential effect of modulating SirT1 activity
Breast cancer results from the accumulation of inherited and/or somatic mutations. Lifetime exposure to estrogen is a major risk factor for breast cancer, and a number of lifestyle factors, such as diet, body fat, alcohol consumption, exercise, parity, and hormone replacement therapy, may influence a woman's exposure to estrogen. The compound personal risk to a woman is difficult to assess because the molecular link between these factors and the effect of estrogen is poorly understood. The result of this study demonstrated that SirT1 positively modulates the efficacy of the estrogen–IGF-1 signal. This suggests that if SirT1 serves as a lifetime sensor to dietary restriction and acute withdrawal of nutrients [14,57], its activity could ultimately influence the risk of breast cancer. Interestingly, an epidemiological study of Dutch famine in 1945 indicated that an exposure to famine at prepubertal age increased the risk of breast cancer, most clearly seen in women who never gave birth [58]. Moreover, the serum levels of estrogen and IGF-1 were increased among those exposed to the famine, which suggests that an epigenetic adaptation phenomenon may take place. Assessing the potential correlation between the enzymatic activity of SirT1 and the risk of breast cancer is therefore of great interest in order to identify feasible targets for chemoprevention against breast cancer.
Several dietary polyphenols as well as small molecules have been identified as either agonists or antagonists of SirT1 [59,60]. The pharmacological effects of these compounds in humans remain elusive given that SirT1 targets multiple transcription factors and exerts pleiotropic effects. Nonetheless, our study of SirT1ko/ko mice demonstrated the potential utility of SirT1 antagonists. Specifically, reducing SirT1 activity alters the homeostasis of cellular responses, including deregulation of the expression of IGFBP-1 and IκBα, which can attenuate the stimulation from the estrogen–IGF-1 signaling and potentially desensitize mammary epithelial cells to NF-κB activation. SirT1 is therefore a candidate target for chemoprevention against breast cancer.
Conclusion
The study of mammary gland development is an excellent model system for unraveling how SirT1 modulates the efficacy of the estrogen–IGF-1 signaling and regulates the timing of ductal morphogenesis. These new mechanistic insights may also aid in understanding the role of SirT1 in breast cancer as well as in other organs in which local IGF-1 plays an important role.
Abbreviations
Atm = ataxia telangiectasia; BrdU = bromodeoxyurdine; CMV = cytomegalovirus; ERα = estrogen receptor alpha; FoxO = forkhead box 'other' protein; GH = growth hormone; IGF-1 = insulin-like growth factor-1; IGFBP-1 = insulin-like growth factor-1 binding protein-1; H & E = hemotoxylin and eosin; IIS = insulin/insulin-like growth factor-1 signaling; IκBα = inhibitors of NF-κB alpha subunit; IKKα = IκB kinase alpha; MEF = murine embryonic fibroblast; NF = nuclear factor; PCR = polymerase chain reaction; RANK = receptor activator of NF-κB; Sir = silencing information regulator; TEB = terminal end bud; TNF = tumor necrosis factor.
Competing interests
The authors declare that they have no competing interests.
Authors' contributions
HL characterized the phenotypes of the SirT1 mutant mice, participated in the design of the study, and helped to draft the manuscript. GKR and NL helped to characterize the phenotypes. CW generated the SirT1 mutant embryonic stem cells and mice. BPR helped analyze the histology data. YG conceived of the study, participated in its design and coordination, and drafted the manuscript. All authors read and approved the final manuscript.
Supplementary Material
Additional file 1
A pdf file containing the materials and methods for the generation of SirT1ko/ko mice. This supporting file documented the details of materials and methods used to generate both SirT1co/co mice and SirT1ko/ko mice.
Click here for file
Acknowledgements
The study is supported by grants from the American Cancer Society (RSG-04-019-01-CNE) and the Nathan Shock Center of Excellence in the Biology of Aging at the University of Washington (to YG). The authors thank Jennifer Blasi, Thomas Gernon, Janet Leath, Jennifer Lee, Bryce Sopher, Loraine Warner, and Heather-Marie Wilson for their contributions in the early phases of the study, and Dr Fred Alt, Dr Mary Helen Barcellos-Hoff, Dr Mark Groudine, Dr Warren Ladiges, Dr George Martin, Dr Peter Rabinovitch, and Dr Jeffrey Schwartz for their support and advice.
Figures and Tables
Figure 1
Generation and characterization of SirT1co/co and SirT1ko/ko mice. (a) Mouse SirT1 wild-type allele (SirT1+), conditional targeted allele (SirT1co), and knockout allele (SirT1ko). B, BamHI. (b) Western blot analysis of SirT1 expression in wild-type (+/+), conditional targeted (co/co), heterozygote (+/ko), and knockout (ko/ko) murine embryonic fibroblasts (MEFs) and mammary tissues (MG). (c) Growth retardation in surviving SirT1ko/ko mice (open bar) using sibling mice or age-matched mice of control genotypes (filled bar). (d) The serum levels of insulin-like growth factor-1 (IGF-1) in wild-type (+/+), SirT1+/ko (+/ko), and SirT1ko/ko (ko/ko) mice. (e) The survival curves of wild-type (WT), SirT1co/co, SirT1ko/ko, and Ku70-/- embryonic stem cells after ionizing radiation. (f) The survival curve of wild-type (WT), SirT1ko/ko, Ku70-/-, and Atm-/- embryonic stem cells after treatment with hydrogen peroxide.
Figure 2
Lactation failure in SirT1ko/ko mice. (a) Whole mount analysis of the mammary tissues from virgin wild-type (+/+) and SirT1ko/ko mice (ko/ko). Scale bar = 2 mm. (b) Histological analysis of the mammary tissues harvested from wild-type (+/+) and SirT1ko/ko female mice on either day 13 of pregnancy (P13) or lactation day 1 (L1). Scale bars = 100 μm. (c) Immunofluorescence staining of milk production in the mammary tissues from wild-type (+/+) and SirT1ko/ko mice on L1. Red and smear stains, mouse milk proteins; blue dots, nuclear staining of adipocytes, epithelial cells, and other cells in mammary tissues. Scale bars = 100 μm.
Figure 3
Pregnancy-induced ductal morphogenesis. (a) Whole mount analysis of mammary gland development. Upper panel: wild-type (+/+) mice show terminal end buds (TEBs) at the onset of puberty, elongated ducts (virgin mice), site branching during pregnancy (P13), and lobuloalveolar structures for milk production on lactation day 1 (L1). Lower panel: virgin SirT1ko/ko (ko/ko) mice show impeded ductal morphogenesis. Pregnancy-induced ductal morphogenesis manifests transitional TEBs, ductal elongation, and side branching with variety in three SirT1ko/ko mice on L1. All scale bars = 200 μm. (b) Western blot analysis of both insulin-like growth factor-1 binding protein-1 (IGFBP-1) and IκBα in mammary tissues of male mice, virgin mice, and L1 female mice of wild type (+/+) and SirT1ko/ko (ko/ko). Actin is used as a loading control. Lower panel scores the estimated density of indicated lineages of cells in the protein extracts, which are based on the morphological analyses in (a).
Figure 4
Estrogen implantation stimulates ductal elongation and site branching. (a) Whole mount analysis of the mammary tissues from the virgin wild-type (+/+) and SirT1ko/ko mice implanted with estrogen pellets (+E2) on day 14 and day 21. Scale bars = 200 μm. (b) Western blot analysis of the expression of insulin-like growth factor-1 binding protein-1 (IGFBP-1) and IκBα, with actin as a loading control, in mammary tissues from the virgin wild-type (+/+) and SirT1ko/ko mice implanted with estrogen pellets on day 21.
Figure 5
Mammary epithelial cell proliferation and apoptosis in SirT1ko/ko mice on lactation day 1. (a) Upper panel: bromodeoxyurdine (BrdU) staining of the mammary sections from SirT1ko/ko (ko/ko) and wild-type (+/+) mice on day 13 of pregnancy (P13) and lactation day 1 (L1). From left to right, three terminal end buds (TEBs) at a scale of 100 μm, a single TEB, ducts, and alveolus of SirT1ko/ko mice at a scale of 20 μm, and ducts and alveolus of wild-type mice on P13 and L1 at a scale of 20 μm. Lower panel: TUNEL (Terminal deoxynucleotidyl transferase mediated dUTP Nick End Labeling) assay for apoptotic cells using sections neighbored to the sections for BrdU staining. (b) The quantitative analysis of BrdU-positive epithelial cells in TEBs, ducts, and alveoli of SirT1ko/ko mice on L1 (open bar) and wild-type mice on P13 (grey bar) and L1 (solid bar). (c) The quantitative analysis of TUNEL-positive cells in neighboring sections.
Figure 6
SirT1 deficiency derails NF-κB signaling in response to TNFα stimulation. (a) Western blot analysis of IκBα expression in SirT1ko/ko (ko/ko) and wild-type (+/+) murine embryonic fibroblasts (MEFs) within the first hour after TNFα stimulation. Actin is used as a loading control. (b) The relative levels of newly synthesized IκBα in three independent SirT1ko/ko MEF lines (red circles, blue and green triangles) and wild-type MEFs after TNFα stimulation. For each line of MEFs, the unit of IκBα protein at each time point is relative to the unit 0 minutes after normalization with actin.
Figure 7
SirT1 modulates estrogen–insulin-like growth factor-1 signaling for ductal morphogenesis: a model. (a) The insulin/insulin-like growth factor-1 (IGF-1) signaling system: both SirT1 and Akt kinases can negatively regulate the transcription activity of forkhead box 'other' protein (FoxO) proteins. SirT1 deficiency deregulates the expression of insulin-like growth factor-1 binding protein-1 (IGFBP-1), which may exert autocrine and/or paracrine effects to inhibit IGF-1. (b) Mammary epithelial precursor cells (EPC) express IGF-1 receptor (IGF-1R) and can differentiate into estrogen receptor (ER)-negative ductal epithelial cells (DEC-I) in response to maternal, circulating, or estrogen-stimulated, stromal cell (S)-derived local IGF-1. At the onset of puberty, ovarian estrogen, in synergy with growth hormone (GH), enhances the production of local IGF-1 and stimulates the differentiation of DEC-I to ER-positive ductal epithelial cells (DEC-II). SirT1 deficiency deregulates the expression of IGFBP-1 in adipose tissues (a), however, which attenuates the efficacy of the IGF-1 signaling and causes impeded ductal morphogenesis in virgin SirT1ko/ko mice. Either pregnancy or exogenous estrogen can overcome the impeded ductal morphogenesis in virgin SirT1ko/ko mice and can stimulate the differentiation of EPC.
Table 1
Perinatal lethality, fertility, and lactation defect in SirT1ko/ko mice
More than 10 pairs of SirT1+/ko male mice and SirT1+/ko female mice were used for breeding. Data in parentheses indicate the number of pups of this genotype found dead within 3 days after their birth. aTotal number of surviving pups at weaning (postnatal day 21). | [
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17206865 | A Role for Alström Syndrome Protein, Alms1, in Kidney Ciliogenesis and Cellular Quiescence
Abstract
Premature truncation alleles in the ALMS1 gene are a frequent cause of human Alström syndrome. Alström syndrome is a rare disorder characterized by early obesity and sensory impairment, symptoms shared with other genetic diseases affecting proteins of the primary cilium. ALMS1 localizes to centrosomes and ciliary basal bodies, but truncation mutations in Alms1/ALMS1 do not preclude formation of cilia. Here, we show that in vitro knockdown of Alms1 in mice causes stunted cilia on kidney epithelial cells and prevents these cells from increasing calcium influx in response to mechanical stimuli. The stunted-cilium phenotype can be rescued with a 5′ fragment of the Alms1 cDNA, which resembles disease-associated alleles. In a mouse model of Alström syndrome, Alms1 protein can be stably expressed from the mutant allele and is required for cilia formation in primary cells. Aged mice developed specific loss of cilia from the kidney proximal tubules, which is associated with foci of apoptosis or proliferation. As renal failure is a common cause of mortality in Alström syndrome patients, we conclude that this disease should be considered as a further example of the class of renal ciliopathies: wild-type or mutant alleles of the Alström syndrome gene can support normal kidney ciliogenesis in vitro and in vivo, but mutant alleles are associated with age-dependent loss of kidney primary cilia.
Author Summary
Alström syndrome is a rare genetic disorder caused by mutations in the ALMS1 gene. The disease is characterized by blindness, deafness, and metabolic disorders. These symptoms are reminiscent of diseases affecting the primary cilium, a cellular appendage with a role in sensing changes to the extracellular environment. In addition, kidney failure is a frequent cause of death in Alström syndrome patients, and recent studies have suggested a causal relationship between defects in primary cilia and cystic kidney diseases. In this paper, we show that ALMS1 protein is required to form cilia in kidney cells. Mutant alleles of the gene that are similar to those seen in the human disease are able to support cilia formation in cell culture and in animals. However, a defect in the function of the disease alleles is uncovered in older mice: cilia are lost from the kidney cells, and this is associated with an increase in cellular proliferation and cell death. The data are consistent with a requirement for ALMS1 in ciliogenesis and suggest inclusion of Alström syndrome among the growing class of cilia-related pathologies.
Introduction
Alström syndrome is a rare autosomal recessive disorder characterized by early onset obesity, type 2 diabetes mellitus, retinal degeneration, and hearing impairment. Other aspects of the disease include cardiomyopathy, liver dysfunction, kidney dysfunction, and a delay in puberty. Renal function declines with age, and end-stage renal disease is a common cause of death in Alström syndrome patients [1]. Additionally, enlarged kidneys have been reported in a previously reported mouse strain with a mutation in Alms1 [2].
The primary cilium is an antenna-like organelle, surrounded by a membrane contiguous with the plasma membrane [3,4]. Typically, cilia extend several micrometers from the apical face of the cell, grounded to the cellular microtubule complex through the basal body. Cilia are conserved through several eukaryotic phyla, including Caenorhabditis elegans, Chlamydomonas, and vertebrates. Immotile primary cilia contain a scaffold of nine microtubule doublets running the length of the axoneme (9 + 0), whereas motile cilia contain an additional central microtubule pair (9 + 2 arrangement). Increasing data demonstrate roles for the primary cilium in sensory functions. These include mechanosensation of lumenal flow in kidney tubules and transduction of extracellular signaling through the hedgehog, Wnt, and platelet-derived growth factor receptor pathways [3,4].
Four lines of evidence suggest the hypothesis that renal failure in Alström patients is secondary to a defect in primary cilia in the kidney. First, mutations in genes that are implicated in the function of primary cilia are associated with kidney diseases. Polycystic kidney disease (PKD) is characterized by progressive development of fluid-filled cysts, ultimately leading to end-stage renal failure [5]. Both autosomal dominant PKD (ADPDK) proteins (Polycystin-1 and −2) are localized to primary cilia and are necessary for cilia-mediated signaling in response to a fluid-flow stimulus [6]. Autosomal recessive PKD (ARPKD) protein (fibrocystin) and nephronophthisis disease proteins, nephrocystin and inversin, are involved in ciliary protein transport [7,8]. Additionally, mouse strains with genetic lesions in ciliary proteins lead to cystic kidney disease [9,10]. Second, the spectrum of phenotypes seen in Alström patients is similar to Bardet-Biedl patients [11], suggesting that mutations in ALMS1 might cause disease through a similar mechanism to BBS mutations. It is now well documented that several of the 11 genes mutated in Bardet-Biedl syndrome (BBS) have roles in the function of primary cilia [12–19]. Third, ALMS1 is localized to centrosome and ciliary basal bodies in vitro [20,21], consistent with a role in the structure of the basal body or in the transport of proteins between the cytoplasm and the ciliary axoneme. Fourth, in vivo phenotypes of Alms1 mutant mice include a lack of sperm flagella, a modified ciliary structure, as well as defective rhodopsin transport through the connecting cilia of photoreceptor cells [2,22].
These data point to a role of ALMS1 in the function of primary cilia, but no defects in cilia formation were observed in human dermal fibroblasts from an Alström syndrome patient [21] or in the kidney collecting duct epithelial cells of a gene-trap Alms1 mutant mouse strain [2]. A possible resolution of this discrepancy comes from the genetic analysis of mutations in ALMS1 that have been found in patients. In all cases, there was at least one allele of ALMS1 that could encode an N-terminal fragment of the ALMS1 protein before the presence of a premature stop codon. Moreover, the alleles in two reported mice models of Alström syndrome would also be predicted to encode the N-terminus of Alms1 [2,22]. Thus, it is possible that ALMS1 is required for cilia formation, and that the disease-associated alleles are able to provide at least part of this activity through expression of the N-terminus of the protein. In this study, we test whether ALMS1 has a role in cilia formation or function and provide evidence that renal failure in Alström syndrome patients might be associated with ciliary dysfunction.
Results
Depletion of Alms1 mRNA and Protein by Short Interfering RNA Causes Defective Ciliogenesis in Kidney Cells
To determine whether Alms1 was necessary for cilium formation and/or function, we used an in vitro model of kidney cell ciliogenesis and signaling. Mouse inner medullary collecting duct (mIMCD3) cells form cilia 5 d after confluency [23]. The cilia can be visualized as long protrusions from the cell surface using an antibody raised against acetylated tubulin (Figure 1A). We tested several short interfering RNA (siRNA) molecules designed against the mouse Alms1 sequence for their effects on formation of cilia in this model. Cilia were formed normally in the presence of a negative control siRNA. However, transfection with two siRNA sequences against Alms1, Alms1a and Alms1b, led to a markedly different phenotype. The acetylated tubulin staining on each cell manifested as a single ball of fluorescence, and very few cells showed the elongated staining typical of ciliary axoneme (Figure 1A). Both of these siRNA species also caused a decrease in the Alms1 mRNA level (Figure 1B). In contrast, two additional siRNAs (Alms1c and Alms1d) did not affect the pattern of acetylated tubulin staining in the ciliogenesis assay, nor did they affect the Alms1 mRNA level (Figure 1A and 1B). To demonstrate that the active siRNA species also decreased levels of Alms1 protein, we raised an antiserum in rabbits against the predicted N-terminal 13 amino acids of the open reading frame of Alms1. Using this antiserum, we detected positive staining at the base of cilia, consistent with previous reports on the subcellular localization of ALMS1 [21]. The antibody signal was reduced below detection in the presence of the active siRNA molecules (Figure 1C).
Loss of Alms1 Does Not Affect Transcriptional Regulation of Ciliary Genes but Does Disrupt Ciliary Mechanosensation
Transcriptional profiling in Chlamydomonas and in C. elegans has identified a set of genes that are regulated during ciliogenesis [24,25]. As an initial characterization of the role of Alms1 in the mIMCD3 in vitro model, we asked whether Alms1 protein was required for the transcriptional response associated with ciliogenesis. A time course of mIMCD3 ciliogenesis showed a steady rise in the level of Alms1 mRNA after confluency. This was paralleled by an increase in cilia length (Figure 2A and 2B). We chose two other cilia-related genes that have increased mRNA levels during ciliogenesis, Bbs4 and Ttc10. Bbs4 (BBS gene 4) encodes a protein localized to the basal body and centrosome that is required for targeting cargo to the pericentriolar region [13]; Ttc10 (IFT88/polaris) is required for transport of cargo from basal body to the distal cilium [26]. Bbs4 and Ttc10 mRNA were upregulated even when ciliogenesis was disrupted with Alms1a siRNA (Figure 2B). Moreover, microarray analysis showed that knockdown of Alms1 by siRNA had no effect on the general transcription program associated with confluency and ciliogenesis in the mIMCD3 cells (Figure 2C and Table S1).
We then used this model to ask whether the stunted cilia formed in limiting amounts of Alms1 were functionally competent. Cilia on kidney epithelial cells are able to induce an intracellular calcium signal in response to fluid flow over the cell surface. Defects in this response are thought to underlie PKD [27,28]. Full-length cilia formed in the presence of a control siRNA were able to respond to flow as expected (Figure 2D). However, the stunted cilia formed in the presence of the Alms1a siRNA were unable to induce calcium flux after a flow stimulus.
Taken together, these in vitro results suggest a critical role of Alms1 in cilia biogenesis of kidney epithelial cells, though without disruption of the transcriptional program that accompanies this process. The abnormal cilia formed after knockdown of Alms1 are characterized by a focal concentration of acetylated tubulin at the cell membrane, and these stunted cilia are unable to respond to fluid flow by calcium flux.
A 5′ Fragment of Alms1 Is Sufficient to Rescue Cilia Formation In Vitro after Knockdown of Endogenous Alms1 and Can Support Normal Ciliogenesis In Vivo
Reported alleles of ALMS1 in human patients, as well as in two reported mouse models of the disease, usually encode premature termination codons in exons 8, 10, and 16 [2,22,29,30]. In these alleles, the 5′ terminus of the gene encodes an intact open reading frame that might give rise to a partially functional protein. Therefore, we tested whether a truncated 5′ cDNA was able to rescue cilia formation after Alms1 knockdown. We transfected a mouse Alms1 cDNA encoding the N-terminal 1,282 amino acids, equivalent to exons 1–8, into mIMCD3 cells in the presence of the Alms1a siRNA. The Alms1a siRNA sequence matches the Alms1 gene 3′ of the cDNA fragment and so was predicted to affect only the levels of the endogenous transcript and not expression of the cDNA encoding the N-terminus of Alms1. As expected, transfection with Alms1a siRNA caused focal acetylated tubulin staining. However, cotransfection of the Alms1 cDNA with Alms1a siRNA was fully capable of rescuing the cilia phenotype (Figure 3A). To monitor the effects of cDNA transfection on knockdown of the endogenous mRNA level, we used a Taqman assay recognizing the boundary of exons 12 and 13 of the Alms1 mRNA. This assay will detect the endogenous full-length transcript but not the transfected cDNA and shows that cotransfection of the cDNA did not affect knockdown of the endogenous transcript. Conversely, using a Taqman assay recognizing the 5′ of Alms1 mRNA that detects both the endogenous and cDNA-encoded transcript, we showed that cotransfection with the siRNA Alms1a did not affect over-expression of the Alms1 5′ cDNA (Figure 3B).
To determine whether a premature truncation allele of the Alms1 gene can support normal ciliogenesis at endogenous expression levels, we used primary cells from a mouse strain with an ethyl nitroso urea–induced, premature truncation mutation in exon 10 of the Alms1 gene (Figure S1). The Alms1 gene in this mutant strain is predicted to encode the N-terminal 2,131 amino acids of the mouse Alms1 protein, and we therefore named the allele Alms1L2131X/L2131X. Similar expression of Alms1 mRNA was observed across eight tissues in wild-type and homozygous Alms1L2131X/L2131X mice (Figure 3C). In particular, levels of Alms1 mRNA in the kidney of mutant Alms1L2131X/L2131X mice are at least as high as in the wild-type control mice. In addition, we detected mutant Alms1 protein localized at the base of cilia of Alms1L2131X/L2131X primary kidney cells (Figure 3D). Thus, despite the presence of a premature stop codon allele in the Alms1 gene of these mice, both mRNA and protein were readily detectable. By isolating primary fibroblasts and kidney cells from Alms1L2131X/L2131X mice and wild-type controls, we found that cells from the Alms1L2131X/L2131X strain showed normal primary cilia when compared to wild-type cells (Figure 3E), which is consistent with previous reports [2,21]. We then tested whether expression of the mutant Alms1 transcript was required for normal cilia formation in cells derived from the Alms1L2131X/L2131X mouse strain. Knockdown of mutant Alms1 mRNA using a 5′ specific siRNA in the Alms1L2131X/L2131X embryonic fibroblasts inhibited ciliogenesis in these cells (Figure 3F), as was seen for wild-type cells (unpublished data).
Age-Dependent Loss of Primary Cilia and Homeostasis in Alms1 Mutant Mice
We characterized the phenotype of Alms1L2131X/L2131X mouse strain with reference to the metabolic and sensory defects reported in Alström syndrome patients to determine whether the pathology of the mouse strain resembled that of human patients. Homozygous mutant mice increased in weight faster than wild-type controls between 7 and 10 wk of age, and body composition analysis showed that this increase was almost entirely explained by an increase in fat mass (Figure 4A). Histological analysis showed hypertrophy of white and brown fat adipocytes and steatosis of the liver in obese mutant mice (Figure 4B). Although one older male mutant mouse in this cohort of 20 became diabetic, most mice had normal glucose levels with hyperinsulinemia. Other serum abnormalities include elevated leptin, triglycerides, total cholesterol, and HDL cholesterol (Figure S2). Finally, we noted that the Alms1L2131X/L2131X strain had defective sperm formation in the testes and defective rhodopsin transport in the retina (Figure 4C and 4D). We conclude from these data that the Alms1L2131X/L2131X mouse strain is a close genetic model of human Alström syndrome [1], and we proceeded to analyze whether there were any defects in the kidney cilia that might relate to renal failure in human patients.
The kidneys of 6-mo-old Alms1L2131X/L2131X mice contained multiple dilated tubules in the cortex (Figure 5A). Consistent with a previous report [2], primary cilia appeared normal in collecting ducts of the renal medulla. However, we noticed that some cortex tubules appeared to be almost completely denuded of cilia in aged Alms1L2131X/L2131X mutants (Figure 5A and 5B). To further characterize these tubules, we stained the kidney sections with Lotus tetragonolobus agglutinin (LTA) as a proximal tubular marker, and with aquaporin-2 antibody as a marker of collecting ducts (Figure 5C). In both the wild-type and Alms1L2131X/L2131X mutant animals, the aquaporin-2-labeled collecting duct cells have clear primary cilia as expected. LTA-labeled proximal tubule cells in the wild-type animals were also ciliated. In contrast, most of the LTA-labeled proximal tubules of Alms1L2131X/L2131X mutants were not ciliated.
Human PKD is thought to originate from renal tubule dilatation, secondary to abnormalities in cellular proliferation and apoptosis. We examined kidney sections from wild-type and mutant mice with a marker for proliferation, Ki67, and by TUNEL staining for apoptotic cells (Figure 5D). Sparse proliferation was seen in wild-type kidney sections. Higher levels of proliferation were detected in the cortex of kidneys from Alms1L2131X/L2131X mice, and this was focused in patches, perhaps representing convolutions of the same tubule coming into the plane of sectioning. In most of the cross sections of dilated tubules, several (20%–50%) of the epithelial cells were Ki67-positive, and these cells lacked cilia. We also noted a dramatic increase in kidney epithelial cell apoptosis in the mutant mice. As with Ki67, it was striking that TUNEL-positive cells had a nonuniform distribution and seemed to be clustered within particular tubules, whereas other tubules appeared identical to wild-type controls. These changes appeared to compromise kidney function, as urinanalysis revealed a mild proteinurea in adult Alms1L2131X/L2131X mice (Figure 5E). None of the kidney phenotypes described above (dilated tubules, loss of cilia, proliferation and apoptosis of epithelial cells, or proteinurea) was seen in 2-mo-old mice (Figure S3 and unpublished data).
Discussion
ALMS1 is among the largest disease genes identified today in the human genome. However, amino acid sequence analysis identifies only a leucine zipper near the N-terminus, allowing limited inferences to be made about the function of this protein. Localization of ALMS1 to the centrosome and ciliary basal body has been determined using a green fluorescent protein fusion to its C-terminus, and with an N-terminal antibody, respectively [20,21]. These data are consistent with a role for ALMS1 in the function of cilia, which is supported by the overlapping clinical phenotypes between Alström syndrome and other ciliopathies, especially BBS. Additionally, mouse models of BBS and Alström syndrome lack sperm flagella and a modified cilium; and show aberrant transport of rhodopsin, pointing to defective function of the connecting cilium in photoreceptor cells.
In this paper, we investigated whether Alms1 protein is required for the formation of cilia and, if so, whether cilia can be formed in human or mouse cells with a mutated ALMS1/Alms1 gene. We used an in vitro model of kidney ciliogenesis to demonstrate that knockdown of Alms1 mRNA caused a striking alteration in the morphology of cilia: knockdown of Alms1 caused stunted or focal staining of acetylated tubulin. The specificity of the siRNA reagents was confirmed by two pieces of evidence. First, two siRNAs that were active in decreasing mRNA expression were also active in decreasing formation of elongated cilia. Conversely, two additional Alms1 siRNAs, as well as a negative control siRNA, did not decrease Alms1 mRNA nor did they affect the ciliary phenotype. Second, the ciliary phenotype that was induced by Alms1 siRNA knockdown could be rescued by cotransfection of a 5′ fragment of Alms1 that was not targeted by the active siRNA. Together these results rule out the possibility that the cilia phenotype caused by siRNA knockdown was an off-target effect, and point to a role of Alms1 in the maintenance or biogenesis of cilia.
Interestingly, previous work had shown that cilia were formed in cells with mutant ALMS1/Alms1. To reconcile those findings with our siRNA knockdown data, we posited a functional role for the truncated Alms1 protein encoded by the mutant alleles. Alms1 mRNA in our mutant mouse strain was present at levels similar to those in wild-type mice, despite the presence of a premature stop codon. We also found that the mutant Alms1 protein was detectable and localized to ciliary basal bodies in cells from these mice, just as was observed for wild-type Alms1. Additionally, in unpublished data, immunohistochemistry with the antibody raised against the N-terminus of Alms1 showed prominent staining in the mutant pancreatic islets, confirming that protein could be stably expressed from the mutant allele. To test whether there was a function for the mutant protein, we showed that knockdown of the mutant Alms1 mRNA inhibited ciliogenesis in primary cells from these mice. We also showed that the ciliary phenotype induced by knockdown of Alms1 could be rescued using a cDNA encoding only the N-terminal 1,282 amino acids of Alms1, confirming that the N-terminus of Alms1 is sufficient to support cilia formation.
A spectrum of nonsense and frameshift mutations cause Alström syndrome, and all have different effects. However, no genotype/phenotype correlation has been observed among human patients [29–31]. Furthermore, two mouse models of Alström syndrome reported to date, as well as a third model reported in this manuscript, contained alleles that were predicted to encode the N-terminus of the protein, and the reported phenotypes are very consistent. Thus, we suggest that truncation of ALMS1/ Alms1, secondary to mutations in (predominantly) exons 8, 10, and 16, leads to similar phenotypes in human and mouse. In particular, we provide evidence for residual function of the disease-associated alleles. This residual function of mutant Alms1 alleles might explain a lack of the more severe developmental phenotypes described in animal models with mutations of intraflagellar transport (IFT) or kinesin motor proteins [26,32,33].
There is growing support for the “antenna” role of primary cilium as a key participant in sensing environmental stimuli, transduction of intracellular signaling, and regulation of morphogenetic pathways. Studies of BBS patients and BBS knockout animal models have revealed the role of primary cilia in sensing of light and olfaction [14,16,17,19]. The kidney primary cilium, extending from the tubular epithelium into the lumen, has been implicated in mechanosensation: genetic or chemical disruption of cilia inhibited intracellular calcium influx in response to laminar flow [34,35]. Recent studies have suggested that cilia constitute an essential platform where signaling processes are initiated. The Hedgehog receptor Smoothened and the platelet-derived growth factor receptor alpha localize to the cilium and, in both cases, ciliary localization is required for signaling transduction [36,37]. We show here that Alms1 is necessary for formation of kidney epithelial cilia, which in turn are known to be essential for mechanosensory signaling.
Well-characterized genetic disorders of primary cilia include the family of PKD, inherited either as ADPKD or ARPKD traits. While ARPKD is congenital and often causes fetal or neonatal death, the onset of the ADPKD is mainly in adulthood with age-dependent progression [5]. Animal models of PKD have been developed by genetic mutation of PKD genes [38–41], loss of kidney cilia by inhibition of IFT components [10,42], or constitutive activation of the Wnt pathway [43].
Recent data have started to build a connection between these seemingly unrelated genetic perturbations in PKD, IFT, and Wnt genes. Localization of PKD proteins has been observed on, or at the base of, cilia and cilia are dependent on IFT for protein transport to and from the cytoplasm [44]. Both PKD1 and PKD2 function on cilia to sense lumenal flow and control cell proliferation. Specifically, fluid flow over the cilia increases expression of inversin, which in turn targets cytoplasmic Dsh, inhibiting the canonical Wnt pathway [45]. It has been proposed that urine flow terminates the canonical Wnt pathway in favor of the noncanonical Wnt pathway. The noncanonical Wnt pathway might then maintain planar cell polarity and restrict cell division in a direction parallel to the long axis of the tubule. Moreover, BBS proteins have been shown to directly interact with proteins which regulate planar cell polarity [46], as well as in ciliary protein transport [47], suggesting a model in which defects in cilia disrupt planar cell polarity signaling and lead to disorientated cell division and cyst formation.
Existing mouse PKD animal models are characterized by the formation of early-onset cysts in the collecting tubules, reminiscent of human ARPKD. In contrast, the Alms1 mutant mouse model presented here shows cilia loss in the proximal tubules. Further, adult mice progress to a breakdown in maintenance of kidney epithelial cellular quiescence, with a dramatic increase in apoptotic and proliferating cells restricted to those tubules that have lost cilia. This is accompanied by tubular dilatation and mild proteinurea in older mutant mice. All these phenotypes are similar to clinical features of ADPKD, making the Alms1L2131X/L2131X mouse strain an interesting animal model to study the relation of cilia loss, altered signaling, and cellular proliferation in the progression of cystic kidney disease in the proximal tubules.
Initiation of ADPKD is suggested to be dependent on a second somatic mutation in the wild-type allele of PKD1 or PKD2. Evidence for this hypothesis is controversial as somatic mutagenesis rates in nontransformed cells are likely to be too low to explain the high prevalence of bilateral disease in heterozygous carriers [48,49] and a reduced expression of PKD1 is sufficient to initiate cystogenesis [41]. In this recessive model of ciliary dysfunction in the kidney, we also observe localized expression of the cellular phenotype (cilia loss, apoptosis, or proliferation). This suggests that in this model, and perhaps in human late-onset kidney diseases, non-genetic and/or epigenetic factors are necessary for expression of the phenotype. The disease-like allele of Alms1, although functionally able to support cilia formation in vitro, in young mice and in older mice in the kidney medulla, might reveal a compromised function in the cortex during aging. We also note that the expression of the cellular kidney phenotype in our model is not random but shows a variegated pattern on a tubule-by-tubule basis. Some tubule cross sections appear to have normal cilia, and little to no proliferation or apoptosis, whereas other tubule cross sections show extensive loss of cilia, sometimes with all or nearly all of the epithelial cells either proliferating or in apoptosis. The presumed somatic event, which, in combination with a genetic predisposition, causes some tubules to appear dysfunctional while neighboring tubules have a wild-type appearance, is not known. However, more detailed characterization of the progression of the phenotype might suggest treatments that maintain tubular integrity and delay disease.
Materials and Methods
Cell culture and ciliogenesis assay.
mIMCD3 cells were obtained from American Type Culture Collection (www.atcc.org) and cultured in DMEM/F12 media (Invitrogen, http://www.invitrogen.com) supplement with 10% FBS, 2.5 mM L-glutamine. For cilia formation assay, 50,000 cells were seeded on 12-mm Transwell filters (Costar). Filters were collected at day 0, day 1, day 3, day 5, and day 7 after seeding for RNA isolation and cilia immunostaining.
Knockdown of Alms1 expression by RNAi.
We designed siRNA specific to Alms1 by online BLOCK-iT RNAi designer (Invitrogen). These siRNA sequences are: 5′-GCTGTATGTAGTCGAATTA-3′ (Alms1a); 5′-GCCTGATTCCTTGTTTCAA-3′ (Alms1b); 5′-GCAGTAGTCTCTTCTGCTT-3′ (Alms1c); 5′-GCTTCAGCTTTGCTGAATT-3′ (Alms1d). SiRNA were synthesized by Qiagen (http://www1.qiagen.com). For RNAi transfection, mIMCD3 cells were seeded as previously described and transfected with Lipofectamine 2000 (Invitrogen) according to manufacturer's instruction.
DNA constructs.
The mouse Alms1 cDNA was generated from mIMCD3 mRNA. N-terminus (aa: 1–1,282) was cloned into p3XFLAG-CMV vector (Sigma, http://www.sigmaaldrich.com) to generate N-Alms1-FLAG plasmid.
RNA isolation and labeling for array analysis.
Cells were transfected on day 0 with either a scrambled siRNA control (one of two siRNAs directed against Alms1) or mock-transfected. Cells were seeded on day 1 and allowed to reach confluence through day 7. Cilia were formed in the mock-transfected and scrambled siRNA-transfected cells. Samples were taken for gene expression on days 0, 1, 3, 5, and 7 for the mock-transfected cells; days 0, 1, 3, and 5 for scrambled siRNA-transfected cells; and days 1, 3, and 5 for cells transfected with siRNAs against Alms1. RNA was isolated using the RNeasy Kit from Qiagen. Total RNA was examined on an Agilent Bioanalyzer (Agilent, http://www.agilent.com) and the 28S/18S ratio exceeded 2.0 for all samples. cDNA and cRNA were prepared from 4 μg total RNA according to standard Affymetrix protocols (http://www.affymetrix.com/support/technical/manual/expression_manual.affx). The in vitro transcription kit used was the GeneChip IVT labeling kit (Affymetrix). All reactions were processed at the same time by the same individual. We selected a group of 98 genes that had the highest relative gene expression change during ciliogenesis in the mock-transfected cells. The query was: (max expression level for days 0, 1, 3, 5, and 7 for the mock-transfected cells) divided by the maximum (the minimum expression level over these samples and a cutoff value of 400 arbitrary units). Four hundred units was approximately the 77th percentile of expression across all 36,000 probesets for a given sample. We selected the top 98 genes by this ratio (ratio >7.4) and then clustered the genes based on their expression levels in the mock-transfected samples. Comparison of this pattern with the samples that were transfected with either a scrambled or targeted siRNA showed no significant changes in the broad transcriptional program that was associated with ciliogenesis, suggesting that Alms1 is unlikely to play a role in transcription regulation associated with ciliogenesis.
Ca2+ imaging assay.
mIMCD3 cells, grown on cover glass, were transfected with Alms1 RNAi or scrambled control siRNA using Lipofectamine 2000 (Invitrogen), and cells were grown for 5 more d to induce cilia formation. Ca2+ imaging was performed as described [6]. Briefly, cells were incubated with 5 μM Fura2-AM (Molecular Probes, Invitrogen) for 30–60 min at 37 °C. Cover glasses were placed in a laminar flow perfuse chamber (Warner Instruments Corporation, http://www.warneronline.com). After 20 min equilibration, cells were perfused with media as described [6] via a local perfusion pipette. Image of Fura-2 loaded cells with excitation wavelength alternating between 340 nm and 380 nm were captured by a CCD camera. Following subtraction of background fluorescence, the ratio of fluorescence at two wavelengths was analyzed using Metafluor (Universal Imaging Corporation). All experiments have been repeated in triplicate and similar results were obtained.
Generation and phenotypic analysis of Alms1 mutant mice.
We identified Alms1L2131X/L2131X mice in an ethyl nitroso urea–forward genetics screen. Mutant mice were generated by successive intercrossing of heterozygote Alms1L2131X animals on a mixed C57Bl6/ NOD genetic background. Body composition was determined by quantitative magnetic resonance according to protocols supplied by the manufacturer (Echo Medical Systems, http://www.echomri.com). All plasma analytes were measured in retinal blood samples. Serum glucose, cholesterol, HDL, and triglyceride levels were determined by Olympus AU400e. Leptin and insulin levels were analyzed by ELISA (Crystal Chemical Incorporated).
Histology and immunohistochemistry.
Mice were anesthetized with Avertin (Sigma, 1.25%, 0.02 ml per gram body weight), perfused with 10% sucrose solution and 4% paraformaldehyde. Tissues were fixed in 4% paraformaldehyde, processed, and embedded in paraffin. 5-μm tissue sections were processed for hematoxylin-eosin (H&E) and immunostaining. For routine immunostaining, the sections were deparaffinized with xylene and rehydrated with graded ethanol. Primary antibodies used in this study were: γ-tubulin; acetylated tubulin (Sigma); β-Galactosidase (Abcam, http://www.abcam.com/index.html?); ki67 (NeoMarker Rabbit Monoclonal); aquaporin-2 (Santa Cruz Biotechnology, http://www.scbt.com). We generated polyclonal antibodies to Alms1 by injecting rabbits with the synthetic peptides MEPEDLPWPDELE, representing amino acids 1–13 of mouse Alms1. Dilutions were following the manufacturer's suggestions. Antibodies were visualized by Vectastain immunodetection kit (Vector Laboratories, Incorporated, http://www.vectorlabs.com). For indirect immunofloresence, Alexa488 and Texas Red conjugated secondary antibodies were obtained from Molecular Probes, Incorporated.
Isolation and culturing primary fibroblasts and kidney cortical duct cells.
Wild-type and Alms1L2131X/L2131X embryos were harvested at embryonic day 12.5 and minced by passage through a syringe with an 18 G needle. Embryonic fibroblasts were cultured in 10% FBS DMEM medium. For generation of primary kidney epithelial cells, mice (postnatal day 5) were anesthetized by intraperitoneal injection of 2.5% Avertin. Kidney cortices were dissected by handheld microtome. The slices were dissociated with DMEM containing 1 mg/ml collagenase. Digested tissues were then passed sequentially through 100-μm and 45-μm sieves, and centrifuged at 1,000 × g for 10 min at room temperature. The pellets were resuspended and cultured in renal epithelial growth medium (REGM Bullet Kit, Cambrex Bioscience, http://www.cambrex.com).
Supporting Information
Figure S1
Identification of Alms1L2131X/L2131X Strain
(A) Genotyping data for F2 mice derived from outcross of G3 mice from an ethyl nitroso urea screen on a C57Bl/6 background to NOD, followed by intercrossing of the F1 offspring. Genotype is marked as “B” for a B6-derived allele, “H” for heterozygote, and “N” for a NOD-derived allele. Based on the phenotype and genotype data, black shading denotes the included region for the position of the mutation and gray shading denotes the excluded region. Map positions refer to public mouse genome assembly M33 (http://www.ensembl.org/Mus_musculus/index.html).
(B) Resequencing Alms1 gene in mutant mice showing nonsense mutation at exon 10. Schematic structure of Alms1 was drawn to scale based on a genomic search of Alms1 cDNA (http://www.genome.ucsc.edu).
(327 KB JPG)
Click here for additional data file.
Figure S2
Serum Analytes in Alms1L2131X/L2131X Mice and Controls
Plasma glucose, triglyceride, insulin, leptin, total cholesterol, and HDL cholesterol measurements in 60- to 80-d-old and 140- to 170-d-old Alms1L2131X/L2131X, Alms1+/L2131X , and Alms1+/+ mice. Arrow, diabetic mutant male.
(445 KB JPG)
Click here for additional data file.
Figure S3
Loss of Cilia in the Kidney Cortex of Aged Alms1L2131X/L2131X Mice
Cilia are lost in the cortex tubules (A) but not in the medulla tubules (B) in an age-dependent manner. Cilia were stained with an anti-acetylated tubulin antibody.
(2.0 MB JPG)
Click here for additional data file.
Table S1
Gene Expression Changes after Knockdown of Alms1
Gene expression levels of 98 genes with the most dramatic changes in mIMCD3 cells transfected with Alms1 siRNAs compared to a scrambled siRNA control and a mock-transfected control.
(126 KB XLS)
Click here for additional data file.
Accession Numbers
The GenBank (http://www.ncbi.nlm.nih.gov/Genbank) accession number for Alms1 mouse cDNA is AF425257 and the RefSeq (http://www.ncbi.nlm.nih.gov/RefSeq) accession number for Alms1 mouse protein is NP660258.
Acknowledgements
We thank Richard Cornall, Samantha Zaharevitz, Conan Liu, Hanh Garcia, David Lloyd, Satchin Panda, James Watson, John Walker, and A. Huang for help and advice throughout the course of this work. We also thank M. Bandell and T. Jegla for technical assistance in Ca2+ imaging.
Abbreviations
ADPDK - autosomal dominant polycystic kidney disease
ARPKD - autosomal recessive polycystic kidney disease
BBS - Bardet-Biedl syndrome
IFT - intraflagellar transport
LTA - Lotus tetragonolobus agglutinin
mIMCD3 - mouse inner medullary collecting duct
PKD - polycystic kidney disease
siRNA - short interfering RNA
Figures and Tables
Figure 1
Suppression of Alms1 Expression Alters Primary Cilium Formation in Kidney Epithelial Cells
(A) Elongated cilia, visualized with staining of acetylated tubulin (green), form normally in mIMCD3 cells after mock-transfection, transfection with a negative control siRNA, or transfection with two inactive siRNAs directed against Alms1 (Alms1c and Alms1d). Focal staining of acetylated tubulin without axoneme extension is seen after transfection with two active siRNAs targeting Alms1 (Alms1a and Alms1b).
(B) Real-time PCR analysis with two mouse Alms1 probes recognizing the junctions of exons 1 and 2 and exons 12 and 13, respectively: Alms1a and Alms1b siRNAs both cause 70%–80% knockdown of Alms1 mRNA; no effect on Alms1 mRNA was seen with the three siRNAs that were inactive in the ciliogenesis assay.
(C) Alms1a siRNA-treated cells lose endogenous Alms1 protein expression. Acet, acetylated. Scale bars, 10 μm.
Figure 2
Loss of Alms1 Does Not Affect Transcriptional Changes during Ciliogenesis but Causes Impairment in Flow-Induced Mechanosensation
(A) Confocal microscopic analysis of cilia biogenesis in mIMCD3 cells. Short cilia can be detected at day 3 after transfection. mIMCD3 cells transfected with Alms1a siRNA have stunted cilia at days 3 and 5. Cells were stained with anti-acetylated tubulin (yellow, cilia) and TO-PRO-3 (red, nuclei).
(B) Suppression of Alms1 does not affect the upregulation of Bbs4 and Ttc10 during ciliogenesis. N, negative siRNA; 1a, Alms1a siRNA; 1b, Alms1b siRNA.
(C) Heat map representation of microarray analysis of mIMCD3 during ciliogenesis. 98 genes with the most dramatic changes in expression showed approximately equivalent expression changes in the presence of a scrambled siRNA control, or in the presence of specific siRNAs that decreased Alms1 mRNA levels and blocked cilia formation.
(D) Stunted cilia formed in the presence of Alms1a siRNA (red) lack flow-induced Ca2+ influx in mIMCD3 cells, compared with a negative control siRNA (blue). Representative data are shown for cytosolic calcium change of individual cells in response to mechanical flow. Arrow points to the start of flow.
Figure 3
N-Terminal Alms1 Protein Can Support Cilia Formation
(A) Cotransfection of Alms1a siRNA-treated cells with a 5′ Alms1 cDNA construct rescues primary cilia formation in mIMCD3 cells.
(B) Real-time PCR analysis of Alms1a siRNA and N-terminal Alms1-transfected cells. Upper panel: over-expression of the 5′ cDNA does not affect knockdown of endogenous Alms1 mRNA with Alms1a siRNA. Lower panel: knockdown of endogenous Alms1 mRNA does not affect overexpression of the 5′ cDNA. N, negative control siRNA; cDNA, 5′ Alms1 cDNA; 1a, Alms1a siRNA.
(C) Stable expression of Alms1 mRNA from the Alms1L2131X/L2131X allele. Real-time PCR analysis of Alms1 gene expression in an Alms1L2131X/L2131X mouse and a wild-type littermate control.
(D) The N-terminal mouse Alms1 antibody detects Alms1 mutant protein at the ciliary basal body in primary kidney cells from the Alms1L2131X/L2131X mouse. Shown are low and high magnifications of the ciliated cells. Arrowheads point out the base of cilia.
(E) Normal appearance of primary cilia in primary fibroblasts (MEF) and primary kidney cells (PKC) from the Alms1L2131X/L2131X mouse strain.
(F) Inhibition of cilia formation in Alms1a siRNA-treated Alms1L2131X/L2131X primary fibroblasts. Scale bars, 10 μm.
Figure 4
Alms1L2131X/L2131X Mice Recapitulate Human Alström Syndrome
(A) Alms1L2131X/L2131X mice gain more fat mass than heterozygote or wild-type controls but equivalent lean mass.
(B) Histological examination of Alms1L2131X/L2131X mice and wild-type littermate control. Insets show oil red O staining of frozen liver sections.
(C) Testis H&E sections show degeneration of seminiferous tubules (arrow), which have reduced numbers of germinal cells. Reduced numbers of sperm flagella with decreased length are observed in the epididymus of Alms1L2131X/L2131X mice (anti-acetylated tubulin, green). H&E, hematoxylin-eosin.
(D) Rhodopsin staining in the outer nuclear layer cell bodies is seen in rare cells in the Alms1L2131X/L2131X animals (arrows) but not in wild-type littermate controls. Insets illustrate higher magnification images. ONL, outer nuclear layer; IS, inner segment; OS; outer segment. Scale bars, 50 μm.
Figure 5
Kidney Abnormalities in Alms1 Mutant Mice
(A) H&E-stained kidney sections of a 6-mo-old Alms1L2131X/L2131X mouse showing dilated cortex tubules compared with an age-matched wild-type control. Lack of kidney cilia is observed in some tubules in the cortex of Alms1L2131X/L2131X kidney, whereas cilia in the medulla appear normal. H&E, hematoxylin-eosin; Acet, acetylated.
(B) Cortex cilia count comparison between Alms1L2131X/L2131X and controls. 300–400 kidney nuclei were examined for each of six fields of Alms1L2131X/L2131X and wild-type controls. The bar chart represents the average and standard deviations of cilia count per 100 kidney cortex epithelial cells from eight mice per group.
(C) In the cortex of Alms1L2131X/L2131X kidney, cilia are lost selectively in LTA-labeled tubules but not in aquaporin-2-expressing tubules.
(D) Upper panel: clusters of Ki67-positive proliferating epithelial cells in the Alms1L2131X/L2131X kidney, potentially lining the same convoluted tubule. Lower panel: TUNEL staining reveals apoptotic cells in Alms1L2131X/L2131X kidneys but rarely in a wild-type control. Arrow, whole tubule cross sections were labeled by TUNEL, suggesting progression of nephropathy in Alms1L2131X/L2131X mutant kidneys. WT, wild-type.
(E) Urinalysis of 3- to 6-mo-old Alms1L2131X/L2131X mice and age-matched littermate controls. Urine from Alms1L2131X/L2131X mice showed slight elevation of protein levels, p = 0.007. Scale bars, 50 μm.
Footnotes
¤ Current address: Genomics Institute of the Novartis Research Foundation, San Diego, California, United States of America
Competing interests. RJG owns stock in Phenomix.
A previous version of this article appeared as an Early Online Release on November 30, 2006 (doi:10.1371/journal.pgen.0030008.eor).
Author contributions. GL, KN, CG, PM, NAH, and RG conceived and designed the experiments. GL, RV, KN, and HW performed the experiments. GL, KN, NAH, and RG analyzed the data. GL, NG, and CG contributed reagents/materials/analysis tools. GL and RG wrote the paper.
Funding. This work was funded by Phenomix Corporation and the Genomics Institute of the Novartis Research Foundation. | [
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17465682 | PPAR gamma 2 Prevents Lipotoxicity by Controlling Adipose Tissue Expandability and Peripheral Lipid Metabolism
Abstract
Peroxisome proliferator activated receptor gamma 2 (PPARg2) is the nutritionally regulated isoform of PPARg. Ablation of PPARg2 in the ob/ob background, PPARg2−/− Lepob/Lepob (POKO mouse), resulted in decreased fat mass, severe insulin resistance, β-cell failure, and dyslipidaemia. Our results indicate that the PPARg2 isoform plays an important role, mediating adipose tissue expansion in response to positive energy balance. Lipidomic analyses suggest that PPARg2 plays an important antilipotoxic role when induced ectopically in liver and muscle by facilitating deposition of fat as relatively harmless triacylglycerol species and thus preventing accumulation of reactive lipid species. Our data also indicate that PPARg2 may be required for the β-cell hypertrophic adaptive response to insulin resistance. In summary, the PPARg2 isoform prevents lipotoxicity by (a) promoting adipose tissue expansion, (b) increasing the lipid-buffering capacity of peripheral organs, and (c) facilitating the adaptive proliferative response of β-cells to insulin resistance.
Author Summary
It is known that obesity is linked to type 2 diabetes, however how obesity causes insulin resistance and diabetes is not well understood. Some extremely obese people are not diabetic, while other less obese people develop severe insulin resistance and diabetes. We believe diabetes occurs when adipose tissue becomes “full,” and fat overflows into other organs such as liver, pancreas, and muscle, causing insulin resistance and diabetes. Peroxisome proliferator activated receptor gamma (PPARg) is essential for the development of adipose tissue and control of insulin sensitivity. PPARg2 is the isoform of PPARg regulated by nutrition. Here we investigate the role of PPARg2 under conditions of excess nutrients by removing the PPARg2 isoform in genetically obese mice, the POKO mouse. We report that removing PPARg2 decreases adipose tissue's capacity to expand and prevents the mouse from making as much fat as a normal obese mouse, despite eating similarly. Our studies suggest that PPARg plays an important antitoxic role when it is induced in liver, muscle, and beta cells by facilitating deposition of fat as relatively harmless lipids and thus prevents accumulation of toxic lipid species. We also show that PPARg2 may be involved in the adaptive response of beta cells to insulin resistance.
Introduction
An adipocentric view of the Metabolic Syndrome (MS) considers obesity as the major factor leading to insulin resistance in peripheral metabolic tissues. However, the link between obesity and insulin resistance is complex, as indicated by the fact that some extremely obese people are glucose tolerant, while others with a mild degree of obesity develop severe insulin resistance and diabetes. This suggests that the absolute amount of fat stored may not be the most important factor determining the relationship between obesity and insulin resistance. Recent work showing the complexity of the molecular mechanisms controlling adipogenesis [1,2] suggests that adipose tissue expandability may be an important factor linking obesity, insulin resistance, and associated comorbidities.
There are two mechanisms that have been proposed to explain how expansion of the adipose tissue stores affects insulin sensitivity. One mechanism suggests that increased adiposity induces a chronic inflammatory state characterized by increased cytokine production by adipocytes and/or from macrophages infiltrating adipose tissue. Cytokines produced by these adipocytes or macrophages may directly antagonise insulin signalling [3,4]. A second nonexclusive hypothesis is lipotoxicity. The lipotoxic hypothesis states that if the amount of fuel entering a tissue exceeds its oxidative or storage capacity, toxic metabolites that inhibit insulin action are formed [5–8]. Of particular relevance to this article, lipid metabolites, such as ceramides and diacylglycerol (DAG) or reactive oxygen species generated from hyperactive oxidative pathways, have been shown to inhibit insulin signalling and to induce apoptosis [9–11].
The nuclear receptor peroxisome proliferator activated receptor gamma (PPARg) is critically required for adipogenesis and insulin sensitivity [12–15]. There are two PPARg isoforms, PPARg1 and PPARg2. PPARg1 is expressed in many tissues and cell types, including white and brown adipose tissue, skeletal muscle, liver, pancreatic β-cells, macrophages, colon, bone, and placenta [16]. Under physiological conditions, expression of PPARg2, the other splice variant, is restricted to white and brown adipose tissue [16,17]. In adipose tissue PPARg is the key regulator of adipogenesis. PPARg2 is the more adipogenic PPARg isoform in vitro, it is also the isoform regulated transcriptionally by nutrition [17–20]. Although under physiological conditions expression of PPARg2 is limited to adipose tissues, we have shown that PPARg2 is ectopically induced in liver and skeletal muscle in response to overnutrition or genetic obesity [2,18]. De novo expression of PPARg2 in liver and muscle in obesity suggests that PPARg2 may have a role in insulin resistance and lipotoxicity in these tissues. Little in vivo research into the metabolic roles for the specific isoforms of PPARg has been carried out, with the studies so far focusing almost exclusively on adipose tissue [2,13,21,22]. PPARg (both isoforms) deletions have been generated in most major metabolic tissues. Liver-specific deletion of both PPARg isoforms caused an impairment in insulin sensitivity, particularly when challenged by different genetic backgrounds (lipoatrophic or leptin-deficiency) [23,24]. The effect of ablating both PPARg isoforms in muscle produced controversial results, with two groups reporting different effects on insulin sensitivity [25,26]. The role of PPARg in pancreatic β-cells is unclear, primarily due to its low expression under physiological conditions [27–29] and secondly because ablation of both PPARg isoforms in β-cells did not result in a metabolic phenotype. However PPARg may play a role in β-cell hyperplasia in response to insulin resistance, an idea supported by the fact that mice that lack PPARg in β-cells do not expand their β-cells mass in response to a high-fat diet [30]. More recently, it has been shown that heterozygous PPARg-deficient mice develop impaired insulin secretion, which is associated with increased islet triacylglycerol (TAG) content [31].
Here we investigate the physiological relevance of PPARg2 under conditions of positive energy balance by ablating PPARg2 in ob/ob mice. We use a new approach that integrates traditional physiological phenotyping with advanced lipidomic technology and transcriptomics. Our results indicate that in the context of positive energy balance, the absence of PPARg2 results in a major metabolic failure. Furthermore, we provide evidence that control of adipose tissue expansion by PPARg2 may be an important variable linking positive energy balance to its metabolic complications including insulin resistance, β-cell failure, and dyslipidaemia. Similarly, our lipidomic results indicate that induction of PPARg2 in nonadipose tissues should be considered as a physiological adaptation that prevents the toxic effects produced by excess nutrients. This antilipotoxic effect of PPARg2 is achieved by increasing the lipid-buffering capacity of peripheral organs and facilitating β-cell hyperplasia in response to insulin resistance.
Results
Ablation of PPARg2 in Ob/Ob Mice (POKO Mouse) Prevents Adipose Tissue Expansion in Response to Positive Energy Balance
PPARg2−/− Lepob/Lepob mice with genetic ablation of the PPARg2 isoform on the obese hyperphagic ob/ob background (POKO) were generated. Matings of PPARg2+/− Lepob/Lep+ mice followed the expected Mendelian distribution (Fisher's test = 0.074 and 0.135 for males and females, respectively). PPARg1 gene expression in white adipose tissue (WAT) from five-week-old POKO mice was similar to PPARg2 KO mice and was not significantly different from wild-type (WT) mice (Figure S1).
Figure 1A shows growth curves for male and female mice of four genotypes (WT, PPARg2 KO, ob/ob, and POKO mice) over a 12-week period. At birth, the body weight of male and female POKO mice was indistinguishable from other genotypes (unpublished data). The ob/ob mice quickly became heavier than their WT littermates, with significantly elevated body weight by four and six weeks of age in female and male mice, respectively. However, the POKO mice did not become obese, and their body weight remained close to WT and PPARg2 KO body weights mice during the 12-week study.
POKO mice were as hyperphagic (Figure 1B) as the ob/ob mice but drank far more water compared with ob/ob littermates (81.85 ± 15.14 versus 9.05 ± 2.32 ml/70 h, p < 0.01, female POKO versus ob/ob, n = 4 at 20 wk) (Figure S2A). Dual-energy X-ray absorptiometry analysis at 20 wk (Figure 1C) confirmed that female POKO mice had slightly increased fat content (4%) compared to WT and PPARg2 KO mice, but significantly reduced fat mass compared to the 40% increase observed in ob/ob mice. At the age of 20 wk, POKO and ob/ob mice had a trend to a decreased total locomotor activity during dark and light cycles compared with the WT and PPARg2 KO mice over the 72-h period. However POKO had similar total locomotor activity compared with ob/ob mice (Figure S2B).
At six weeks of age, female POKO mice consumed a similar amount of oxygen as ob/ob mice (vO2 = 25.06 ± 0.89 versus 23.10 ± 0.99 ml/kg bodyweight 0.75/min, p = 0.07 POKO versus ob/ob, n = 6–8) showing a lower respiratory exchange ratio (0.916 ± 0.011 versus 0.952 ± 0.007, p = 0.01, female POKO versus ob/ob) in the fed state, but similar respiratory exchange ratio in the fasted state (0.73 ± 0.014 versus 0.75 ± 0.018, p-value = 0.59 POKO versus ob/ob mice). Water intake was already significantly increased in POKO compared to ob/ob mice (13.59 ± 1.88 versus 8.15 ± 0.89 ml/d, p-value < 0.05, POKO versus ob/ob). Furthermore, levels of glucose in urine were higher in POKO mice compared with ob/ob mice (403.4 ± 49.2 versus 34.13 ± 13.5 mMol/l, POKO versus ob/ob mice, p-value = 0.001), showing an energy loss of 15.43 ± 3.06 kJ/d through urine compared with 0.70 ± 0.19 kJ/d in ob/ob mice. At this age, POKO mice showed similar locomotor activity compared with the ob/ob mice during the day, but increased locomotor activity during the night (Figure S2C).
Histomorphometric analysis of adipose tissue from 16-wk-old male mice revealed that POKO mice had fewer small adipocytes than the ob/ob mice (Figure 1D and 1E). This analysis of adipocyte size suggests that ablation of PPARg2 in the ob/ob background impairs the potential for adipocyte recruitment.
Early Insulin Resistance in POKO Mice Independent of Body Weight
As expected the reduced adipose tissue expandability of the POKO mouse was associated with severe insulin resistance. Surprisingly insulin resistance developed very early in life with elevated insulin levels and blood glucose compared to ob/ob mice (Table 1). We investigated whether peripheral insulin resistance and/or a severe defect in insulin secretion may cause hyperglycaemia in the POKO mouse. No differences in plasma glucose levels were detected three to five days after birth amongst the four genotypes for both genders (unpublished data). At weaning (three weeks of age) total body weight was indistinguishable amongst the four genotypes, and blood glucose levels were similar in males and females (Figure 2A). However, by the age of four weeks, coincident with the change to a chow diet, male and female POKO mice developed severe hyperglycaemia compared to the other genotypes. Insulin plasma levels in the POKO mice at four weeks of age were increased compared to ob/ob mice (Table 1). Insulin resistance in POKO mice was confirmed by an insulin tolerance test (ITT) in four-week-old male and female mice (Figure 2B). Furthermore insulin resistance in adipose tissue was demonstrated by the extremely low levels of glucose transporter4 (GLUT4) protein in POKO adipose tissue when compared with GLUT4 levels in adipose tissue from ob/ob mice (Figure S3). Of note, insulin resistance in the POKO mice was associated with hypertriglyceridaemia as early as four-weeks of age (Table 1).
Adult POKO Mice are Hyperglycaemic and Have Low Plasma Insulin Levels
Given the early insulin resistance and hyperinsulinaemia in the young POKO mice, we expected to see increased insulin levels in mature POKO mice. At 16 weeks, male POKO mice exhibited severe hyperglycaemia in the fasted and fed states compared to littermate controls. Male POKO mice had inappropriately low levels of insulin (Table 2). A similar, but milder phenotype was also observed in POKO female mice (unpublished data). Of note, adult ob/ob mice compensated for their insulin resistance with increased insulin levels (Table 2). POKO mice also had hypertriglyceridaemia when compared to WT, ob/ob, or PPARg2 KO mice.
Impaired Beta-Cell Function in the POKO Mice
The inappropriately low insulin levels in the adult POKO mice suggested a defect in β-cells. Insulin resistance in ob/ob mice was compensated for by increasing pancreatic insulin secretion, islet number, and size (Figure 3A). However, despite being more insulin resistant than ob/ob mice, POKO mice did not increase their β-cell mass, resulting in lower plasma insulin levels than the ob/ob controls. Morphometric analysis of pancreatic sections from 16-week-old male mice confirmed that the islet-to-pancreas volume ratios were similar in the POKO, WT, and PPARg2 KO mice (0.023 ± 0.005, 0.013 ± 0.006, and 0.016 ± 0.005, respectively) and markedly increased in ob/ob mice (0.077 ± 0.017, p < 0.01 ob/ob versus POKO). Additionally, POKO mice had significantly decreased islet number and size (average area of islets POKO = 18.40 ± 2 mm2) compared to ob/ob mice (ob/ob = 61.59 ± 8 mm2). Insulin staining demonstrated that islets from POKO mice contained fewer insulin-positive cells than islets from ob/ob mice (Figure 3A). The normal cellular organization of the islet, abundant β-cells (insulin staining) in the centre of the islet and a rim of α-cells at the periphery (glucagon staining), was retained in the insulin resistant ob/ob mice but was disrupted in the islets of POKO mice (Figure 3A). Islets from POKO mice had decreased number of insulin positive β-cells when compared to islets from ob/ob mice and a scattered pattern of α-cells, which are morphological changes associated with islet remodelling in the context of β-cell failure. Gene expression analysis of islets from 16-week-old mice revealed decreased expression of pancreatic duodenal homeobox-1, insulin receptor substrate 2, Glut2, and insulin in islets from POKO mice when compared with those from WT or ob/ob (Figure S4).
The changes seen in the β-cells of POKO mice were not the result of an inherent failure of the β-cell to develop properly as indicated by histological studies of neonatal pancreas (day 3 to day 5) (unpublished data) and four-week-old pancreas (Figure 2C), showing no morphological differences in the size, number, or insulin staining of islets from POKO mice when compared to ob/ob controls.
Impaired Glucose-Stimulated Insulin Secretion in POKO Mouse Islets
We measured glucose-stimulated insulin secretion in 16-week-old female POKO mice and their ob/ob littermates. Islets isolated from POKO mice were 30% smaller than those from ob/ob mice. Moreover, whereas normal islets were pure white with a smooth surface, islets from POKO mice were gray; their surface was irregular and required less time for collagenase digestion (only ten minutes instead of 30 minutes), suggesting that they were also more fragile.
Insulin content in islets from ob/ob mice was more than 30-fold greater than in those from POKO mice (Figure 3B). Insulin secretion from the islets of POKO mice was strikingly impaired compared to those of ob/ob mice, even when expressed relative to insulin content (Figure 3C). This was observed under basal (1 mM glucose) and stimulated (16 mM glucose, 16 mM glucose + tolbutamide) release.
Decreased Steatosis in POKO Mice Compared to Ob/Ob Mice
As expected, the POKO mice had increased hepatic fat deposition compared to WT and PPARg2 KO mice (Table S1), but surprisingly the POKO mouse had much milder hepatosteatosis than the ob/ob mouse (Figure 3D), suggesting that ectopic expression of the PPARg2 isoform in the liver of ob/ob mice (see below), might contribute to the deposition of TAGs in the liver.
Ablation of PPARg2 Induces a Lipotoxic Lipid Profile in Adipose Tissue, Pancreatic Islets, Liver, and Skeletal Muscle
To investigate lipotoxicity as a potential pathogenic mechanism we used liquid chromatography/mass spectrometry (LC/MS) [32] to compare a broad spectrum of cellular lipids in the adipose tissue, pancreatic islets, liver, and skeletal muscle between the POKO mouse and controls (Protocol S1).
Adipose tissue from POKO mice has decreased TAG but increased DAG, ceramides, and other reactive lipid species associated with insulin resistance.
Lipidomic analysis using LC/MS identified 74 molecular species differentially present in POKO, ob/ob, and WT mice (Protocol S1). POKO adipose tissue had decreased short chain TAGs compared to ob/ob adipose tissue (Protocol S1). Conversely, the concentration of DAGs was increased in the WAT of the POKO mice compared to ob/ob littermates. There was also an increased concentration of reactive lipid species in the WAT of POKO mice compared to that of ob/ob. The WAT of both POKO and ob/ob mice (Protocol S1) had increased levels of two ceramide species (with 16:0 and 24:1 fatty acid chains, respectively) and three proinflammatory lysophosphatidylcholine species [33] compared to WT mice. Partial least squares discriminant analysis indicated these changes in ceramides were greater in the POKO than ob/ob mice (Protocol S1). Sphingomyelin (d18:1/16:0), the precursor of ceramide (d18:1/16:0) and antioxidant ethanolamine plasmalogen (36:1) [34] were markedly decreased in POKO and ob/ob mice (Figure 4A).
Decreased TAG and accumulation of reactive lipid species in islets from POKO mice.
Partial least-squares discriminant analysis of lipidomic profiles of isolated pancreatic islets of 16-week-old mice identified 44 lipid species accumulated at different concentrations in WT, PPARg2 KO, and POKO mice (Protocol S1). Short chain TAGs were decreased in islets from POKO and PPARg2 KO mice when compared to those from WT. This was associated with up-regulation of phosphatidylethanolamine (36:2), down-regulation of ethanolamine plasmalogen (36:2), and preferential accumulation of reactive lipid species, particularly of two ceramides (20:0 and 22:0 fatty acids) in islets from POKO mice (Figure 5A and Protocol S1).
Decreased TAG and increased reactive lipid species in liver of POKO mice.
Multivariate analysis of lipidomic profiles (192 lipid species) revealed large changes between the POKO, PPARg2 KO, ob/ob, and WT genotypes (Protocol S1). These included decreased levels of short and medium chain TAGs and DAGs (Figure 5B) in livers from POKO mice compared to those of ob/ob mice. Livers from POKO mice also had decrease levels of phosphatidylcholine lipid species (Protocol S1) utilised during the formation and secretion of very low density lipoproteins [35]. Conversely, POKO livers were enriched in ceramides compared to ob/ob livers, which correlated with the extent of increased levels of lysophosphatidylcholines in POKO and ob/ob mice (Protocol S1).
Decreased TAG and accumulation of reactive lipid species in POKO skeletal muscle.
The same lipidomic pattern of decreased TAG and increased reactive lipid species previously observed in adipose tissue, β-cell, and liver was found to a milder degree in the skeletal muscle of POKO mice (Protocol S1). Briefly, when compared to ob/ob skeletal muscle, POKO skeletal muscle showed a decrease in very short-chain fatty acid TAGs and a slight decrease in levels of medium and long chain TAGs (Protocol S1). The skeletal muscle of POKO mice also had increased reactive lipids including ceramide (d18:1/18:0), DAGs, lysophosphatidylcholines, and sphingomyelins (precursors of ceramides) when compared to that of ob/ob mice.
Transcriptomic Analysis in POKO Mice Correlates with Lipidomic Changes
Given the lipotoxic profiles identified in the POKO mouse, we hypothesised changes in the expression of metabolic genes directly related to PPARg2 ablation and also compensatory changes in genes associated with cellular stress (Table S4).
Gene expression analysis in WAT.
Target genes of PPARg such as Glut4, adipsin, aP2, and adiponectin were decreased to a larger extent in the WAT of five- and 16-week-old POKO mice than in PPARg2 KO mice (Figure S1 and Figure 4B). At five weeks of age, when differences in body fat between female WT, ob/ob, and POKO mice are only starting to become evident, levels of GLUT4, aP2, and adiponectin mRNA levels were similar in WT and ob/ob mice, yet were markedly decreased in POKO mice. As the ob/ob mice aged (16 wk) and became obese and insulin resistant, the expression pattern of these PPARg targets in the WAT of ob/ob mice became similar to that of the POKO mice.
Results from the lipidomic analysis suggested major changes in the expression of genes involved in lipid metabolism (Figure 4B). Expression of stearoyl-coenzyme A desaturase 1 (Scd1) and sterol regulatory element-binding protein-1c (SREBP1c) were significantly lower in WAT from POKO mice compared to ob/ob mice. Furthermore, the decrease in TAGs and increased DAGs correlated with decreased expression of DAG acyltransferase 2, a key enzyme catalysing the final step in TAG synthesis, in the WAT of POKO mice compared with WAT from ob/ob mice. Again supporting the lipidomic profile, the expression of hormone-sensitive lipase, a rate-limiting enzyme for hydrolysis of diacylglycerides, was decreased in the WAT of POKO, PPARg2 KO, and ob/ob mice compared with WT mice, with the lowest levels observed in the POKO mice. Adipose triglyceride lipase levels were decreased in ob/ob and POKO compared with WT and PPARg2 KO mice, but without significant differences between ob/ob and POKO mice.
Oxidative stress has recently been suggested as a common mechanism of insulin resistance. Adipose tissue from POKO mice had increased oxidative stress compared to that of ob/ob mice as indicated by decreased gene expression levels of extracellular CuZn-superoxide dismutase, disruption of the glutathione pathway as indicated by decreased levels of gluthatione synthase, and increased levels of peroxidase and several gluthatione transferases (Table S2). We examined macrophage infiltration of adipose tissue as a potential marker of inflammation associated insulin resistance. Expression of CD68 and F4/80, both macrophage markers, was increased in the WAT of both POKO and ob/ob mice compared with WT and PPARg2 KO mice (Figure 4B). However their expression levels were lower in the POKO mice than the ob/ob mice suggesting that macrophage infiltration was not directly related to the exacerbated insulin resistance of the POKO mouse compared to the ob/ob mouse.
Gene expression in the POKO liver.
Reduced hepatic steatosis accompanied by altered lipid profiles suggested that lack of hepatic ectopic expression of PPARg2 might be affecting lipid storage and metabolism in the liver of the POKO mice. Expression of genes involved in lipid metabolism in liver (Figure 5C) revealed that, proportional to the accumulation of TAGs in the liver, fatty acid synthase, Scd1, and the fatty acid translocase (FAT/CD36) were increased in ob/ob and POKO livers compared to WT mice and were significantly decreased in liver from POKO mice compared with liver from ob/ob mice. Other lipogenic PPARg target genes such as Lpl were also decreased in the POKO liver compared to the ob/ob mice. The ob/ob mice also had a compensatory increase in the expression of genes involved in β-oxidation (e.g., Pparg, Lcad, Aox, Cpt1, and Ucp2). Interestingly expression of these pro-oxidative genes was decreased in the liver of POKO mice when compared to that of ob/ob mice suggesting PPARg2 may contribute to their regulation [36].
Although β-cell failure could account for the severe hyperglycaemia observed in the POKO genotype, hepatic gluconeogenesis function might be affected. We observed a robust up-regulation of PPARg coactivator 1 alpha (PPARGC1a, also known as PGC1a) expression in the POKO liver compared with the WT and ob/ob mice. PPARGC1a is up-regulated in fasting and is thought to induce gluconeogenesis [37]. In parallel with the increase in PPARGC1a, microarray analysis revealed increased mRNA levels of the progluconeogenic genes phosphoenolpyruvate carboxykinase 1 (Pepck1) and glucose-6-phosphatase (G6pc) in the livers of POKO mice when compared to those of ob/ob mice (Table S2), suggesting hepatic gluconeogenesis may contribute to the hyperglycaemia observed in POKO mice.
Gene expression analysis in skeletal muscle of POKO mice.
In 16-week-old POKO-mice skeletal muscle we observed down-regulation of Srebp1c and Ppargc1a and up-regulation of Ucp2 expression in skeletal muscle from POKO mice compared to that of WT mice. Similarly, expression of Lpl and Scd1 was down-regulated in the skeletal muscle of POKO mice when compared with that from ob/ob mice (Figure S5; Table S2). Gene set enrichment analysis of microarray data showed decreased expression of oxidative phosphorylation and mitochondrial components including electron transport chain complex components, in skeletal muscle from POKO mice when compared with that from ob/ob mice (Table S3).
Discussion
The link between obesity, insulin resistance, and diabetes while epidemiologically very clear is still not properly understood at a mechanistic level. An emerging concept is that the absolute amount of fat stored may be less important than the remaining storage capacity of the adipose tissue. Here we show that the PPARg2 isoform may be an important factor controlling obesity-induced comorbidities through two mechanisms: (a) by regulating nutritionally induced adipose tissue expandability and (b) when de novo expressed in nonadipose tissues, by allowing the storage of energy in the form of relatively harmless TAG species.
Previously we described the metabolic phenotype of the adult PPARg2 KO mouse [2], characterised by mild insulin resistance observed only in males. Given the greater adipogenic potency of PPARg2 compared with PPARg1 in vitro, we expected PPARg2 KO mice to have many more severe defects in adipose tissue than we observed, and therefore insulin sensitivity. As PPARg2 is the PPARg isoform regulated in response to nutrition and obesity [17–20], we hypothesised that PPARg2 would only become essential for adipose tissue function in the face of positive energy balance. The metabolic challenge we opted for was PPARg2 ablation in the obese (ob/ob) background (PPARg2−/− Lepob/Lepob, POKO mouse). The POKO mouse had severely decreased body-fat mass due to impaired adipose tissue expandability. Despite eating as much as an ob/ob mouse and expending a similar amount of energy, the POKO mouse was unable to store fat efficiently in its adipose tissue. This mismatch between increased energy availability and lack of adipose tissue expandability lead to a global metabolic failure characterised by severe insulin resistance, β-cell failure, and dyslipidaemia.
The observation of reduced fat mass and increased insulin resistance in the POKO mouse compared to the ob/ob mouse strongly supports two of our hypotheses. First, we hypothesised that PPARg2 is required to recruit new adipocytes in overnutrition, but it is not required to make adipocytes during development. This is reflected by similar expression of aP2, a late marker of adipocyte differentiation, in POKO and ob/ob mice. The absence of small adipocytes was markedly different to other forms of lipodystrophy [38,39]. Additionally, and again in contrast with other lipodystrophic models that have markedly less adipose tissue than WT controls [38–40], the POKO mice had a percentage body fat that was similar (only 4% more) to WT and PPARg2 KO mice, as opposed to ob/ob mice, which had 40% fat as a proportion of body mass. This suggests that the remaining PPARg1 isoform is sufficient to support development of adipose tissue and fat deposition requirements of a lean mouse model. However, under conditions of positive energy balance, adipose tissue expandability mainly relies on the PPARg2 isoform. This idea is also suggested by the studies in heterozygous mice harbouring the murine equivalent of the human mutation (P465L) in PPARg on an ob/ob background [41]. These mice were able to accumulate fat and become obese even though showing a body mass 14% lower than ob/ob controls. In humans there is also evidence for a role for PPARg2. We have observed that metabolically healthy, nondiabetic, morbidly obese individuals have elevated levels of PPARg2 in their adipose tissue when compared to lean individuals [19]. Our second hypothesis, that the mismatch between energy availability and adipose tissue expandability is more important than fat mass itself as a predictor of insulin resistance, is also supported by our data. In fact the ob/ob mouse is much more obese than the POKO mouse but is much less insulin resistant. Furthermore, the POKO mice were already more insulin resistant than the ob/ob mice by the age of four weeks, with very low levels of GLUT4 in adipose tissue, before large differences in body weight developed, suggesting that the bioenergetic mismatch rather than the total amount of fat stored is important for the development of insulin resistance.
Although we hypothesised that the POKO mice would become insulin resistant, the degree of hyperglycaemia in these animals was in excess of what we expected. We found that the normal adaptive response of β-cells to insulin resistance did not occur in the POKO mice as indicated by the pathological changes observed by histology and the lack of β-cell hypertrophy. Although it has been shown that genetic background can affect the ability of ob/ob mice to undergo β-cell hypertrophy [42,43], we found that the ob/ob controls on our mixed 129Sv × C57BL/6J background underwent adaptive β-cell hyperplasia and hypertrophy, suggesting that the lack of PPARg2 was responsible for the failure of the POKO β-cells to adapt to insulin resistance. Interestingly the mass of pancreatic islets in POKO mice remained similar to the noninsulin resistant WT and PPARg2 KO mice. Furthermore, these defects in POKO β-cells did not appear to be the result of a developmental defect, as new born and four-week-old mice had morphologically normal islets.
The severe β-cell phenotype of the POKO mouse contrasts with the absence of hyperglycaemia observed in the pancreatic β-cell specific PPARg KO mouse [30]. However it should be kept in mind that in the β-cell specific PPARg KO mouse, the expression of PPARg and the lipid storage capacity of other tissues, most importantly adipose tissue, were not affected, and that insulin sensitivity was only mildly affected by high fat feeding in these mice when compared to the severe insulin resistance observed in POKO mice. Therefore the challenge to the pancreatic β-cells in this model was milder than in POKO mice. This is a clear example of how tissue-specific genetic manipulations are not always the best approach to understand the physiology of an organ in the context of the global energy homeostasis. The potential importance of the de novo expression of PPARg2 isoform in β-cells is also supported by the observation that humans harbouring the Pro12Ala mutation in PPARg2, a mutation that is located in the g2 isoform and makes PPARg2 less active, has only been associated with insulin deficiency and disease severity in obese individuals with type 2 diabetes [44].
The liver of the POKO mouse also displayed an unusual phenotype. We expected the POKO mice to have worse hepatosteatosis with increased triglyceride deposition in liver compared to ob/ob mice, because the POKO mice could not store fat in adipose tissue. However POKO mice had less hepatosteatosis than ob/ob mice suggesting that the PPARg2 isoform may directly contribute to facilitate triglyceride deposition in the liver.
A common mechanistic link for the phenotypes observed in the POKO liver and β-cell was not immediately obvious. To try to determine the role of PPARg2 in these locations we performed lipidomic and gene expression analyses of the adipose tissue, pancreatic islet, liver, and skeletal muscle of the POKO mouse. The lipid pattern of adipose tissue from POKO mice was characterised by decreased TAGs and increased DAGs in parallel with decreased gene expression of DGAT2, hormone-sensitive lipase, and adipose triglyceride lipase. This decrease in TAGs in the POKO adipose tissue was associated with increased levels of reactive lipid species and a gene expression profile suggestive of increased oxidative stress [45–49]. Although it has been described that oxidative stress and insulin resistance may be related to infiltration of adipose tissue by macrophages, resulting in a chronic state of inflammation [50–52], we did not observe increased macrophage infiltration in the adipose tissue of POKO mice compared to that of ob/ob mice.
Lipidomic analysis of POKO derived islets also showed decreased levels of triacyl and DAGs and increased levels of ceramides, suggesting that PPARg2 may contribute to increasing the lipid-buffering capacity of β-cells by promoting formation of TAGs and thus preventing lipotoxic insults. Liver and skeletal muscle lipidomics also showed reduced TAG and increased formation of reactive lipid species such as ceramides and lysophosphatidylcholines in POKO mice compared to ob/ob mice. This lipid profile was associated with impaired expression of pathways controlling de novo lipogenesis, transport of fatty acids, and beta oxidation in the POKO mice compared with the ob/ob mice. Of interest, Ppargc1a and other gluconeogenic genes were induced in the liver of POKO mice compared to that of ob/ob mice, suggesting a potential mechanism contributing to marked hyperglycaemia in POKO mice [53,54].
Overall, our lipidomic studies identify a remarkably similar pattern of changes in lipid species in the four tissues studied. The reduced adipose tissue mass and hepatosteatosis in the POKO mouse compared to the ob/ob mouse is explained by reduced levels of mature TAG in the POKO mouse. Similarly, ablation of PPARg2 resulted in accumulation of reactive lipid species implicated in causing insulin resistance, not only in adipose tissue, but also in other organs involved in whole-organism glucose metabolism. These results indicate that expression of PPARg2 in the pancreas, liver, and muscle of the ob/ob mouse may be performing a protective role, by increasing the capacity of these organs to buffer toxic lipid species by allowing accumulation of relatively harmless TAGs. The importance of this peripheral antilipotoxic role of PPARg2 becomes more evident if we consider that POKO and ob/ob mice are under the same degree of positive energy balance as determined by similar food intake, locomotor activity, and energy expenditure, that both models lack leptin, and that the only difference between ob/ob and POKO mice is the presence or absence of PPARg2. Given the decreased adipose tissue expandability of the POKO mice compared to ob/ob, it was anticipated that, as in the liver, muscle, or β-cells of lipodistrophic mice, the POKO mouse would accumulate more fat than the ob/ob. However, our results clearly indicate that mice lacking PPARg2, despite massive nutrient availability, are unable to deposit TAG in peripheral tissues and instead accumulate reactive lipid species in these organs. Therefore the pathologies of the liver and β-cell observed in the POKO mouse may be a result of a common lipotoxic insult facilitated by the absence of PPARg2 (Figure 6).
In summary, in this study we provide new insights into the physiological relevance of the PPARg2 isoform and identify adipose tissue expandability as an important determinant of metabolic complications. Ablation of PPARg2 decreases adipose tissue expandability, but its pathophysiological effects only become relevant in the context of a mismatch between energy availability and adipose tissue expansion. We show that PPARg2 also plays protective role when expressed de novo in peripheral organs by increasing their capacity to buffer toxic lipids. Ablation of PPARg2 under conditions of positive energy balance determined by absence of leptin produced early development of severe insulin resistance, β-cell failure, diabetes, and hyperlipidaemia. Extrapolation of this model to humans may suggest that normal to overweight individuals with positive energy balance and inappropriately severe manifestations of the MS may have a defect in PPARg2 and/or alternative mechanisms that control adipose tissue expandability.
Materials and Methods
Generation of mice homozygous for PPARg2 KO and leptin deficiency (ob/ob).
Mice heterozygous for a disruption in exon B1 of PPARg2 on a 129Sv background (PPARg2+/−) [2] were crossed with heterozygous ob/ob (Lepob/Lep+) mice on a C57Bl/6 background to obtain mice heterozygous for both the PPARg2 ablation and the leptin point mutation (PPARg2+/− Lepob/Lep+). These mice were crossed to obtain the four experimental genotypes: WT (PPARg2+/+ Lep+/Lep+), PPARg2 KO (PPARg2−/− Lep +/Lep+), ob/ob (PPARg2+/+ Lepob/Lepob), and POKO (PPARg2−/− Lepob/Lepob). Genotyping for deletion of PPARg2 and the point mutation in the ob gene was performed by PCR using standard protocols [2,55].
Animal care.
Animals were housed at a density of four animals per cage in a temperature-controlled room (20–22 °C) with 12-h light/dark cycles. Food and water were available ad libitum unless noted. All animal protocols used in this study were approved by the UK Home Office and the University of Cambridge.
Blood and urine biochemistry, food intake, and body composition analysis.
Mice of the four experimental genotypes were placed at weaning (three weeks of age) on a normal chow diet (10% of calories derived from fat; D12450B, Research Diets, http://www.researchdiets.com). Enzymatic assay kits were used for determination of plasma FFAs (Roche, http://www.roche.com) and TAGs (Sigma-Aldrich, http://www.sigmaaldrich.com). Elisa kits were used for measurements of leptin (R & D Systems, http://www.rndsystems.com), insulin (DRG Diagnostics International Limited, http://www.drg-international.com), and adiponectin (B-Bridge International, http://www.b-bridge.com) according to manufacturers' instructions. Dual-energy X-ray absorptiometry (DEXA, Lunar Corporation, http://www.lunarcorp.com) was used to measure body composition; glucose in blood and in urine and food intake were monitored in the four experimental genotypes as previously shown [2].
Oxygen consumption, water intake, and locomotor activity.
Oxygen was measured using an eight-chamber open-circuit oxygen-monitoring system attached to and sampled from the chambers of a Comprehensive Laboratory Animal Monitoring System (CLAMS; Columbus Instruments, http://www.colinst.com). Water consumed was also measured using CLAMS. Mice were housed individually in specially built Plexiglass cages maintained at 22 °C under an alternating 12:12-h light-dark cycle (light period 08:00–20:00). Sample air was sequentially passed through oxygen (O2) and carbon dioxide (CO2) sensors (Columbus Instruments) for determination of O2 and CO2 content. Mice were acclimatized to monitoring cages for 72 h before data collection. Mice were weighed before each trial. Ambulatory activity of individually housed mice was evaluated using an eight-cage rack OPTO-M3 Sensor system (Columbus Instruments). Cumulative ambulatory activity counts were recorded every 5 min throughout the light and dark cycles.
Calculations of energy lost in urine.
Energy lost in urine was calculated accordingly as previously shown before [56] using the following calculations:
Energy lost in urine kJ/day = (glucose in urine [mMol/l]/1,000) × molecular weight glucose × (water intake [ml/day]/1,000) × E densitycarb; E densitycarb = energy density related to oxidations within the body for carbohydrates as glucose = 15.76 kJ/g.
RNA preparation and real-time quantitative RT-PCR.
Total RNA was isolated from islets and tissues samples according to the manufacturer's instructions (RNAeasy kit, Qiagen, http://www.qiagen.com) and STAT60 (Tel-Test, http://www.isotexdiagnostics.com/tel-test.html). Real-time quantitative PCR was performed using a TaqMan 7900 (Applied Biosystems, http://www.appliedbiosystems.com) according to standard protocols.
Western blot analyses.
The tissue samples (40 μg) were subjected to SDS-PAGE on 8% polyacrylamide gels. Proteins were then electrophoretically transferred to polyvinylidene difluoride filters. After transferring, the filters were blocked with 5% nonfat dry milk in TBS-Tween 20 followed by incubation with primary GLUT4 and extracellular signal-regulated kinase 1/2 (ERK1/2) antibodies (Promega, http://www.promega.com) overnight. The bands were quantified by scanning densitometry.
Light microscopy and immunohistochemcal analysis.
Tissue samples for morphological and immunohistochemcal analysis were prepared according to published protocols [2]. Morphometric analyses of adipose tissue and pancreas sections were acquired using a digital camera and microscope (Olympus BX41, http://www.olympus.com), and cell areas were measured using AnalySIS software (Soft Imaging System, http://www.soft-imaging.net). For adipose tissue, two fields from each section were analysed to obtain the mean cell-area per animal (n = 5 per genotype). The Computer Assisted Stereology Toolbox (CAST) 2.0 system from Olympus was used to perform all measurements in the pancreas according to published protocols [57].
Isolation and culture of pancreatic islets.
The pancreas was injected via the bile duct with cold Hank's solution containing 0.4% (w/v) liberase (Roche). The pancreas was removed, digested for 15–30 min, and islets collected by handpicking. Isolated islets were cultured overnight in h-cell medium (SBMI 06, hcell technology, http://www.hcell.com) at 37 °C in 5% CO2 in air. Islets were used the day after isolation for insulin secretion studies or RNA extraction.
Insulin secretion studies.
Insulin secretion from isolated islets (five islets/well) was measured during 1-hr static incubations in Krebs—Ringer Buffer containing either 1 mM glucose, 16.7 mM glucose, or 16.7 mM glucose plus 200 μM tolbutamide in DMSO. The supernatants were assayed for insulin. Insulin content was extracted using 95:5 ethanol/acetic acid. Insulin was measured using a Mouse Insulin ELISA kit (Mercodia, http://www.mercodia.com). Islets were isolated from three mice of each genotype for these experiments. Thus, the data are the mean of three separate experiments, in which data were collected for each test solution from six samples each of five islets. For each sample, insulin release was normalised to insulin content.
ITT.
ITTs on four-week-old mice were performed as previously published [58].
Lipid profiling.
For WAT and muscle, the tissue sample (50 mg) was homogenized with 0.15 M sodium chloride (300 μl), and the lipids were extracted with 2 ml of chloroform: methanol (2:1) and used for LC/MS as previously described [2].
For liver and islets, an aliquot (20 μl for liver or 10 μl for islets) of an internal standard mixture (11 reference compounds at concentration level 8–10 μg/ml), 50 μl of 0.15 M sodium chloride (for liver), and chloroform:methanol (2:1) (200 μl for liver or 90 μl for islets) was added to the tissue sample (20–30 mg). The sample was homogenized, vortexed (2 min for liver or 15 s for islets), let to stand (1 h for liver, 20 min for islets), and centrifuged at 10,000 RPM for 3 min. From the separated lower phase, an aliquot was mixed with 10 μl of a labelled standard mixture (three stable isotope-labelled reference compounds at concentration level 9–11 μg/ml), and 0.5–1.0 μl injection was used for LC/MS analysis.
Total lipid extracts were analysed on a Waters Q-Tof Premier mass spectrometer (http://www.waters.com) combined with an Acquity Ultra Performance LC (UPLC). The column, which was kept at 50 °C, was an Acquity UPLC BEH C18 10 × 50 mm with 1.7 μm particles. The binary solvent system (flow rate 0.200 ml/min) included A, water (1% 1 M NH4Ac, 0.1% HCOOH), and B, LC/MS grade (Rathburn, http://www.rathburn.co.uk) acetonitrile/isopropanol (5:2, 1% 1 M NH4Ac, 0.1% HCOOH). The gradient started from 65% A/35% B, reached 100% B in 6 min, and remained there for the next 7 min. The total run time per sample, including a 5 min re-equilibration step, was 18 min. The temperature of the sample organizer was set at 10 °C.
Mass spectrometry was carried out on Q-Tof Premier (Waters) run in ESI+ mode. The data were collected over the mass range of m/z 300–1,200 with scan duration of 0.2 s. The source temperature was set at 120 °C, and nitrogen was used as desolvation gas (800 l/h) at 250 °C. The voltages of the sampling cone and capillary were 39 V and 3.2 kV, respectively. Reserpine (50 μg/l) was used as the lock spray reference compound (5 μl/min; 10 s-scan frequency).
Data processing was performed using the MZmine software [59]. Identification was performed based on an internal reference database of lipid species, or alternatively utilizing the tandem mass spectrometry. The statistical analyses were performed using Matlab (Mathworks, http://www.mathworks.com) and the Matlab library PLS Toolbox (Eigenvector Research, http://www.eigenvector.com).
Tandem mass spectrometry was used for the identification of selected lipid species. MS/MS runs were performed by using ESI+ mode, collision energy ramp from 15–30 V, and mass range starting from m/z 150. The other conditions were as shown in the Protocol S1.
Statistics.
Results were expressed as mean ± standard error of mean. Statistical analysis was performed using a two-tailed unpaired t-test between appropriate pairs of groups, and significance declared if p-values were less than 0.05.
Supporting Information
Figure S1
Adipose Tissue and Liver Gene Expression
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Figure S2
Water Consumed and Locomotor Activity
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Figure S3
GLUT4 protein expression in WAT
(65 KB PPT)
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Figure S4
Gene Expression in Islets
(25 KB PPT)
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Figure S5
Gene Expression in Muscle
(32 KB PPT)
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Protocol S1
POKO Mouse Model Lipidomics Dataset
(208 KB PDF)
Click here for additional data file.
Table S1
Tissue Weights of 16-Wk-Old Male POKO, Ob/Ob, PPARg2 KO, and WT mice
(29 KB PPT)
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Table S2
Microarray Data
(105 KB DOC)
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Table S3
Pathway Analysis from Microarray Data
(112 KB XLS)
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Table S4
Accession Numbers
GenBank (http://www.ncbi.nlm.nih.gov/Genbank) accession numbers for the genes and gene products discussed in this paper.
(58 KB DOC)
Click here for additional data file.
Acknowledgements
Animal care and husbandry provided by J. Carter, S. Shelton, H. Wetsby, H. Williams, A. Kant, J.P. Whiting, and G. Bevan. We thank D. Lam, M. Dale, and K. Burling for their technical assistance. We also thank J. Skepper and P.M. Coan for their help with morphometry analysis in pancreas and Peter Murgatroyd for his help with oxygen consumption experiments. We acknowledge Paradigm Therapeutics (http://www.paradigm-therapeutics.co.uk) for generating the PPARg2 KO mouse.
Abbreviations
DAG - diacylglycerol
GLUT - glucose transporter
H and E - haematoxylin and eosin
ITT - insulin tolerance test
LC/MS - liquid chromatography/mass spectrometry
MS - Metabolic Syndrome
PPARg - peroxisome proliferator activated receptor gamma
PPARg2 - peroxisome proliferator activated receptor gamma 2
PPARGC1a - PPARg coactivator 1 alpha
Scd1 - stearoyl-coenzyme A desaturase 1
TAG - triacylglycerol
WAT - white adipose tissue
WT - wild type
Figures and Tables
Figure 1
Physiological Characterisation of POKO Mouse
(A) Body weights (black circles, WT; black squares, ob/ob; white circles, PPARg2 KO; white squares, POKO) are shown for males (left) or females (right) (n = 5–12). *, p < 0.05 POKO versus ob/ob and §, p < 0.01 POKO versus WT.
(B) Food intake from 20-wk-old female mice (n = 4) is shown.
(C) Body composition analysis from 20-wk-old females is shown: WT, ob/ob, PPARg2 KO, and POKO mice fed chow diet mice (n = 4–7). *, p < 0.05 POKO versus WT and ###, p < 0.001 POKO versus ob/ob.
(D) Haematoxylin and eosin (H and E)-stained sections (10×) from epididymal WAT from 16-wk-old male WT, ob/ob, and POKO mice.
(E) Percent relative cumulative frequency analysis (PRCF) from epididymal WAT adipocytes from 16-wk-old male WT, ob/ob, PPARg2 KO, and POKO mice. (n = 4–5).
Figure 2
Early Insulin Resistance in POKO Mice Independent of Body Weight
(A) Body weight and plasma glucose levels from three, four, and five-week-old female WT, ob/ob, PPARg2 KO, and POKO. *, p < 0.05; **, p < 0.01; ***, p < 0.001 POKO versus ob/ob.
(B) Plasma glucose levels during ITT on 4-wk-old male (left) and female (right) mice on chow diet (black triangle, WT; white triangle, PPARg2 KO; black square, ob/ob; black diamond, POKO) (n = 7). *, p < 0.05; **, p < 0.01 POKO versus ob/ob.
(C) Morphological analysis of H and E-stained sections (10×) in pancreas from 4-wk-old males ob/ob and POKO mice (n = 5).
Figure 3
Impaired β-Cell Function and Hepatic Morphological Analyis in the POKO Mice
(A) H and E-stained sections (10×) and immunohistochemical (20×) analysis of insulin and glucagon in pancreas from 16-wk-old males WT, ob/ob, and POKO mice (n = 5).
(B) Insulin content of islets isolated from POKO (black bars), and ob/ob (grey bars) mice. Each data point is the mean of six samples each of five islets.
(C) Insulin secretion from islets isolated from POKO (black bars) and ob/ob (grey bars) mice in response to glucose (1, 16 mM) or glucose 16 mM + tolbutamide (200 μM). Data were collected from six samples each of five islets from three mice of each genotype. For each sample, insulin release was normalised to insulin content. *, p < 0.05; **, p < 0.01; ***, p < 0.001 POKO versus ob/ob.
(D) H and E-stained sections (4×) in liver from 16-wk-old males WT, ob/ob, and POKO mice (n = 5).
Figure 4
Lipidomic and Gene Expression Analysis of POKO WAT
(A) Lipidomic profiling of WAT from 16-wk-old males WT, ob/ob, and POKO mice.
(B) Adipose tissue mRNA levels from different genes from 16-wk-old male WT, PPARg2 KO, ob/ob, and POKO mice (n = 6–8). *, p < 0.05; **, p <0.01; ***, p <0.001 POKO versus ob/ob.
Figure 5
Lipidomic and Gene Expression Analysis in Islets and Liver from POKO Mice
Lipidomic profiling of islets (A) and liver (B) from 16-wk-old males WT, PPARg2 KO, ob/ob, and POKO mice. TG, TAGs; DAGs, diacylglycerols; SM, sphingomyelins. (C) Liver gene expression from 16-wk-old male WT, ob/ob, PPARg2 KO, and POKO mice fed chow (n = 6–8). *, p < 0.05; **, p, <0.01; ***, p <0.001 POKO versus ob/ob; ###, p < 0.001 POKO versus WT; ‡‡‡, p < 0.001 ob/ob versus WT.
Figure 6
Storage of Lipids—Antilipotoxic Role of PPARg2
Antilipotoxic role of PPARg2 mediated by (a) expansion of adipose tissue and facilitation of triglyceride deposition and (b) facilitating deposition of fat in liver, skeletal muscle, and pancreas in the form of TAG. Ob/Ob mice can induce PPARg2 expression in liver, muscle, and β-cell, facilitating deposition of excess of energy in these organs in the form of TAG. Absence of inducibility of PPARg2 in POKO mouse liver, muscle, and β-cells results in increased deposition of reactive lipid species and decreased TAG, leading to marked insulin resistance and β-cell failure.
Table 1
Metabolic Parameters in Fed 4-Wk-Old Male and Female POKO, Ob/Ob, PPARg2 KO, and WT Mice
Table 2
Metabolic Parameters in 16-wk-Old Male POKO, Ob/Ob, and WT Mice
Footnotes
Competing interests. The authors have declared that no competing interests exist.
Author contributions. GMG, SLG, FMA, and AVP conceived and designed the experiments. GMG, SLG, LY, KS, SV, MB, ML, and MO performed the experiments. GMG, SLG, LY, KS, SV, MC, RKC, MJL, TSL, and MO analysed the data. GMG, LY, KS, SV, MC, RKC, MB, and GSHY contributed reagents/materials/analysis tools. GMG and AVP wrote the paper.
Funding. This work was supported by the European Union FP6 Hepatic and Adipose Tissue and Functions in the Metabolic Syndrome (Hepadip) integrated program (http://www.hepadip.org) (LSHM-CT-2005–018734); Diabetes UK; Medical Research Council; Wellcome Trust Integrative Physiology program; Academy of Finland (grant number 111338); and Marie Curie International Reintegration Grant from the European Community. | [
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17503968 | A KATP Channel-Dependent Pathway within α Cells Regulates Glucagon Release from Both Rodent and Human Islets of Langerhans
Abstract
Glucagon, secreted from pancreatic islet α cells, stimulates gluconeogenesis and liver glycogen breakdown. The mechanism regulating glucagon release is debated, and variously attributed to neuronal control, paracrine control by neighbouring β cells, or to an intrinsic glucose sensing by the α cells themselves. We examined hormone secretion and Ca2+ responses of α and β cells within intact rodent and human islets. Glucose-dependent suppression of glucagon release persisted when paracrine GABA or Zn2+ signalling was blocked, but was reversed by low concentrations (1–20 μM) of the ATP-sensitive K+ (KATP) channel opener diazoxide, which had no effect on insulin release or β cell responses. This effect was prevented by the KATP channel blocker tolbutamide (100 μM). Higher diazoxide concentrations (≥30 μM) decreased glucagon and insulin secretion, and α- and β-cell Ca2+ responses, in parallel. In the absence of glucose, tolbutamide at low concentrations (<1 μM) stimulated glucagon secretion, whereas high concentrations (>10 μM) were inhibitory. In the presence of a maximally inhibitory concentration of tolbutamide (0.5 mM), glucose had no additional suppressive effect. Downstream of the KATP channel, inhibition of voltage-gated Na+ (TTX) and N-type Ca2+ channels (ω-conotoxin), but not L-type Ca2+ channels (nifedipine), prevented glucagon secretion. Both the N-type Ca2+ channels and α-cell exocytosis were inactivated at depolarised membrane potentials. Rodent and human glucagon secretion is regulated by an α-cell KATP channel-dependent mechanism. We propose that elevated glucose reduces electrical activity and exocytosis via depolarisation-induced inactivation of ion channels involved in action potential firing and secretion.
Author Summary
Glucagon is a critical regulator of glucose homeostasis. Its major action is to mobilize glucose from the liver. Glucagon secretion from α cells of the pancreatic islets of Langerhans is suppressed by elevated blood sugar, a response that is often perturbed in diabetes. Much work has focused on the regulation of α-cell glucagon secretion by neuronal factors and by paracrine factors from neighbouring cells, including the important islet hormone insulin. In contrast, we provide evidence in support of a direct effect of glucose on α cells within intact rodent and human islets. Notably, our work implicates an α-cell glucose-sensing pathway similar to that found in insulin-secreting β cells, involving closure of ATP-dependent K+ channels in the presence of glucose. Furthermore, we find that membrane depolarisation results in inhibition of Na+ and Ca2+ channel activity and α-cell exocytosis. Thus, we propose that elevated blood glucose reduces α-cell electrical activity and glucagon secretion by inactivating the ion channels involved in action potential firing and secretion.
Introduction
Blood glucose levels are under the control of two hormones released from the pancreatic islets of Langerhans. Islet β cells secrete insulin when glucose is high, decreasing glucose production by the liver and increasing glucose storage in multiple tissues. Regulated insulin secretion is relatively well understood, involving the metabolic stimulation of electrical activity, Ca2+ entry, and exocytosis [1]. Islet α cells secrete glucagon in response to decreased blood glucose, whereas elevated glucose levels suppress glucagon release. Glucagon is the principal factor stimulating glucose production by the liver. In diabetes, baseline glucagon release is elevated, and glucagon secretion in the low-glucose condition is blunted [2–4]. These effects contribute to chronic hyperglycaemia and to an increased risk for acute hypoglycaemic events.
The mechanism regulating glucagon secretion is poorly understood and remains hotly debated [5]. Glucagon release in rodents may be regulated by paracrine signals, including γ-aminobutyric acid (GABA) [6,7], Zn2+ [8], and insulin [9,10]. Conversely, glucose may suppress glucagon secretion through a direct effect on α-cell activity [11–14]. There are also studies suggesting that glucagon secretion is under hypothalamic control [15,16]. Human in vivo studies provide conflicting evidence regarding the control of glucagon secretion by paracrine or intrinsic regulation of α cells [17–30], and very little work has been done to examine this question in isolated human islets.
Islet α cells express ATP-dependent K+ (KATP) channels [9,13,14,31,32] that can be closed by ATP [9,31]. Glucose increases intracellular free ATP in α cells [8,10], although reports vary as to the ability of glucose to inhibit α-cell KATP channels [13,31,33]. Evidence from SUR1−/− mice implicate KATP channels as regulators of glucagon release [13,34,35]. However, because both α and β cells possess molecularly identical KATP channels [36,37], it is not clear how KATP-mediated depolarisation would stimulate insulin secretion but suppress glucagon secretion.
The answer may lie in the downstream machinery regulating electrical activity and Ca2+ entry. Unlike β cells, α cells possess a large voltage-dependent Na+ current that is essential for glucagon release [14,33]. We have previously proposed that the depolarisation-induced inactivation of this channel contributes to the cessation of action potential firing [14]. Additionally, activation of voltage-dependent Ca2+ channels (VDCCs) is essential for Ca2+ entry and α-cell function [38,39]. Accordingly, the α-cell intracellular Ca2+ ([Ca2+]i) oscillations (reflecting α-cell electrical activity) are suppressed in parallel with glucagon release by glucose [40]. Multiple VDCCs regulate glucagon release [14,41–43], although the N-type channels appear to be particularly important for glucagon release evoked by hypoglycaemia alone [33,44,45], at least in mouse islets. This is in contrast to the β cell in which the L-type VDCC functionally predominates [46].
We have now compared insulin and glucagon release and α- and β-cell Ca2+ responses in intact mouse, rat, and human pancreatic islets. We show that glucose retained the ability to suppress glucagon release from isolated islets during blockade of the Zn2+ and GABA paracrine pathways, and in the absence of stimulated insulin secretion or β-cell Ca2+ responses. Thus we now provide evidence in both rodent and human islets supporting the direct (intrinsic) glucose regulation of glucagon release from pancreatic α cells.
Results
Glucose Can Regulate Glucagon Secretion Directly
To examine a role for GABA and Zn2+ as paracrine mediators of glucagon secretion, we examined the ability of the GABAA receptor antagonist SR-95531 and Zn2+ chelation with Ca2+-EDTA to prevent the glucose-dependent suppression of glucagon release. Glucose, at concentrations (7 and 8.3 mM) just above the threshold for insulin release (see below), suppressed glucagon secretion from isolated mouse (Figure 1A) and rat islets (Figure 1B) by 60% (n = 15, p < 0.001) and 57% (n = 10, p < 0.001), respectively. In both mouse and rat islets, the ability of glucose to inhibit glucagon secretion persisted in the presence of Ca2+-EDTA (43% and 48%, n = 10, p < 0.001, respectively) and SR-95531 (31% and 46%, n = 8 and 10, p < 0.01 and p < 0.001, respectively) (Figure 1).
It is worth noting that in the presence of the GABAA antagonist, glucagon secretion was increased under both low- and high-glucose conditions, and furthermore, glucose was approximately 50% less effective in suppressing glucagon release from mouse islets (31% versus 60%, respectively) (Figure 1A). Additionally, somatostatin released from pancreatic δ cells is suggested to be a potential paracrine regulator of glucagon secretion. However, the somatostatin receptor 2 (SSTR-2) antagonist PRL-2903 does not interfere with the ability of glucose (at 3 and 7 mM) to inhibit glucagon secretion from mouse islets [47]. Like the GABAA receptor antagonist, however, PRL-2903 increased glucagon secretion in low-glucose conditions [47]. Thus, although the present data do not entirely rule out these pathways as modulators of glucagon secretion, glucose is clearly able to suppress glucagon secretion independently of these.
α-Cell KATP Channels Regulate Glucagon Secretion
We next examined glucagon and insulin secretion during pharmacological manipulation of KATP channel activity. Here we have examined the role of KATP channels in intact islets by applying an indirect, but minimally invasive, technique. It would have been difficult (if not impossible) to study the effects of glucose on α-cell KATP channel activity using the patch-clamp technique because of the smallness of the α-cell resting conductance (0.15 nS/pF in the absence of glucose, of which two thirds is attributable to KATP channels) [13]. We have instead used increasing concentrations of diazoxide and tolbutamide to “titrate” the influence of KATP channel activity on α-cell [Ca2+]i and glucagon secretion. Increasing concentrations of the KATP channel activator diazoxide demonstrated that moderate activation of KATP channels (0.3–10 μM diazoxide) relieved the suppression of glucagon secretion from both mouse (Figure 2A) and rat (Figure 2B) islets. Stimulation of glucagon release was half-maximal at approximately 1 μM diazoxide, which is well below that required to inhibit insulin release, suggesting that “re-activation” of glucagon release was not secondary to reduced β-cell secretion. Increasing the concentration of diazoxide beyond 10 μM inhibited glucagon secretion in parallel with an inhibition of insulin release from both mouse and rat islets (Figure 2A and 2B). When instead applied in the presence of 1 mM glucose (at which concentration glucagon secretion is stimulated), increasing diazoxide produced a monotonic inhibition of glucagon secretion (Figure 2C) with a half-maximal inhibitory concentration (IC50) that is much lower than what is seen under high-glucose conditions (∼2 μM versus ∼50 μM).
In the complete absence of glucose, when KATP channels are expected to be open, the KATP channel antagonist tolbutamide also produced a biphasic effect on glucagon release. Augmentation of glucagon secretion was seen at tolbutamide concentrations of up to 1 μM (stimulation being half-maximal at 0.1 μM), whereas inhibition was observed at higher concentrations (Figure 2D). Importantly, in the presence of a maximally inhibitory tolbutamide concentration (0.5 mM), glucose was unable to produce any further inhibition of glucagon release (Figure 3A). These data are inconsistent with the idea that β-cell secretion is the primary inhibitor of α-cell glucagon release. They also suggest that glucagon secretion is maximal within a “window” of intermediate KATP channel activity.
To further investigate the role of α-cell KATP channels as regulators of glucagon secretion, we studied islets from Kir6.2Y12X mice, which posses a Tyr12STOP mutation in the Kcnj11 gene, leading to premature termination of the KATP channel pore-forming subunit [48]. At low-glucose concentrations, glucagon secretion from the Kir6.2Y12X islets was already suppressed compared with that from wild-type islets (Figure 3B), similar to the effect of glucose stimulation or pharmacological KATP channel inhibition observed in Figures 2D and 3A. Consistent with a recent report [49], higher glucose levels stimulated glucagon secretion, perhaps due to a direct effect of metabolism on secretion [33]. In wild-type islets, glucagon secretion exhibited a nadir at 5 mM glucose, and glucagon secretion at 20 mM glucose was 40% higher than at 5 mM glucose. Insulin secretion in the Kir6.2Y12X islets was elevated at low-glucose concentrations (Figure 3C), whereas glucose-stimulated insulin secretion was blunted. It is notable that the increase in glucagon release from the Kir6.2Y12X islets was coincident with increased insulin secretion, reinforcing the view that inhibition of glucagon is not mediated by a factor released by the β cells. Furthermore, in control islets, the inhibition of glucagon secretion is maximal at a concentration of glucose (5 mM) that is without stimulatory action on insulin secretion.
We next examined the function of α and β cells in situ by monitoring the [Ca2+]i responses of single cells within intact mouse islets. These were functionally identified by their [Ca2+]i response to 0.5, 2, and 11 mM glucose [50]. Glucose stimulation suppressed the [Ca2+]i response of α cells by 51 ± 3% (n = 63 cells in 15 islets, p < 0.001) (Figure 4A and 4B). The suppressive effect of glucose was alleviated (to 101 ± 11% of initial values) by application of 2 μM diazoxide (Figure 4A and 4B). This concentration of diazoxide was similar to that which produced maximal stimulation of glucagon secretion and much (∼15-fold) lower than necessary to inhibit insulin secretion (Figure 2B) or β-cell [Ca2+]i responses to glucose (Figure 4A).
KATP Channels Regulate Human α-Cell Activity
Little work has been done to examine the mechanism of glucose-regulated glucagon secretion from isolated human islets [51]. Glucagon and insulin secretion from islets isolated from healthy donors was examined. Similar to above, glucagon secretion from human islets was suppressed by 58 ± 3% (n = 9, p < 0.05) upon raising the glucose concentration from 0 to 10 mM. Tolbutamide (200 μM) inhibited glucagon release by 43 ± 14% (n = 7, p < 0.05). At the same time, insulin secretion was increased 6.4- and 4.4-fold by glucose and tolbutamide stimulation, respectively (Figure 5A). Moderate re-activation of KATP channels with 2 μM diazoxide antagonized the glucose-induced suppression of glucagon release (n = 7, p < 0.01) (Figure 5B). A 10-fold higher concentration of diazoxide had significantly less stimulatory effect (p < 0.001 versus 2 μM diazoxide), and in the presence of 200 μM diazoxide, glucagon secretion was not different from that observed in the absence of the KATP channel activator. Importantly, the lowest concentration of diazoxide (2 μM), which produced the strongest stimulation of glucagon secretion, had no inhibitory effect on glucose-stimulated insulin secretion (n = 5), whereas 20 and 200 μM produced partial or complete inhibition of insulin secretion. The GABAA receptor antagonist SR95531 had no effect on glucose-induced inhibition of glucagon secretion from human islets, and whole-cell voltage-clamp measurements indicate that human α cells, unlike human β and δ cells, express few if any GABAA receptor Cl− channels (M. Braun, R. Ramracheya, and P. Rorsman, unpublished data).
Examination of the Ca2+ responses of single α and β cells within intact human islets demonstrated that glucose decreased [Ca2+]i by 62 ± 3% (n = 42 cells in 7 islets, p < 0.001) in α cells, and that this effect could be completely reversed by 2 μM diazoxide (Figure 6A and 6B). Furthermore, the re-activation of α-cell [Ca2+]i responses by diazoxide could be prevented by application of 100 μM tolbutamide, confirming the role of KATP channels (Figure 6C). An analysis of the dose–response relationship to diazoxide demonstrated a maximally effective concentration of 1.7 μM (Figure 6D and 6E; n = 38 cells in 9 islets), far below the concentration necessary to block β-cell responses (Figure 6D). Increases in diazoxide above 20 μM blocked the [Ca2+]i responses of α cells and β cells in parallel, consistent with the effect on glucagon and insulin release.
Na+ Channels Regulate Low-Glucose–Stimulated Glucagon Secretion
The above data suggest that glucose-dependent inhibition of α-cell KATP channels is involved in the suppression of glucagon secretion. It is not clear however, how KATP channel inhibition and membrane depolarisation result in suppression of secretion. Unlike in β cells, α-cell Na+ channels are active in the physiological range of membrane potentials. Previous work from our group suggested that the voltage-dependent inactivation of Na+ channels, which in mouse α cells is half-maximal at −42 mV [43], contributes to cessation of electrical activity upon α-cell depolarisation [14]. We thus used the voltage-dependent Na+ channel antagonist tetrodotoxin (TTX) to test a role for these channels in α-cell function and glucagon secretion. TTX (0.1 μg/ml) suppressed glucagon release from mouse islets by 56 ± 5% (n = 10, p < 0.001) under low-glucose conditions (Figure 7A). This effect of TTX in the low-glucose condition was similar to what we observed with high-glucose stimulation alone. With TTX present, glucose was without further inhibitory action, suggesting that glucose inhibits glucagon secretion through a Na+ channel-dependent mechanism. TTX has no effect on glucose-stimulated insulin secretion in mouse islets (Figure 7A); again suggestive of a direct rather than indirect effect on α cells. In accordance with this, application of 0.1 μg/ml TTX to mouse islets reversibly abolished the α-cell [Ca2+]i response evoked by low-glucose concentrations (Figure 7B and 7C; n = 18 cells in 4 islets), but had no effect on β-cell [Ca2+]i (unpublished data).
N-Type Ca2+ Channels Mediate the Ca2+ Influx That Triggers Glucagon Secretion and α-Cell Exocytosis
Downstream of Na+ channel activation, the opening of VDCCs allows Ca2+ into α cells, triggering the exocytosis of glucagon-containing vesicles. We applied whole-cell patch-clamp recordings to establish the α-cell Ca2+ channel complement. The integrated whole-cell Ca2+ current measured during 50-ms depolarisations from −70 mV to 0 mV amounted to 6.2 ± 0.8 pC (n = 15) under control conditions, 2.6 ± 0.8 pC (n = 10; p < 0.05) in the presence of 50 μM nifedipine, and 4.1 ± 0.5 pC (n = 12; p < 0.01) in the presence of 1 μM ω-conotoxin. Thus, L- and N-type Ca2+ channels account for 58% and 34% of the α-cell Ca2+ current, respectively.
Figure 8A compares glucagon secretion at 1 and 20 mM glucose under control conditions and in the presence of 100 nM ω-conotoxin and 20 μM nifedipine, respectively. We found that whereas the L-type channel blocker nifedipine had no effect on glucagon release at 1 or 20 mM glucose, the N-type Ca2+ channel blocker ω-conotoxin inhibited the release of the hormone to a level similar to that of high glucose, tolbutamide, and TTX. Furthermore, glucose exerted no additional inhibitory action in the presence of ω-conotoxin. Thus, glucagon secretion stimulated by low-glucose levels depends principally on Ca2+ influx through N-type channels, although these only account for one third of the Ca2+ current. We confirmed this Ca2+ channel dependence by conducting high-resolution single-cell capacitance measurements of exocytosis (Figure 8B). Exocytosis elicited by 500-ms step depolarisations from −70 to 0 mV averaged approximately 150 fF (corresponding to the fusion of ∼75 secretory granules with the plasma membrane [43]) under control conditions (Ctrl). This response was not significantly affected by 50 μM nifedipine (nif), but was nearly abolished by 1 μM ω-conotoxin (ω-con; Figure 8C).
Inactivation of N-Type Ca2+ Channels Underlie Glucose Inhibition of Glucagon Secretion
We next examined the mechanism by which N-type channels regulate glucagon release in response to membrane depolarisation/hyperpolarisation in mouse α cells. Non–L-type (isradipine-resistant) Ca2+ currents were elicited by step depolarization to 0 mV in the absence (black) or presence (red) of 1 μM ω-conotoxin following a 200-ms conditioning pulse to −70 mV (Figure 9A) or +10 mV (Figure 9B). It is evident that the ω-conotoxin–sensitive component was abolished by the +10 mV conditioning pulse. Figure 9C summarizes the relationship between the conditioning voltage and the Ca2+ current amplitude in the absence (open squares) and presence (circles) of ω-conotoxin (upper panel). The ω-conotoxin–sensitive N-type Ca2+ current component is shown in the lower panel of Figure 9C, and underwent voltage-dependent inactivation that was half-maximal (V0.5) at −31 ± 6 mV (n = 6). A residual, ω-conotoxin–insensitive Ca2+ current also appears to undergo voltage-dependent inactivation (Figure 9C). This accounts for less than 15% of the inactivating current, and may be attributable to T-type Ca2+ channels that have been reported in α cells [14].
In Figure 9D, we examined the voltage-dependent activation of α-cell exocytosis using step-wise depolarisations (500 ms) from −70 mV to between −50 and 20 mV (Figure 9D). The capacitance response versus voltage relationship (Figure 9E) demonstrates a marked increase in the capacitance response between −10 and 10 mV. Thus any reduction in action potential amplitude within this range would severely attenuate α-cell exocytosis.
The voltage-dependent inactivation of α-cell exocytosis was examined next (Figure 9F). Cells were held at conditioning potentials between −70 and −30 mV, after which exocytosis was elicited by depolarisation to 0 mV. Exocytosis stimulated from the −30 mV conditioning potential was only 25% of that elicited from −70 mV (Figure 9G). This voltage-dependent decline in exocytosis is attributable to the voltage-dependent inactivation of the N-type Ca2+ channels. The fact that inhibition of exocytosis appears to occur at more-negative voltages than that documented for the inactivation of the N-type Ca2+ channels can be attributed to the brevity of the conditioning pulses in Figure 9A–9C (200 ms), whereas in Figure 9G the holding potential was varied.
Discussion
Regulated glucagon secretion from pancreatic α cells is a major component of the counter-regulatory response to hypoglycaemia. Diabetes mellitus is associated with defects of glucagon secretion that exacerbate the consequences of impaired insulin secretion [2–4]. The mechanism regulating the release of this important hormone is incompletely understood and currently the source of much debate. In the present study we examined glucagon secretion and the [Ca2+]i response of in situ α cells of isolated rodent and human islets. We further characterised the Ca2+ currents and capacitance changes of single isolated α cells, to dissect the mechanism regulating downstream control of glucagon release.
Previous work with genetic mouse models, including SUR1−/− [13,34,35] and Kir6.2−/− [16] mice, suggests an important role for KATP channels in glucagon secretion. Three pieces of evidence argue for the importance of islet KATP channels in glucagon secretion. First, introduction of the Tyr12STOP mutation into the KATP channel subunits in the Kir6.2Y12X islets results in suppression of glucagon secretion at low-glucose levels and causes loss of glucose-induced inhibition of secretion. Second, the KATP channel inhibitor tolbutamide when applied at maximally effective concentrations inhibits glucagon secretion from isolated islets. Third, a low dose of the KATP channel activator diazoxide restores α-cell Ca2+ responses and glucagon secretion in high-glucose conditions. What is not immediately clear from these arguments, and from the previous SUR1−/− genetic studies, is whether glucagon is being controlled by the α-cell or the β-cell KATP channels, as the latter may regulate α-cell function indirectly via paracrine pathways. We therefore examined insulin and glucagon secretion, and the Ca2+ responses of α and β cells in situ in parallel to determine the functional relationship between these cells. Furthermore, we measured the glucagon response to glucose during blockade of putative paracrine signalling pathways.
Studies on rat islets support an important role for paracrine signals as regulators of glucagon release [33,52,53], whereas the case for paracrine regulation of glucagon secretion from mouse and human α cells is less clear [9,10,14,17–21,27]. Although it is clear that Zn2+, GABA, and somatostatin can exert a paracrine control of glucagon secretion under certain conditions, the data shown here firmly establish that glucose can suppress glucagon secretion independently of these pathways as demonstrated in Figure 1 (and in [47]). Furthermore, and in agreement with recently published results [47], maximal inhibition of glucagon release occurs at levels equal to or lower than 5 mM glucose, whereas the stimulation of insulin release requires glucose levels greater than 5 mM (Figure 3), suggesting that products of β-cell secretion are not required for suppression of glucagon release.
This conclusion is further underpinned by the significant discordance between insulin and glucagon release, and the β- and α-cell Ca2+ responses, under several conditions: (1) low doses of the KATP channel opener diazoxide that stimulate glucagon secretion while not affecting insulin release; (2) high doses of diazoxide at which suppression of β-cell secretion would be expected to elevate glucagon release; (3) in the Kir6.2Y12X islets; and (4) in response to the Na+ channel blocker TTX. Therefore, our data argue that glucagon-producing α cells possess an intrinsic mechanism for regulation by glucose and that involves KATP channels. This is at variance with the data of Liu et al. [12] who report that tolbutamide has no effect on [Ca2+]i in single, isolated α cells, but in agreement with the conclusion of Ostenson et al. [54], that sulfonylureas can inhibit glucagon secretion by a direct, non-paracrine mechanism.
On the basis of our findings, we propose that α-cell glucagon secretion occurs within a narrow window of intermediate KATP channel activity (and thus membrane potential) (Figure 10). That is, if the α cell is either too hyperpolarised (maximal KATP activity) or too depolarised (maximal KATP inhibition), then glucagon secretion is suppressed. This is supported by the biphasic effects of both diazoxide and tolbutamide. Whereas the former (in high glucose) brings the α cell in a dose-dependent manner through the membrane potential window supporting glucagon secretion in the depolarised (low KATP) to hyperpolarised (high KATP) direction (Figure 10C), the latter (in zero glucose) brings the α cell through the window in the opposite direction (from hyperpolarised to depolarised) (Figure 10B). Thus, glucose likely leads to the suppression of glucagon secretion by depolarising the α-cell membrane potential above the range that supports glucagon secretion (Figure 10A). Indeed, in some (but not all, see [55]) studies, glucose was found to depolarize mouse and rat α cells and reduce action potential amplitudes [13,52]. It is interesting that whereas low concentrations of glucose were without stimulatory effect, tolbutamide (0.1–1 μM) stimulated glucagon release beyond that observed at zero glucose. Thus, small depolarisations (as previously documented for arginine [13]) exert a positive “chronotropic” effect in α cells and thus stimulate glucagon release. The fact that glucose did not share this ability indicates that the depolarisation produced by 1 mM glucose results in sufficient inactivation of the currents to balance any increase in action potential frequency.
Previous human studies have employed pharmacological KATP channel antagonists [23,24] or agonists [25,26] to examine the regulation of glucagon release in vivo. These were interpreted with the assumption that pharmacological KATP modulation only affects the β cells, and that any change in glucagon release was therefore secondary to altered β-cell function. Our data establish that KATP channel modulation has dramatic and direct effects on glucagon secretion and Ca2+ signalling in human α cells under conditions in which insulin secretion is unaffected. Thus, the in vivo manipulation of α-cell KATP channel activity in the above studies may well have involved direct effects on α-cell function that contributed to the observed changes in glucagon release.
The importance of the human KATP channel pathway for glucagon release is nicely highlighted by a study investigating the effects of the common Glu23Lys (E23K) polymorphism of the Kir6.2 subunit of the KATP channel on insulin and glucagon secretion in non-diabetic human patients [29]. This variant of the channel leads to a slight decrease in the ATP sensitivity of the channel. The functional significance of this was examined by comparing hormone release during hyperglycaemic clamps in individuals carrying the polymorphism or not. Although insulin secretion in homozygous Glu23Lys individuals was not different from controls, glucose-induced suppression of glucagon release was blunted [29]. This becomes understandable in light of the effect of diazoxide on isolated human islets (Figures 4 and 5). Half-maximal activation of KATP channels occurs at diazoxide concentrations of 20–100 μM (depending on the intracellular ATP level) [56]. If the Glu23Lys polymorphism increases KATP channel activity to the same extent as 0.3–1.5 μM diazoxide, the concentration at which an effect on glucagon release is first seen, then the effect will be very difficult to detect with electrophysiology, perhaps explaining why some studies have failed to detect a functional effect of the polymorphism (reviewed in [57]). Nevertheless, such small changes can have significant biological effects, as illustrated by the glucagon release data, and may contribute to pathological states such as impaired glucose tolerance and diabetes. Thus, the reduced ability of glucose to inhibit glucagon secretion in individuals carrying the Glu23Lys variant of the KATP channel likely results from the failure of these channels to undergo complete inhibition in response to glucose.
We have proposed that glucagon secretion is stimulated within a window of intermediate α-cell KATP channel activity, and that glucagon release is suppressed by either increases or decreases in KATP channel activity. How is this accomplished? Briefly, we suggest that this window is the result of (1) the ability of intermediate KATP channel activity to support regenerative electrical responses through the activation of voltage-dependent Na+ and N-type Ca2+ channels (grey in Figure 10); (2) the failure of the Na+ and Ca2+ channels to activate when α cells are hyperpolarised by the opening of a major fraction of KATP channels (above the grey in Figure 10); and (3) the voltage-dependent inactivation of the Na+ [14] and N-type Ca2+ channels when KATP channels are closed and the α-cell membrane potential is depolarised (below the grey in Figure 10). Thus, the differential responsiveness of α and β cells to diazoxide does not result from differential sensitivities of the KATP channels in these cells, but to the downstream responses to titrated KATP channel activity.
One important difference between mouse α and β cells is that whereas the latter rely exclusively on voltage-gated Ca2+ channels for the upstroke of the action potentials, glucagon-producing α cells are equipped with voltage-gated Na+ channels. These channels undergo voltage-dependent inactivation at voltages more positive than −50 mV [14]. This will reduce the action potential amplitude, and indeed it is reported that glucose reduces the peak voltage in α cells from +11 mV to −1 mV [13]. This will in itself result in an approximately 35% reduction of exocytosis, which is steeply dependent on voltage between −10 and +10 mV (Figure 9E). The functional significance of this is illustrated by the observations that glucagon secretion and α-cell Ca2+ responses at low-glucose concentrations are suppressed by the Na+ channel blocker TTX. The exact ion channel complement of human α cells remains to be established. However, the fact that glucagon secretion from human islets shows the same bell-shaped diazoxide concentration dependence as in mouse islets suggests that the depolarization-induced inactivation of ion channels involved in α-cell regenerative electrical activity underlies glucose-induced suppression of glucagon secretion in man as well.
Although L-type VDCCs mediate the majority of Ca2+ current in α cells, this work and previous studies [33,44,45] demonstrate that it is the N-type (ω-conotoxin–sensitive) VDCCs that mediate the Ca2+ influx necessary for α-cell exocytosis and glucagon secretion under hypoglycaemic conditions. We now show that the α-cell N-type VDCCs are also subject to voltage-dependent inactivation at voltages more positive than −50 mV and furthermore that this is associated with reduced exocytotic capacity. It is pertinent that N-type VDCC-deficient mice exhibit reduced serum glucagon levels and improved glucose tolerance despite a parallel reduction in plasma insulin [44]. Although Ca2+ influx through L-type Ca2+ channels does not appear to contribute much to glucagon secretion under the experimental conditions used in this study, these channels become the predominant conduit of Ca2+ entry in the presence of agents increasing cAMP [41] (unpublished data). The mechanism underlying this switch in Ca2+ channel dependence remains obscure, but may depend on the strength of depolarisation because glucagon secretion stimulated by strong depolarisation with increased K+ in combination with KATP channel block can be prevented by inhibition of L-type Ca2+ channels [47].
We finally point out that the model we propose here for the control of glucagon secretion shares many features with what is known about the metabolic control of insulin secretion. This in turn means that processes that interfere with the ability of, for example, glucose to stimulate insulin secretion will have the opposite effect on glucagon secretion. This would provide a simple explanation for the fact that both insulin and glucagon secretion become perturbed in diabetes and why oversecretion of glucagon exacerbates the hyperglycaemic effects of insufficient insulin secretion.
Materials and Methods
Islet isolation and culture.
Islets from female NMRI mice were isolated by collagenase digestion and cultured in RPMI-1640 media (5 mM glucose) at 37 °C and 5% CO2 for 2−24 h prior to secretion or intracellular Ca2+ assays. For single-cell studies, islets were dispersed in a Ca2+-free buffer and plated in 35-mm plastic dishes. Generation of the Kir6.2Y12X mice, which possess a Tyr12STOP mutation in the Kcnj11 gene on a BALB/c background, has been described previously [48]. This results in nonfunctional KATP channels in both α and β cells. These animals are not overtly diabetic, exhibiting normal fasting glucose, but are glucose intolerant (R. Cox, unpublished data). For these experiments, age- and weight-matched wild-type C3HB mice were used as controls because the Kir6.2Y12X mice were back-crossed into this background over several generations. Human islets from four healthy donors were obtained from the Oxford DRWF Human Islet Isolation Facility. Experiments were performed in duplicate or triplicate using islets from each donor. Human islets were cultured in RPMI-1640 (10 mM glucose) at 37 °C and 5% CO2, and experiments were performed within 48 h of isolation.
Intracellular Ca2+ measurements.
Intact islets were loaded with fluo-4-AM (1 μM) and fura red (5 μM) in 0.5 mM glucose buffer (see below) with 0.01% pluronic acid for 30 min at 37 °C. Islets were fixed with a wide-bore holding pipette within a continuously superfused and temperature-controlled (37 °C) bath on an Axioskop 2 FS-mot microscope (Carl Zeiss, http://www.zeiss.com). The perfusion buffer contained (in mM): 140 NaCl; 3.6 KCl; 2 NaHCO3; 0.5 NaH2PO4; 0.5 MgSO4; 5 HEPES; 2.5 CaCl2; 0.5, 3, or 11 glucose; (pH 7.4 with NaOH). Laser scanning confocal microscopy was performed using an LSM 510meta system (Zeiss). Excitation was with a 488-nm argon laser, and emitted fluorescence was collected through 500–550-nm and 650–710-nm band-pass filters for the fluo-4 and fura red (FuraR) signals, respectively. Increases in intracellular Ca2+ are displayed as upward deflections. Images were acquired at 1.5-s intervals. Individual cells were selected as regions of interest, and the average ratio intensity (RI = [Fluo+1]/[FuraR+20] × 128+1) of these were analysed over time with Origin v7.0220 (OriginLab Corporation, http://www.originlab.com). Prior to experimental recordings, islets were perfused for 10 min with the appropriate control solution, and Ca2+ responses were monitored periodically during this time to ensure that responses (or lack thereof) were stable prior to beginning the experimental recording. Ca2+ responses were determined by baseline subtraction and calculation of the integrated response (i.e., the area under the curve). The slope of this response, reported here as arbitrary units (AU), was calculated for the final 60%–90% of a given treatment period to allow for equilibration of the responses.
Hormone release.
Insulin and glucagon secretion were measured as described elsewhere [45]. Briefly, batches of ten freshly isolated islets were pre-incubated in 1 ml of Krebs–Ringer buffer (KRB) supplemented with 1 mM glucose for 30 min (rodent) or 1 h (human) followed by 1-h incubation in 1 ml of test KRB medium supplemented as indicated. For experiments in which Zn2+ was chelated or GABAA receptors were antagonised (Figure 1), the Ca2+-EDTA and SR-95531, respectively, were present in both the pre-incubation and test solutions. When extracellular KCl was increased, NaCl was correspondingly reduced to maintain iso-osmolarity.
Single-cell capacitance and ion current measurements.
Whole-cell currents and exocytosis were recorded using an EPC-9 patch-clamp amplifier (HEKA Electronics, http://www.heka.com) and Pulse software (version 8.50). Single α cells were identified by their small size and Na+ current inactivation properties [58]. This method was validated by combining the electrophysiological recordings with subsequent immunostaining for glucagon and insulin. The extracellular medium contained (in mM): 118 NaCl; 20 TEA-Cl (tetraethylammonium chloride); 5.6 KCl; 2.6 CaCl2; 1.2 MgCl2; 5 HEPES (pH 7.4 with NaOH); and 5 glucose. Except for Figure 8B and 8C, in which the standard whole-cell configuration was used, the electrophysiological measurements were conducted using the perforated patch technique, and the pipette solution contained (in mM): 76 Cs2SO4; 10 NaCl; 10 KCl; 1 MgCl2; and 5 HEPES (pH 7.35 with CsOH). Exocytosis was monitored as changes in α-cell capacitance using the software-based lock-in function of the Pulse software. The standard whole-cell measurements (Figure 8B and 8C) were conducted using a pipette solution (dialyzing the cell interior) consisted of (in mM) 125 CsOH; 125 glutamate; 10 CsCl; 10 NaCl; 1 MgCl2; 5 HEPES (pH 7.15 with KOH); 3 Mg-ATP; and 25 μmol/l EGTA (measured resting free Ca2+, ∼0.2 μmol/l). Pulses were applied at low frequency (<0.05 Hz) to allow the exocytotic capacity to recover fully between the pulses.
Statistical analysis.
Data are presented as means and standard errors. Significance was examined by either the unpaired t-test or by multiple-comparisons analysis of variance (ANOVA) and post-test, as appropriate.
Acknowledgements
PEM was supported initially the European Foundation for the Study of Diabetes (EFSD)/AstraZeneca Fellowship in Islet Biology and is currently an Alberta Heritage Foundation for Medical Research Scholar, Canadian Diabetes Association Scholar, and the Canada Research Chair in Islet Biology. PR is a Wolfson Royal Society Merit Award Research Fellow.
Abbreviations
[Ca2+]i - intracellular Ca2+
FuraR - fura red
GABA - γ-aminobutyric acid
KATP - ATP-dependent K+, TTX, tetrodotoxin
VDCC - voltage-dependent Ca2+ channel
Figures and Tables
Figure 1
Glucose Suppresses Glucagon Release Independently of Paracrine Signals Mediated by Zn2+ or GABA.
(A) Glucagon release from isolated mouse islets was suppressed by 60% at 7 mM glucose compared with 1 mM (filled bars). Glucose retained its suppressive effect on glucagon release under conditions of Zn2+ chelation (Ca2+-EDTA) (open bars) and antagonism of GABAA receptors (SR-95531) (shaded bars). Antagonism of GABAA receptors increased both basal and glucose-suppressed glucagon secretion, suggesting a paracrine role for GABA independent of the glucose effect.
(B) As in (A), but using rat islets.
*, p < 0.05; **, p < 0.01; ***, p < 0.001, compared with 1 mM glucose, or as indicated.
Figure 2
Glucagon Release from Isolated and Intact Mouse Islets Is Regulated by a KATP Channel-Dependent Pathway
(A) Glucagon (filled circles) and insulin (open circles) secretion measured from mouse islets in the presence of 8.3 mM glucose at increasing concentrations of diazoxide.
(B) As in (A), but using rat islets. The glucagon responses to 1 mM glucose (filled square) and 100 μM tolbutamide (filled triangle) are indicated.
(C) As in (A), but in the presence of 1 mM glucose and only measuring glucagon secretion.
(D) As in (A), but examining the effect of tolbutamide in the absence of glucose. Glucagon secretion in response to 20 mM glucose is indicated by the filled square.
*, p < 0.05; **, p < 0.01; ***, p < 0.001, compared with zero diazoxide (A–C) or zero tolbutamide (D).
Figure 3
Tolbutamide and Glucose Effects Are Non-Additive and Glucagon Response Is Altered in Kir6.2Y12X Islets That Express a Truncated Kir6.2 Subunit
(A) Glucagon release from isolated mouse islets at 1 (open bars) and 20 mM glucose (filled bars) under control conditions and presence of 0.5 mM tolbutamide.
(B) A glucose dose-response of glucagon release from control C3HB islets (filled circles) and Kir6.2Y12X islets (open circles).
(C) As in (B), but insulin was measured.
*, p < 0.05; **, p < 0.01; ***, p < 0.001, compared with control.
Figure 4
The Intracellular Ca2+ Response of Single α Cells within Intact Islets Can Be Re-Activated by Low Concentrations of Diazoxide
(A) Representative intracellular Ca2+ responses from α and β cells within an intact mouse islet exposed to 0.5 mM glucose, 11 mM glucose, and 11 mM glucose plus 2 μM diazoxide (diaz) as indicated above the traces.
(B) The Ca2+ response of α cells was suppressed by 11 mM glucose, and could be reactivated with low concentrations of the KATP channel agonist diazoxide. ***, p < 0.001, compared with the low-glucose condition, or as indicated.
Figure 5
Glucagon Release from Isolated and Intact Human Islets Is Regulated by a KATP Channel-Dependent Pathway
(A) Glucagon (open bars) and insulin (filled bars) release was measured from isolated human islets under control conditions and following addition of 10 mM glucose or 200 μM tolbutamide.
(B) Glucagon (open bars) and insulin secretion (filled bars) measured in the presence of 10 mM glucose and increasing concentrations of diazoxide (0–200 μM).
*, p < 0.05; **, p < 0.01; ***, p < 0.001, compared with controls, unless otherwise indicated.
Figure 6
Intracellular Ca2+ Responses of α Cells within Intact Human Islets Are Regulated by a KATP Channel-Dependent Mechanism
(A) Ca2+ responses measured in two human α cells within the same islet at 0.5 mM and 11 mM glucose (glu), in the presence of 2 μM diazoxide (diaz).
(B) Summary of the Ca2+ responses at 0.5 mM glucose, at 11 mM glucose, in the presence of glucose (11 mM) and diazoxide (2 μM), and following the removal of diazoxide, but in the continued presence of 11 mM glucose.
(C) The re-activation of human α-cell Ca2+ responses by 2 μM diazoxide was reversed upon application of the KATP channel antagonist tolbutamide (100 μM).
(D) The effects of increasing concentrations of diazoxide on the Ca2+ response of a human α cell and β cell within the same islet exposed to 11 mM glucose. At the end of the experiment, diazoxide was withdrawn and glucose lowered to 0.5 mM.
(E) Dose-response curve for the effect of diazoxide on α-cell Ca2+ responses. The grey horizontal line indicates the response with 11 mM glucose alone.
*, p < 0.05; **, p < 0.01; ***, p < 0.001, compared with controls, unless otherwise indicated.
Figure 7
Glucagon Secretion and α-Cell Ca2+ Responses Are Dependent upon the Activity of Voltage-Dependent Na+ Channels
(A) Glucagon (open bars) and insulin (filled bars) release from isolated mouse islets at 1 mM and 20 mM glucose, under control conditions and in the presence of the Na+ channel antagonist TTX (0.1 μg/ml).
(B) Intracellular Ca2+ response of single α cells to 0.5 mM glucose. TTX (0.1 μg/ml) was included in the perfusion medium during the indicated period.
(C) The effects of TTX and glucose on α-cell Ca2+ responses. Note that TTX inhibits Ca2+ responses as effectively as glucose and that the action is at least partially reversible.
*, p < 0.05; **, p < 0.01; ***, p < 0.001, compared with controls, unless otherwise indicated.
Figure 8
Glucagon Secretion Is Regulated by N-Type Ca2+ Channels
(A) Glucagon release measured at 1 (open bars) and 20 mM glucose (filled bars) under control conditions and in the presence of 100 nM ω-conotoxin (middle) or 20 μM nifedipine (right).
(B) Exocytosis was elicited by 500-ms depolarisations from −70 to 0 mV under control conditions (Ctrl) and in the presence of either nifedipine (50 μM; nif) or ω-conotoxin (1 μM; ω-con).
(C) Summary of the exocytotic response under control conditions and in the presence of nifedipine and ω-conotoxin.
*, p < 0.05; **, p < 0.01, compared with 1 mM glucose and the control capacitance response, unless indicated otherwise.
Figure 9
Voltage-Dependent Inactivation of α-Cell N-Type Ca2+ Currents and Exocytosis
(A) N-type Ca2+ currents were evaluated during blockade of the L-type channels with isradipine (2 μM). The N-type channel antagonist ω-conotoxin (1 μM; red traces) reduced the Ca2+ current elicited by a step depolarisation from −70 to 0 mV (right).
(B) As in (A), but the pulse to 0 mV was preceded by a 200-ms conditioning depolarization to +10 mV. ω-Conotoxin was without effect on the current measured during the depolarization to 0 mV under these conditions (right).
(C) Top: peak Ca2+ currents measured in the presence of 2 μM isradipine alone (open squares) or together with 1 μM ω-conotoxin (red circles) during a depolarization to 0 mV following 200-ms conditioning pulses to between −70 and +70 mV. Lower: inactivation of the ω-conotoxin–sensitive component. Half-maximal inactivation of the N-type current was at −31 ± 6 mV (n = 5).
(D) Exocytosis was elicited with 500-ms depolarisations from −70 mV to between −50 and 20 mV.
(E) The voltage dependence of the exocytotic response.
(F) Exocytosis elicited by 500-ms depolarisations to 0 mV from holding potentials of between −70 and −30 mV.
(G) Summary of effects of holding potential on exocytotic responses elicited by depolarisations to 0 mV. Data have been normalized to responses obtained using a holding potential of −70 mV.
*, p < 0.05; **, p < 0.01; ***, p < 0.001, compared with ω-conotoxin (C, top) or with the initial response.
Figure 10
A Model for the Suppression of Glucagon Secretion by an Intrinsic α-Cell Pathway
Schematic representation of the effects of glucose, tolbutamide, and diazoxide on α-cell KATP, Na+, and N-type Ca2+ (VDCC) channel activities and glucagon secretion is shown. The insulin response is also shown for comparison with our experimental results (dashed lines, lower panels). The grey gradient represents a “window” of α-cell KATP channel activity that supports the activation of Ca2+ and Na+ channels. Above this window, the cell is hyperpolarized and Ca2+ and Na+ channel activation is prevented, whereas KATP channel activity below this window depolarizes the cell and causes voltage-dependent inactivation of Ca2+ and Na+ channels.
(A) High-glucose concentration reduces α-cell KATP channel activity, reducing glucagon secretion.
(B) Graded application of tolbutamide (in zero glucose) transiently increases glucagon secretion as KATP channel activity is reduced through, and eventually below, the window supporting glucagon release.
(C) The graded application of diazoxide in high-glucose conditions increases α-cell KATP channel activity into, and then above the window supporting glucagon secretion. The result is a transient “re-activation” of glucagon secretion at low-diazoxide concentrations.
(D) In low-glucose (1–2 mM) conditions, graded application of diazoxide increases KATP channel activity above the window supporting glucagon secretion, causing a monotonic inhibition of glucagon release.
Footnotes
Competing interests. The authors have declared that no competing interests exist.
Author contributions. PEM, LE, and PR conceived and designed the experiments. PEM, YZDM, RR, AS, XM, and LE performed the experiments. PEM, YZDM, RR, AS, LE, and PR analyzed the data. PRVJ and RC contributed reagents/materials/analysis tools. PEM and PR wrote the paper.
Funding. Work in the UK was supported by grants to PR from the Wellcome Trust and the European Union (Eurodia LSHM-CT-2006–518153 and BioSim LSHB-CT-2004–005137). Work conducted in Sweden was supported by the Swedish Strategic Foundation, the Göran Gustafsson Stiftelse, the Swedish Research Council, the Swedish Diabetes Association, and the Novo Nordisk Foundation (LE). | [
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17565376 | CARD15/NOD2 Is Required for Peyer's Patches Homeostasis in Mice
Abstract
Background
CARD15/NOD2 mutations are associated with susceptibility to Crohn's Disease (CD) and Graft Versus Host Disease (GVHD). CD and GVHD are suspected to be related with the dysfunction of Peyer's patches (PP) and isolated lymphoid follicles (LFs). Using a new mouse model invalidated for Card15/Nod2 (KO), we thus analysed the impact of the gene in these lymphoid formations together with the development of experimental colitis.
Methodology/Principal Findings
At weeks 4, 12 and 52, the numbers of PPs and LFs were higher in KO mice while no difference was observed at birth. At weeks 4 and 12, the size and cellular composition of PPs were analysed by flow cytometry and immunohistochemistry. PPs of KO mice were larger with an increased proportion of M cells and CD4+ T-cells. KO mice were also characterised by higher concentrations of TNFα, IFNγ, IL12 and IL4 measured by ELISA. In contrast, little differences were found in the PP-free ileum and the spleen of KO mice. By Ussing chamber experiments, we found that this PP phenotype is associated with an increased of both paracellular permeability and yeast/bacterial translocation. Finally, KO mice were more susceptible to the colitis induced by TNBS.
Conclusions
Card15/Nod2 deficiency induces an abnormal development and function of the PPs characterised by an exaggerated immune response and an increased permeability. These observations provide a comprehensive link between the molecular defect and the Human CARD15/NOD2 associated disorders: CD and GVHD.
Introduction
Caspase Recruitment Domain 15 (CARD15) also known as Nucleotide oligomerisation domain 2 (NOD2) has been associated with Crohn's Disease (CD) [1], [2] and graft versus host disease (GVHD) [3], [4]. NOD2 belongs to a family of genes involved in innate immunity [5]. It can be activated by muropeptides which are components of the bacterial cell wall. When activated, NOD2 interacts with Rick/Rip2 which in turn activates the NF-kB pathway, resulting in the production of pro-inflammatory cytokines.
Half of CD patients have one or more NOD2 mutations [6]. Some of the CD associated mutations were found unresponsive to muropeptides [5]. By consequence, CD is usually considered as an immune deficiency toward bacteria present in the gut lumen [7]. However, the exact mechanism by which NOD2 mutations are able to induce CD lesions is still subject to debate [7]–[10].
Holler et al. reported that the three major mutations associated with CD (R702W, G908R and 1007fs) are also associated with severe acute GVHD and bone marrow transplantation (BMT) related mortality [3]. Mutations in both donor and recipient were found deleterious, suggesting a role of epithelial and circulating cells in disease mechanisms. Despite some differences in their conclusions, other groups recently confirmed the association between NOD2 and BMT complications [4], [11].
CD is a chronic relapsing inflammatory bowel disease (IBD) with mucosal ulcerations of the digestive tract. CD lesions are characterised by a T helper (Th) 1 immune response and several authors have shown that they are related with gut associated lymphoid tissue (GALT), known as lymphoid follicles (LFs). LFs are mainly encountered in the colon where they are isolated and in small bowel where they are grouped forming Peyer's patches (PP) which are known to be pivotal sites for the host immune response and for the entry of enteropathogen bacteria. CD lesions are most often localized in colon and distal ileum, where the LFs are the most abundant [12]. Fujimura et al. found that aphtoïd ulcerations (which are often considered as the earliest CD lesions) are centred by LFs [13]. In addition to this spatial relationship, a temporal link between CD and PP development has also been suggested [14]. PPs develop from birth to 10–15 years of life and then undergo involution. The age-dependent incidence curve of CD is roughly parallel to the number of PP with a delay of about 10 years. Finally, ileal lesions are uncommon in young children and seniors where PP are rare [15], [16].
The role of GALT in GVHD has been suspected on both experimental models and clinical observations. Animal studies showed that death after BMT was prevented by gut decontamination and that prevention of mucosal damage also prevents lethal GVHD [17]. In clinical practice, gut decontamination reduces the frequency and severity of GVHD. Finally, Murai et al. recently showed that PP deficient mice are resistant to GVHD, arguing for a crucial role of PPs in GVHD, at least in models that do not use conditioning of the host prior to adoptive transfer of the allogeneic donor cells [18], [19].
Considering all these elements, we hypothesized that Nod2 may play a role in the structure and function of the GALT. Consequently, we used a new mouse model deficient for Card15/Nod2 i) to evaluate the involvement of Nod2 on the number and PPs size; ii) to assess whether Nod2 modified cellular composition and cytokine expression of PPs; and iii) to determine whether Nod2 may alter paracellular permeability and bacterial translocation of PPs in adult mice. Finally, we also examined if Card15/Nod2 deficiency affects the colonic response to 2,4,6-trinitrobenzene sulphonic acid (TNBS), a classic experimental model of colitis in mouse.
Results
Body weight, intestinal length and intestinal weight were similar in KO and WT mice (supplementary information (SI) Table S1). Macroscopically, no inflammation was visible in KO mice according to Wallace and Ameho criteria (data not shown). KO mice exhibited an increased number of PP in comparison with WT mice at weeks 4, 12 and 52 after birth (Fig 1 A). At weeks 12 and 52, the number of isolated LFs per small intestine was also higher in KO mice than in WT mice (Fig 1 B). In contrast, the number of PPs at birth was similar between KO and WT mice (6.2±0.5 vs. 6.0±0.4; P>0.05) (Fig 1 A).
Macroscopically, the size of PP formations appeared to be larger in adult KO mice (data not shown). To quantify this difference, the three biggest PPs of each mouse were pooled and their cells were counted. At weeks 4, 12 and 52, KO mice exhibited a higher cell number per PP (fig 1 C). In contrast, cell counts were comparable in spleens of KO and WT mice (Fig 1 D).
Microscopic analyses of PP formations from KO mice revealed no gross abnormalities (Fig 1 E). Finally, as NOD2 modulates the NF-κappaB pathway and putatively the apoptosis, we investigated the number of apoptotic cells inside PPs. Immunohistochemistry experiments did not show differences between KO and WT mice for the number of caspase 3 positive cells (1.01±0.11 vs. 1.32±0.10; P>0.05) (Fig 1 E).
In order to better characterise the phenotype of the cells present in PPs, we performed flow cytometry experiments using the B220, CD3 and CD11c antibodies. At week 12, no differences were seen regarding the relative proportions of B220+ B-cells, CD3+ T-cells and CD11c+ dendritic cells between KO and WT mice either for PPs or spleens (Fig 2 A–B).
Because some cells present in a limited number may be functionally important, we also investigated the relative proportion of Ly-6G+ polymorphonuclear neutrophils (PMN) present in the PPs (Fig 2 C) and the number of M cells located inside the follicle associated epithelium (FAE) (Fig 2 D). No difference of PMN relative proportion was observed in PPs of KO and WT mice (Fig 2 C). At the opposite, M cell number was increased in the FAE of KO mice in comparison with WT mice (Fig 2 D).
As T-cells are known to play a pivotal role in CD, we further analyzed the CD3+ T cells. At week 12, KO mice exhibited an increase of CD4+ T-cell relative proportion within their PPs, whereas the proportion of CD8+ T-cells remained constant (Fig 3 A). As a mirror image, PPs from KO mice exhibited significantly fewer CD3+CD4−CD8− T-cells (Fig 3 A). In contrast, no comparable differences were seen in the spleens (Fig 3 B). Similar data were obtained at week 4 (SI Fig S1 A–B).
Finally, we investigated the phenotype of CD4+ T-cells present in the PPs and spleens by examining the relative proportions of naive CD25−CD45Rb+, regulatory CD25+CD45Rb− and memory CD25−CD45Rb− CD4+ T-cells. KO and WT mice had a similar relative proportion of naive, regulatory and memory CD4+ T-cells in PPs (SI Fig S2 A) and spleen (SI Fig S2 B). Flow cytometry analyses also failed to reveal a difference between KO and WT mice when investigating the annexin V positive CD3+ and CD3+CD4+ T-cells extracted from PPs (SI Fig S2 C) suggesting that the excess of CD4+ T-cells observed in KO mice does not result from a defect of apoptosis.
Considering the differences of PP cell number and cellular composition, we further evaluated whether PPs from KO and WT mice might exhibit differences regarding their cytokine profile. Thus, the expression of IL-1β, IFNγ, TNFα , IL-12, and IL-4 in PPs, PP-free ileum and spleen were determined by ELISA, at weeks 4 and 12. Levels of IFNγ, TNFα, IL-12, and IL-4 were significantly increased in PPs of KO mice (Fig 4). In contrast, less marked differences were seen between KO and WT mice in PP-free ileum where TNFα and IFNγ were the only cytokines differentially expressed at week 4 but not in adult mice (Fig 4). Finally, no differences were observed in spleens from KO and WT mice (Fig 4).
Cell composition and cytokine production may affect the function of PPs. We thus used Ussing chambers for determining the paracellular permeability through PP and PP-free ileum. PPs and PP-free ileum of KO mice exhibited a significant increase in paracellular permeability (Fig 5 A) while the electrical resistances were comparable (data not shown). In parallel to this change, mRNA expression levels of TJ proteins were affected in PP of KO mice. Indeed, expression of mRNAs encoding ZO-2 and ZO-1 were decreased by 45% and 36% (P<0.01 and P<0.06 respectively) (Fig 5 B). In contrast, Occludin expression was unchanged (Fig 5 B).
Because Nod2 is involved in innate immunity we hypothesised that Card15/Nod2 deficiency may affect the gut microflora. We thus counted the numbers of bacteria classified as Enterobacteriaceae, Pseudomonas, Staphylococcus, Streptococcus, Enterococcus or Lactobacillus in the ileum of KO and WT mice. However, we did not observed differences between groups (SI Table S2).
Ussing chamber experiments were also performed to investigate the bacterial passage through PP. The passage of a chemically killed Escherichia coli (K-12) was higher in PP of KO mice (P<0.01) (Fig 5 C). This increased translocation was also observed using living non pathogenic Escherichia coli strain J53 (Fig 5 D). In contrast, translocation of Escherichia coli across free-PP ileum mucosa was very low and not significantly increased (P>0.05, fig 5 C). The translocation through PPs of a chemically killed Staphylococcus Aureus (Wood strain without proteinA) and Saccharomyces cerevisiae was also higher in KO than in WT mice (Fig 5 E and F)).
TNBS colitis is a well studied model of acute colitis in mice but no data are currently available in the literature on the effect of Card15/Nod2 in this experimental model. Three days after intracolonic instillation of TNBS, the body weight loss was higher in KO than in WT mice but this difference did not reach significance (Fig 6 A). Nevertheless, the colonic mucosal damage score was higher in KO mice (Fig 6 B) as well as mucosal concentrations of IL-1β, TNF-α and IL-12 (Fig 6 C).
Discussion
Since the discovery of an association between CARD15/NOD2 mutations and both CD [1], [2] and GVH [3], [4], the pathophysiological functions of CARD15/NOD2 involved in CD and GVHD development are poorly understood [9]. However, because an association between GALT dysfonctions and GVHD, as well as spatial and temporal links between CD lesions and PP have been suggested by several authors, we used a new model of Card15/Nod2 deficient mouse, to explore the impact of CARD15/NOD2 in PP development and function. We observed that adult KO mice exhibit an excess of PP and isolated LFs in the gut. PP from KO mice are characterized by an excess of M cells and CD4+ T-cells and a higher expression of Th1 and Th2 cytokines. These differences are associated with increased paracellular permeability and bacterial and yeast passage through PPs. Finally, we observed that Card15/Nod2 deficient mice are more susceptible to TNBS induced colitis. As a whole, the here reported phenotype of the Card15/Nod2 KO mouse is reminiscent to the observations made in the Human diseases associated with CARD15/NOD2 mutations: CD and GVHD.
Our data demonstrate first that Card15/Nod2 deficient mice have an elevated number of PPs and isolated LFs after birth. As intestinal weight and length are similar between KO and WT mice, this finding does not seem to be secondary to an intestinal overgrowth. In addition, PPs from deficient mice are larger, as indicated by the macroscopic examination of the intestines and by the count of PP cells. As a result, Card15/Nod2 modulates the development of the GALT and Card15/Nod2 deficiency is characterised by an overgrowth of the lymphoïd tissue present in the gut. This over-development of the lymphoïd tissue seems to be specific to the GI tract as non significant change were observed in a systemic immune organe like spleen. The lack of difference in PP number between WT and KO mice at birth indicates that Card15/Nod2 plays its role during post natal development of the GALT. While lymphotoxin and IL-7 signalling are essential for the organogenesis of PP during the embryonic stage [20]–[22], it is widely believed that gut commensal bacteria are critical for the postnatal development of gut mucosal immune system, as demonstrated by studies on germ-free animals [23]. Such animals have an underdeveloped GALT and are resistant to experimental colitis and to severe GVHD [18]. Thereby, pattern recognition receptors, including TLRs and NOD molecules, can be seen as good candidates by which the resident flora stimulates the development of GALT. However, TLRs play only a limited role in PP development at early postnatal stage in mice [24]. At the opposite, our data suggest that Card15/Nod2 plays a pivotal role in the postnatal development of GALT.
Because gut flora is important in PP development, it can be questioned if Card15/Nod2 deficiency affects gut flora composition of the host. In order to answer this question, we counted the bacteria most represented in the ileum and able to cultivate in standard conditions. We failed to demonstrate differences between KO and WT mice suggesting that Card15/Nod2 deficiency does not induce gross abnormalities of the gut microflora. However, more discrete alterations cannot be discarded and additional experiments using germ-free animals and/or molecular methods for bacterial detection are required to further explore the relationship between Card15/Nod2 and the normal gut flora.
PPs have B-cell follicles and germinal centres surrounded by areas that contain predominantly T cells. The analysis of the PP from Card15/Nod2 deficient mice failed to reveal gross abnormalities in terms of microachitecture, apoptosis or cell composition (at least for the three main cell lineages present in PP, namely B-cells, T-cells and dendritic cells). This observation is in accordance with the previous descriptions of other Card15/Nod2 KO models of mice which failed to reveal gross intestinal abnormalities under basal conditions [7], [25]. Interestingly, the phenotype observed in the deficient mouse is reminiscent with the Human CD condition where large lymphoid aggregates with normal microarchitecture and cell composition have been reported [26].
LFs are covered by a specialised epithelium including M cells. These cells are able to transfer bacterial and food antigens from the gut lumen to antigen presenting cells [23]. They have thus a pivotal role in the function of PP. M cells were found more numerous in KO mice. In addition, PP of KO mice exhibited a higher proportion of CD4+ T-cells and a decreased percentage of CD3+CD8−CD4− T-cells. As Nod2 may modulate the apoptosis, we have hypothesized that this increase of CD4+ T-cell number may result from an apoptosis defect in the lymphoid cells of KO mice. However, flow cytometry analyses revealed that the relative proportion of apoptotic CD4+ T-cells from PP was similar between KO and WT mice. As a result, our experiments do not support the opinion that Card15/Nod2 deficiency is characterised by a general apoptosis defect in the lymphoid tissue of KO mice. Finally, the altered immune cell composition is concomitant with an increase of pro-inflammatory Th1 but also anti-inflammatory Th2 cytokine expression. Altogether, these results indicate that under basal condition, PPs of KO mice are characterised by an exaggerated immune response.
Cell composition and cytokine production may affect the function of PP. We thus used Ussing chambers for determining the paracellular permeability through PP and PP-free ileum. KO mice exhibited a significant increase of paracellular permeability. This result is in accordance with the altered intestinal permeability reported in CD patients and their healthy relatives, especially in case of CARD15/NOD2 mutations [27]–[30]. Pro- and anti-inflammatory cytokines are known to modulate intestinal paracellular permeability. IFNγ, TNFα and IL-4, act on membrane receptors of epithelial cells to increase tight junction permeability [31]–[34]. For example, on T84 cells IFNγ decreased levels of ZO-1 and altered apical actin organisation, which leads to disorganisation of TJ and increased permeability [35]. Similarly, PP from KO mice exhibited a decrease of ZO-1 and ZO-2 mRNA expression in comparison with WT mice. Consequently, the excessive concentration of IFNγ observed in KO mice may down regulate the transcription of ZO-1 and ZO-2 mRNA expression and contribute to the increase paracellular permeability in KO mice. It is to note that this phenotype is reminiscent to the CARD15/NOD2 associated Human disorders. Indeed, in CD patients, increased permeability has been reported to be mediated by TNFα and to precede the clinical relapse while GVHD has been treated by anti-TNF antibodies [36], [37].
Because of the changes in gut permeability, we finally studied the role of Card15/Nod2 in bacterial translocation. Bacterial passage of Staphylococcus aureus and Escherichia coli and yeast ingress of Saccharomyces cerevisiae were higher through PP of KO mice. It is widely accepted that CD is related with an excessive bacterial translocation through the intestinal epithelium even if this hypothesis is not perfectly documented. The present data further support this opinion. In addition, this excess of yeast and bacteria translocation through PP of KO mice is in agreement with recent reports showing that mutated CD patients and their unaffected relatives develop more frequently antibodies to Saccharomyces cerevisiae, Pseudomonas fluorescens–related protein, Escherichia coli outer membrane porin C and CBir1 flagellin [38]. The excessive bacteria and yeast passage reported here may participate in the enhancement of adaptive immune responses to microbial antigens.
The increased number of M cells can contribute to the high translocation rate observed in KO mice. However, because M cell differentiation is inducible by microbial challenge [39], it may also be a consequence of bacterial translocations and additional experiments are required to further dissect this complex relationship. It is also possible to consider that CARD15/NOD2 dysfunction facilitates bacterial entry through defective antibacterial peptide expression [7], impaired intracellular bactericidal capacity or reduced epithelial immune defence. Finally, bacterial translocation may also be secondary to primitive local cytokine changes. IFNγ is known to increase the epithelial adherence of selected species of enteric bacteria [40]. Ferrier et al. have shown that a chronic stress in mice drives an organ-specific cytokine expression pattern which in turn, alters the colonic mucosal barrier functions and favours bacterial translocation [31]. This effect is dependent on the presence of CD4+ T-cells and requires IFNγ production. It is thus possible that bacterial translocation is a result of the immune changes rather than its cause and additional experiments are now required to answer this question.
Finally, we have shown that Card15/Nod2 invalidation exacerbates the severity of TNBS induced colitis, as evidenced by the increase in all parameters characterising colonic inflammation (damages scores and mucosal levels of IL-1β, TNFα and IL-12). This enhanced colitis-induced by TNBS might be explained by different mechanisms. Firstly, since microflora is required for TNBS-induced colonic mucosal damages and since Card15/Nod2 signaling has been shown to inhibit the TLR2-driven activation of Th1 response [8], one can hypothesize that the Th1 inflammatory response mediated by TLR-2 is increased in Card15/Nod2 deficient mice. Secondly, since GALT is reported to modulate the severity of TNBS-induced colitis [41], [42] it can be hypothesised that the TNBS induced colitis is exacerbated in Card15/Nod2 KO mice because of GALT overdevelopment. Finally, an alternative explanation could be that the observed defect in terms of intestinal permeability and bacterial translocation makes the intestine more susceptible to TNBS.
Altogether, the present data demonstrate that Card15/Nod2 is required to maintain the homeostatis of the PPs. Because CARD15/NOD2 is a well demonstrated etiological factor for CD/GVHD, this conclusion is of particular importance for the understanding of disease mechanisms. Indeed, it further supports the opinion that the defect involved in the development of the gut lesions is related with PP and LF function. The GALT dysfunction observed in KO mice is associated with an excessive gut immune response and an increased bacterial translocation. These findings are consistent with our knowledge on the Human diseases associated with CARD15/NOD2 mutations namely GVHD and CD and for which similar observations have been reported. As a result, the phenotype of the Card15/Nod2 KO mice can be seen as an attenuated model of CD/GVHD. It is to note that the absence of a full phenotype is not unexpected considering the multifactorial nature of the Human diseases where exposure to several additional unknown genetic and environmental risk factors is required for disease expression. At the opposite, our work indicates that the Card15/Nod2 KO mouse is a relevant model for investigating these other risk factors associated with CD and GVHD.
Material and Methods
Animals
Card15/Nod2 was disrupted in mice by replacing the first coding exon carrying the majority of the sequence encoding the CARD domains including the start codon with a EGFP cassette. (Fig. 7 A). In the first step of the strategy, the Card15/Nod2 locus was targeted with the Card15/Nod2 KO targeting fragment. In the second step, the PGKHygromycin selection cassette was removed by Cre recombinase. Targeted disruption was determined by Southern blot and long-range PCR analysis (Fig. 7 B–C). Genomic DNA from mice was amplified by PCR using the primers: 5-GTCATTTCCTGACCTCTGACC-3 and 5-AACCGCATTATTCCATGGGGC-3 to detect WT DNA and primers 5-AACCGCATTATTCCATGGGGC-3 and 5-GCCTGCTCTTTACTGAAGGCTC-3 to detect the disrupted sequence. The loss of mRNA Nod2 expression was demonstrated in splenocytes by RT-PCR (Fig. 7 D) using the following primers: (5-CTTTGAACTGTATGGGTCC-3 and 5-CTCCACTGCCTCTGCCTTA-3). As expected, no signal was observed in KO mice.
The Card15/Nod2−/− (KO) mice used for this study were back-crossed five times with the inbred strains C57BL/6. WT and KO mice were housed and generated in the animal facility at Robert Debré Hospital, Paris, France. The absence of enteropathogens was monitored. All experiments were approved by the institutional committee for animal use.
Peyer's patches and isolated lymphoid follicle numbers
The entire small intestines were removed and the number of PPs was determinated by macroscopic observation except at birth where PPs were too small to be seen by the naked eye. At birth, small intestines were fixed in formalin, stained with 0.5%methylene blue and decolorized in fresh 2% acetic acid. For LF counts, small intestines were fixed in formalin and rolled up into a ‘Swiss-roll’, embedded in paraffin blocks and cut into 5 µm sections. LFs were counted in a blind fashion on two sections per blocks after haematoxylin-eosin staining.
Cell composition of Peyer's Patches and spleens
Cell suspensions from PPs were prepared by pressing the three largest PPs for each mouse with a 5 ml syringe piston. The preparation was then incubated with 100 U/ml collagenase D (Roche, Mannheim, Germany) for 30 minutes at 37°C in DMEM media. Cells from spleen were isolated using the same procedure with an additional step of erythrocytes lysis (Gey's-solution). After centrifugation, cells were re-suspended in DMEM media and submitted to flow cytometry analyses on a FACScalibur (Becton-Dickinson), and analyzed by Cell Quest 3.3 (Becton Dickinson). Monoclonal antibodies used to stain cell suspensions were purchased from BD Biosciences (Pharmingen Heidelberg, Germany) : PE-Cy5-anti-CD3 (17A2), PE-Cy7-anti-CD3 (145-2C11), PE-Cy5-anti-CD4 (H129.19), PE-anti-CD4 (RM4), FITC-anti-CD8 (53-6.7), PE-anti-CD11c (HL3), PE-anti-CD25, PE-anti-CD45R/B220 (RA3-6B2), FITC-anti-CD45RB (16A), APC-anti-annexin V and eBioscience (San Diego, CA) : APC-anti-Ly-6G (RB6-8C5).
Immunohistochemistry
M cells were counted in the FAE using a fluorescence microscope after immunostaining of PP cryostat sections (5 µm) with anti-Ulex Europeaeus antibodies (1/250, 2 h) (Sigma, France), revealed by anti-Ulex Europeaus agglutinin I (1/500, 30 min.) (Vector laboratories) and anti-rabbit FITC conjugate (1/80, 30 min.) (Sigma, France). Caspase 3 immunostaining was done using rabbit polyclonal antibodies to Cleaved Caspase-3 (Asp 175, dilution: 1/100) ( Cell Signaling Technology, Inc Ozyme, Beverly, MA, USA).
Cytokine Enzyme-Linked Immunosorbent Assay (ELISA)
PPs, ileum and spleen from WT and KO mice were removed, washed with cold PBS and the concentration of protein was determined using commercial kit (Biorad, Marnes la Coquette, France). TNFα, IFNγ, IL-4 and IL-1β were determined by ELISA assays (BD Biosciences) according to the manufacturer's instructions. All experimental groups were tested in duplicates.
Ussing chamber experiments
Biopsies from ileum with or without PPs were placed in a chamber exposing 0.196 cm2 of tissue surface to 1.5 ml of circulating oxygenated Ringer solution at 37°C. PP and ileum permeability were assessed by measuring steady-state (from 1 to 2 h) mucosal-to-serosal flux of 4 kDa FITC-dextran (Sigma, St. Quentin Fallavier, France). Bacterial translocation was studied using chemically killed fluorescein-conjugated Escherichia coli K-12 or Staphylococcus Aureus BioParticles (Molecular Probes, Leiden, the Netherlands) or a viable Escherichia coli (the J53 strain resistant to rifampicin) at a final concentration of 1.107 CFU/ml in the mucosal reservoir. Saccharomyces cerevisiae translocation was studied using chemically killed fluorescein-conjugated S. cerevisiae BioParticles (Molecular Probes, Leiden, the Netherlands) at a final concentration of 1.107 CFU/ml in the mucosal reservoir.
Real time reverse transcription-polymerase chain reaction (RT-PCR)
After extraction from PP of ileum by the NucleoSpin® RNA II Kit (Macherey-Nagel, Hoerdt, France), total RNA was converted to cDNA using random hexonucleotides and then used for PCR. We conducted PCR with QuantiTect SYBR Green PCR Kit (Applied, Courtaboeuf, France) using sense and antisense primers specific for: G3PDH, 5′-CACCATCTTCCAGGAGCGAG-3′ and 5′-GCCTTCTCCATGGTGGTGAA-3′; Occludin (Occ), 5′-AGCCTTCTGCTTCATCGCTTC-3′ and 5′-GTGGCAATAAACACCATGATGC-3′; Zonula Occludens-1 (ZO-1), 5′- GACTCCAGACAACATCCCGAA-3′ and 5′- AACGCTGGAAATAACCTCGTTC -3′; Zonula Occludens-2 (ZO-2), 5′-CAGCCACAATCAACGTGAATTC-3′ and 5′-CTGTCCTTCAAGCTGCCAAAC-3′. After amplification, we determined the threshold cycle (Ct) to obtain expression values.
Bacterial content of ileum
The entire ileum (5 cm) was removed and ileal content was collected using 3 mL of steril water (Biorad, France) administered with a polypropylene syringe. Then, ileal content was homogenized and serial dilution (50 µL) of each aliquot were plated onto 5 selective gelose (URI 4, Drigalski, Columbia ANC+5% of sheep blood, Chapman and Coccosel). Plates were incubated for 24 hours at 37°C under aerobic condition and the number of colony forming units was counted and expressed as cfu/mg of ileal content.
TNBS induced colitis
Under anaesthesia colitis was induced in 12 week old mice by a single intracolonic administration of 120 mg/kg TNBS (Sigma, France) dissolved in 50% ethanol. A 50 µl aliquot of the freshly prepared solution was injected into the colon, 4 cm from the anus, using a 3.5 F polyethylene catheter. The mice were weighed and killed 72 h after TNBS administration. Then, body weight, macroscopic damage score according the Wallace scores [43], and cytokines levels were assessed.
Statistical Analyses
Values are expressed as mean±SEM. Statistical analysis were performed using GraphPad Prism 4.00 (GraphPad Software, San Diego, CA, USA) software package for PC. Single comparisons were performed by unpaired Student's t-test. A value of P<0.05 was considered as statistically significant. All P values were two sided.
Supporting Information
Table S1
Impact of Nod2 on body weight, gut weight and intestine length. At weeks 4, 12 and 52, we investigated the body and gut weight and the intestine length of KO and WT mice. No difference was observed between KO and WT mice (P>0.05). Data represent the means±SEM of 8 mice per group.
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Table S2
Ileal microflora under basal condition. Under basal condition, no difference was observed between KO and WT mice (P>0.05 for each bacterial group). Data represent the means±SEM of 10 mice per group.
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Click here for additional data file.
Figure S1
PPs from KO mice exhibit higher rates of CD4+ and CD4-CD8-T-cells at week 4. At week 4, CD3+ T-cells recovered from PPs (A) and spleen (B) were stained with antibodies to CD3, CD4, and CD8 from KO (▪) and WT (□) mice. Data were gated for CD3+ T-cells. Relative proportions of both CD3+CD4+ and CD3+CD4-CD8- T-cells were significantly higher in the PPs but not in the spleen (P>0.05) of KO mice. Data represent the means±SEM of 8 mice per group. *P<0.05; **P<0.01.
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Figure S2
Nod2 and CD3+ T -cells in Peyer's Patches. (A and B) Relative proportions of naïve, regulatory and memory T-cells in PPs (A) and spleens (B) of KO (▪) and WT (□) mice at week 12. CD4+ T-cells were stained with antibodies to CD25 and CD45RB. (C) Relative proportions of apoptotic CD3+ and CD3+CD4+ T-cells. Apoptotic CD3+ and CD3+CD4+ T-cells were investigated by flow cytometry using antibodies to CD3, CD4 and annexin V. Data were gated for CD3+CD4+ T-cells. Data represent the means±SEM of 8 mice per group.
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Click here for additional data file.
Acknowledgements
We acknowledge Veronique Mégras, Catherine Martinet, Michel Peuchmaur, Régine Paris, Pascal Blain, Jean-Baptiste Huguet, Latifa Ferkdadji, Françoise Merlin, Robert Ducroc, Christèle Madre, Sarah Cheriet and Pauline Thabuteau for their excellent assistance.
Figures and Tables
Figure 1
Nod2 and postnatal development of gut associated lymphoid formations.
(A) PP count on the whole intestines of KO (▪) and WT (□) mice at birth and at weeks 4, 12 and 52. (B) Number of isolated LFs identified on microscopic examination of the small intestine at weeks 12 and 52. (C and D) Number of cells per PP and spleen at weeks 4, 12 and 52). (E) Number of apoptotic cells identified by caspase-3 immunostaining at week 12. Data represent the means±SEM of 8 mice per group. *P<0.05; **P<0.01, ***P<0.001.
Figure 2
Nod2 and Peyer's patch cellular composition.
(A and B) Relative proportions of T-, B- and dendritic cells from PPs and spleens of KO (▪) and WT (□) mice at week 12. T-, B- and dendritic cells were investigated by flow cytometry using antibodies to CD3, B220 and CD11c. (C) Relative proportion of polymorphonuclear neutrophils was analyzed using Ly-6G antibody. (D) M cells number inside the follicle associated with epithelium was investigated by immuno-histochemistry. Arrows indicated the presence of M-cell inside the follicle associated with epithelium. Data represent the means±SEM of 8 mice per group. **P<0.01.
Figure 3
Nod2 and T-cells subset in PP and spleen from mice at 12 weeks of age.
Relative proportion of CD4+, CD8+ and CD4−CD8− T-cells from PPs (A) and spleen (B) of KO (▪) and WT (□) mice. At week 12, CD3+ T-cells were stained with antibodies to CD3, CD4, and CD8. Data were gated for CD3+ T-cells. Data represent the means±SEM of 8 mice per group. *P<0.05.
Figure 4
Nod2 invalidation increases cytokine expression in PP.
Expressions of Il-1β, IFNγ, TNFα, IL-12, IL-4 were determined by ELISA in PPs, Ileum and spleens from KO (▪) and WT (□) mice at weeks 4 (left panel) and 12 (right panel). Data represent the means±SEM of 8 mice per group. *P<0.05; **P<0.01.
Figure 5
Paracellular permeability and bacterial translocation are increased in Nod2 invalidated mice.
Ussing-chamber and Real-time PCR experiments were performed on PP and PP-free ileum from WT (□) and KO (▪) at week 12. (A) Paracellular permeability was analysed by FITC-dextran flux from PP and PP-free ileum under basal condition. (B) mRNA expression levels of TJ proteins (ZO-1, ZO-2 and Occ) from PP were analysed by real-Time-PCR under basal condition. (C) Bacterial translocation of chemically killed fluorescent Escherichia Coli K-12. (D) Translocation of the viable non enteropathogen Escherichia Coli strain J53. (E) Translocation of a chemically killed fluorescent Staphylococcus Aureus. (F) Translocation of chemically killed fluorescein-conjugated Saccharomyces cerevisiae. Data represent the means±SEM of 8 mice per group. *P<0.05 and **P<0.01, significantly different from WT.
Figure 6
Nod2 invalidation increased the suceptibility of TNBS induced colitis.
(A) Body weight, (B) Macroscopic damage score, (C) cytokines levels were assessed in 12 week old mice. These parameters were determined three days after intracolonic administration of TNBS. Values are mean (SEM) (n = 8 in each group). *P<0.05 and **P<0.01 between WT and KO mice.
Figure 7
Targeting disruption of the murine Nod2 gene by homologous recombination.
(A) Generation of the Card15/Nod2 KO allele: targeting strategy. The restriction maps of the Card15/Nod2+allele (5′ portion), the Card15/Nod2 KO targeting fragment, and the modified Card15/Nod2 allele after homologous recombination and Cre-mediated recombination of an Hygromycin selection cassette are shown. Exons 1, 2, and 3 (black boxes) and restriction sites used for cloning and screening (X) XbaI, (A) AgeI, (P) PmlI, (S) SalI, (N) NotI are indicated. The Card15/Nod2 KO targeting fragment comprises the EGFP gene (Clontech) in frame with Card15/Nod2 ATG and the floxed PGKHygromycin (Clontech) selection cassette, introduced between the AgeI site downstream of Card15/Nod2 exon 1 and the SalI site upstream of exon 1 in the Card15/Nod2 orientation. All loxP sites are represented by open triangles. Recombination of loxP1 and loxP2 results in the loxP1+2 site. In the first step of the strategy, the Card15/Nod2 locus was targeted with the Card15/Nod2 KO targeting fragment. In the second step, the PGKHygromycin selection cassette was removed by Cre recombinase. The double-headed arrows indicate the DNA fragments resulting from digestions with different enzymes expected to hybridize with probes A, B or Hyg. Also depicted are combinations of PCR primers P1-4 that detect the different Card15/Nod2 alleles. (B) Southern blot analysis of restricted DNA resulting from digestion with XbaI.Hybridation with Probe A and B show complete integration of the targetting fragment after homologous recombination. Probe Hyg show the differents integrations of targetting fragments after homologous recombination. (C) Genotyping of Nod2-deficient mice by PCR. Genomic DNA from mice was amplified by PCR to detect the disrupted sequence (PCR product of 459 bp). (D) Expression of Nod2 mRNA in spleen. RT-PCR was performed on purified mRNA from the spleen of WT and KO mice. As expected, no signal was observed in KO mice. GAPDH expression was used as positive control of expression.
Footnotes
Competing Interests: The authors have declared that no competing interests exist.
Funding: This work was supported by the Institut National de la Santé et de la Recherche Médicale, the Broad Medical Research Program, la Mairie de Paris, la Fondation pour la Recherche Médicale, BREMICI, l'association François Aupetit and la région Ile de France. | [
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17677002 | Complex Seizure Disorder Caused by Brunol4 Deficiency in Mice
Abstract
Idiopathic epilepsy is a common human disorder with a strong genetic component, usually exhibiting complex inheritance. We describe a new mouse mutation in C57BL/6J mice, called frequent-flyer (Ff), in which disruption of the gene encoding RNA-binding protein Bruno-like 4 (Brunol4) leads to limbic and severe tonic–clonic seizures in heterozygous mutants beginning in their third month. Younger heterozygous adults have a reduced seizure threshold. Although homozygotes do not survive well on the C57BL/6J background, on mixed backgrounds homozygotes and some heterozygotes also display spike-wave discharges, the electroencephalographic manifestation of absence epilepsy. Brunol4 is widely expressed in the brain with enrichment in the hippocampus. Gene expression profiling and subsequent analysis revealed the down-regulation of at least four RNA molecules encoding proteins known to be involved in neuroexcitability, particularly in mutant hippocampus. Genetic and phenotypic assessment suggests that Brunol4 deficiency in mice results in a complex seizure phenotype, likely due to the coordinate dysregulation of several molecules, providing a unique new animal model of epilepsy that mimics the complex genetic architecture of common disease.
Author Summary
Epilepsy is a very common brain disorder characterized by recurrent seizures, resulting from abnormal nerve cell activity in the brain. Some cases of epilepsy are caused by brain trauma, such as stroke, infection, tumor, or head injury. Others—so called “idiopathic”—do not have a clear cause. Many idiopathic epilepsies run in families, but the inheritance patterns and complex seizure types suggest that they are not due to a single defective gene but instead are caused by multiple gene defects that are inherited simultaneously in a patient. This complex inheritance makes it difficult to pinpoint the underlying defects. Here, we describe a new mutant mouse, called “frequent-flyer,” which has several different types of seizures. Although these seizures are caused by a mutation in a single gene, because this gene regulates the expression of many other genes, which, in turn, cause abnormal nerve cell activity, frequent-flyer mice provide a unique animal model of epilepsy—mimicking the complex genetic architecture of common disease.
Introduction
Epilepsy, defined by recurrent seizures resulting from abnormal, synchronized neuronal firing in the brain, is a very common neurological disorder. Idiopathic epilepsies do not have any antecedent disease or injury to the brain and many are suspected to have a genetic basis. The difficulty of elucidating defective genes underlying common inherited epilepsies is that they are genetically complex—being caused by multiple variants that are coinherited in affected individuals [1,2]. To date, most mutations involved in idiopathic epilepsy have been found in genes encoding ion channels or their accessory subunits with a few exceptions, for example, LGI1 [3] and EFHC1 [4] in humans, [5] and JRK/JH8 [6,7] in both humans and mice. Such exceptions are of interest in that they may lead to further understanding of epilepsy disease mechanisms beyond primary excitability defects, for example, by identification of genes that modulate the expression or function of the more proximal candidates for epilepsy—ion channels, neurotransmitter receptors, and synaptic proteins.
Here we describe the disruption of the expression of an RNA-binding protein, BRUNOL4 (Bruno-like 4) leading to partial limbic and tonic–clonic seizures in a new mouse model of epilepsy called “frequent-flyer” (abbreviated Ff; gene symbol: Brunol4Ff). BRUNOL4 (also known as CELF4, CUG-BP, and ETR-3 like factor 4) belongs to a family of RNA-binding proteins involved in multiple aspects of RNA processing such as pre-mRNA splicing [8], mRNA editing [9], and RNA stability and translation [10]. There are six family members in both humans and mice with orthologs in nematode and fruit fly [11]. The murine and human BRUNOL4 are 99.6% identical at the amino acid level [12]. Mouse knockouts have very recently been published for Brunol1 and Brunol2, which display spermatagonial and varied developmental defects, respectively [13,14]. UNC-75, a neuron-specific ortholog in C. elegans, shares 47% identity with the human BRUNOL4 protein. UNC-75 deficiency in the nematode leads to behavioral phenotypes indicative of abnormal neurotransmission. Human BRUNOL4 can rescue the unc-75 mutant phenotype, suggesting that UNC-75 and BRUNOL4 may be involved in fine-tuning synaptic transmission through regulating RNA processing in the nervous system [15].
In this study we describe the seizure phenotypes of mice carrying Brunol4 disruption, and begin to explore the molecular consequences using gene expression profiling and genetic interaction tests. Our studies suggest that Brunol4 deficiency alters the expression of several molecules involved in synaptic function, which, when combined, account for the complex seizure disorder of frequent-flyer mice.
Results
Origin of the Mutation and Convulsive Seizure Phenotype
The Ff mutation arose from an independent project in which a series of transgenic mouse lines was generated on the C57BL/6J (B6) strain background. One line (9/9 transgene carriers) developed frequent seizures from about three months of age, precipitated by routine handling such as cage transfer. Since the transgene construct was not expressed in all the lines and since other lines using the same construct did not have seizures, together suggested that the seizures were not caused by transgene expression per se. To distinguish between an unlinked spontaneous mutation and insertional mutagenesis, affected mice were outcrossed to normal B6 mice. In the next generation, seizures cosegregated with the presence of the transgene (25/28 carriers displayed seizures versus 0/22 non-carriers), suggesting a high-penetrance, dominant mode of inheritance for the seizure phenotype.
Convulsive seizures ranged in severity, the mildest being muscle twitching in the face and neck, forelimb clonus, and salivation (Figure 1A). More severe seizures included rearing and falling, myoclonic jerks, and arching of the back and tail. In many cases, convulsions were followed by a very wild running–bouncing phase with occasional tonic–clonic hindlimb extension, but which, unlike the equivalent phase of some induced seizures, did not result in lethality. Hence, the allele symbol “frequent-flyer” (Ff) was assigned. The incidence of these handling-associated seizures was higher in male than in female mice.
Figure 1
Phenotypes of Ff Mutant Mice
(A) The posture of a 3-mo-old Ff/+ heterozygote mouse at the beginning of a seizure episode. Note the contraction of the muscles of the neck, ears, and forelimbs and the vertical position of the tail.
(B) Reduced ECT of Ff/+ mice. The median response level (CC50) for minimal seizures were as follows +/+ = 9.17 mA (95% CI 8.83–9.44), Ff/+ = 7.69 mA (95% CI 7.48–7.9). Squares (□) and triangles (▵) indicate individual data points used to construct CC curves in +/+ and Ff/+ mice, respectively.
(C) Late-onset body-weight gain in Ff/+ heterozygous mice. We monitored the body weight between the age of 12 wk and 34 wk of more than 50 male mice with similar number of mutants and controls. Single-caged mice were excluded from this study. Each mouse was weighed every other week. The body weights of mutants did not diverge significantly from those of controls until 28 wk of age and the divergence remained afterwards (t-test, assuming two-tailed distribution and two-sample equal variance, the p-values are 0.024, 0.004, 0.0011, and 0.001 from 28 wk to 34 wk). The average weight of each genotype at a fixed time-point was plotted against time to generate a growth curve using Microsoft Excel (http://www.microsoft.com/). (D) EEG recording from a Ff/Ff homozygous mouse. Shown are the six differential recordings from four subcranial electrodes, one in each quadrant corresponding to front-right (FR), front-left (FL), back-right (BR), and back-left (BL). These traces correspond to one of the longer spike-wave discharge (SWD) episodes observed.
(E) Treatment of SWD in Ff/Ff homozygous F2 hybrid mice from crosses to FVB (white box) or 129S1 (black). Recording sessions began at least 1 h prior to ethosuximide, saline, or fluoxetine (Prozac) treatment, and the rate of SWD (minimum criteria: ≥ 0.5 s duration, amplitude ≥2× baseline) was determined and plotted (“pre”). Animals were then injected with drug, monitored for an additional hour, and SWD incidence was recorded (post). Lines between datapoints correspond to each animal pre- and post-treatment. Only the ethosuximide and fluoxetine results were significantly different comparing treatments (ETX, p = 0.009; fluoxetine, p = 0.019; saline, p = 0.804 matched pairs test).
Although handling-associated seizures did not begin until the third month of age, by 7 wk heterozygotes had markedly reduced electroconvulsive thresholds (ECT) (Figure 1B). In addition to convulsive seizure phenotypes, heterozygotes were also slightly hyperactive, and while slightly smaller at weaning age, they had a late-onset body weight gain in Ff/+ heterozygotes (on average 10% heavier than littermate controls, Figure 1C). Despite the high frequency and the severity of seizures, Ff/+ heterozygotes do not have a reduced life span (analyzed up to 24 mo of age). The morphology of the Ff/+ brain appears normal, as evident in the proper cortical and hippocampal layering and the lack of overt gliosis (unpublished data).
Strain Background Effects on Frequent-Flyer Phenotypes
Ff/Ff homozygotes, however, had a much more severe phenotype; they were born alive at close to Mendelian ratios but most died during the first day. From matings between heterozygotes, only 1.1% (expect 25%) survived until 4 wk of age (Table 1). While alive, homozygotes did not display obvious signs of convulsion or respiratory stress, nor was there any obvious pathology seen in mutant brains (unpublished data). Future work will be needed to clarify the cause of perinatal lethality in Ff/Ff homozygotes. However, when we examined the F2 generation of matings between B6-Ff/+ and six different inbred mouse strains, a range of survival rates of homozygotes were observed was with the highest being 8.2% in crosses with 129S1 (Table 1), suggesting that homozygosity for B6 allele(s) makes the homozygous phenotype worse, as is the case with many neurological mutations in mice (e.g., see [16]). Interestingly, although these F2 hybrid homozygotes often lived for more than 6 mo, they were smaller than littermates and also exhibited spontaneous limbic and tonic–clonic seizures, similar in appearance to those of Ff/+ heterozygotes, except they were observed as early as 8 wk (and we suspect that lethal seizures occurred as early as 4 wk). In addition, Ff/+ heterozygotes on F1 hybrid backgrounds experienced a lower incidence of convulsive seizures later in life (unpublished data). These results show that inbred strains have polymorphisms that attenuate the frequent-flyer phenotypes.
Table 1
Survival of Brunol4Ff/Ff Homozygous Mutants in F2 Generation
Absence Epilepsy in Frequent-Flyer Mutants
The availability of Ff/Ff homozygotes on a mixed genetic background afforded us the opportunity to determine whether they show spike-wave discharges (SWD), the electroencephalographic manifestation of absence seizures—events not observed in Ff/+ heterozygotes on the B6 background (unpublished data). Ff/Ff homozygotes tested on the F2 hybrid backgrounds (B6 × 129S1 or FVB/NJ) experienced very frequent SWD (e.g., see Figure 1D and 1E). Interestingly, heterozygotes in the FVB/NJ cross, but not in the 129S1 cross, also showed a significant rate of SWD (unpublished data). Together, these results suggest that not only do Ff/Ff homozygotes have SWD, but that the penetrance or severity is also modulated by genetic background. Although SWD in Ff/Ff homozygotes were synchronous, rhythmic, and generalized, when compared to those of other SWD-prone mice, such as stargazer or C3H/HeJ (e.g., see [17,18]), the episodes were relatively short, averaging 1.5 s in length, and the rhythmicity was more erratic than in other mutants (Figure 1D). Nevertheless, the animals remained motionless during SWD episodes, and SWD were suppressed by the anti-absence drug ethosuximide (Figure 1E, left), suggesting that they are absence seizures.
Transgenic Insertion Mutation in Brunol4
To determine the identity of the gene disrupted by transgenic insertion, we cloned and sequenced a unique transgene-genomic junction fragment (see Materials and Methods) and found a 100% match to intron 1 of the Brunol4 gene on mouse Chr 18. We then evaluated the impact of transgene insertion on Brunol4 expression. The insertion expanded the 74-kb intron 1 of Brunol4 by at least 20 kb, well upstream of the exons encoding RNA binding motifs (www.ensembl.org/Mus_musculus, Figure 2). Multiple splice donor and acceptor sites were detected in the transgene, suggesting the possibility of the insertion interfering with normal Brunol4 splicing. In total RNA samples from newborn mice, no Brunol4 transcript was detected in Ff/Ff homozygotes and approximately 45% reduction was seen in heterozygotes (Figure 3A); by real-time reverse-transcriptase PCR (RT-PCR), similar reduction was observed in the adult brain of Ff/+ heterozygotes (Figure 3B). We also examined the potential impact of the transgene on the expression of the neighboring genes. The genomic region where murine Brunol4 resides is gene poor. Genes with strong annotation are at least 0.5 Mb either 5′ or 3′ away from Brunol4. Expression analysis of the neighboring genes with potential brain function did not reveal difference between Ff/Ff homozygous mutants and normal controls (unpublished data), suggesting that Brunol4 is the only brain-expressed gene affected by the transgene insertion. This is consistent with preliminary assessment of a gene-targeted null allele of Brunol4 that we made recently, which displays handling-associated seizures in older adults, resembling those of frequent-flyer mice with both limbic and wild tonic-clonic phases; younger heterozygotes also have an unusually low threshold to electroconvulsion (C. L. Mahaffey, W. N. Frankel, unpublished results).
Figure 2
Trangene Insertion into the Brunol4 Locus in Ff Mice
(A) Physical map of the transgene insertion point in the first intron of Brunol4 gene on mouse Chromosome 18. The exact number of copies of the transgene inserted is currently unknown. It appears to be more than four copies based on a Southern blot probed with a transgene-specific probe. The transgene insertion event was a simple addition at the 5′ end, but appeared complex at the 3′ end. The last intact copy of the transgene was inverted to the opposite orientation and there was an 18-kb deletion in the genomic sequence at the 3′ insertion site. Further, a 397-nt fragment from the transgene was left between the last intact copy of the transgene and the intronic sequence from Brunol4. This 397-nt fragment was in the same orientation as that of the transgene at the 5′ insertion site. The telomeric breakpoint of the transgene insertion was 28,987 nt from the exon 1 splice donor and the centromeric breakpoint was 27,056 nt from the exon 2 splice acceptor.
(B) The exon/intron structure of the mouse Brunol4 gene. Note the orientation of the gene is reversed from (A) to facilitate viewing. The open reading frame starts in exon 1 and ends in exon 12. Three RNA recognition motifs are predicted in the BRUNOL4 peptide sequence and the respective coding exons are marked.
Figure 3
Expression of Brunol4 in Mutant and Wild-Type Mice
(A) Reduced Brunol4 transcript abundance in Ff/+ mice. Newborn brain total RNA blot was probed with Brunol4 p1 probe (upper portion) and a mouse β-actin probe (lower portion) sequentially. Brunol4 transcript was absent from Ff/Ff homozygous newborns, and reduced in abundance in littermate +/+ controls. An arrow indicates the position of the Brunol4 transcript.
(B) Relative fold change of Brunol4 by real-time RT-PCR) shown in total RNA from adult cortex in three independent mice of each genotype. Similar results were observed from all other brain regions.
(C) The expression of Brunol4 is restricted to the brain in adult mice. An RNA blot containing poly-A selected RNA extracted from various adult mouse tissues was sequentially hybridized with probes specific to Brunol4 and β-actin as a loading control. The position of the size markers (Ambion, http://www.ambion.com) is shown to the left side of panels.
In order to evaluate the expression pattern of Brunol4 in adults, we examined RNA from a variety of tissues. Only brain samples showed robust signal, despite prolonged exposure time, suggesting that BRUNOL4 is brain specific in adults (Figure 3C), consistent with a recent survey of organ protein expression in mice that detected BRUNOL4 only in the brain [19]. At a higher resolution, Brunol4 showed a predominantly neuronal expression pattern in the brain—labeling was seen in the cerebral cortex, hippocampus, olfactory bulbs, and the granule cell layer of the cerebellum (Figure 4). However, strongest expression in the brain was observed in the hippocampus where high expression was detected in the pyramidal neurons of the CA2 and CA3 region. Pyramidal neurons in CA1, the dentate gyrus granule cells, and the dentate subgranular zone had weaker expression (Figure 4); the latter is interesting in light of the observation that some types of seizure activity may induce neurogenesis in this region (e.g., see [20]).
Figure 4
Brunol4 Expression in Normal Adult Mouse Brain by RNA In Situ Hybridization
A 33P labeled antisense riboprobe (A) or a sense control probe (B) was hybridized to 7-μm parasagittal paraffin sections of brain. Note the positive staining in the olfactory bulb, cerebral cortex, hippocampus, and cerebellum. Despite several attempts, persistent minor background signal was observed at the edges of the brain section, especially in the cerebellum. In order to increase the specificity and better monitor the signal development, a DIG-labeled antisense riboprobe (C) or a sense control probe (D) was hybridized to 15-μm parasagittal cryosections of brain. Representative images are shown focusing on the entire hippocampus. Note the intense labeling (blue color) in the CA2 and CA3 regions, consistent with the strong hipocampal signal seen in panel (A).
CB, cerebellum; CO, cerebral cortex; Hi, hippocampus; OB, olfactory bulb.
Neuroexcitability Candidates Down-Regulated in Mutant Mice
How might BRUNOL4 deficiency result in a complex seizure phenotype? We hypothesize that BRUNOL4 is involved in the processing events of one or more mRNA-encoding proteins that are themselves more directly involved in synaptic function. Thus, in the absence or reduction of BRUNOL4, these molecules become dysregulated, leading to imbalance in neuronal excitability. We carried out microarray analysis to detect genes differentially expressed between coisogenic wild-type and mutant mice. For the primary screen, newborn Ff/Ff homozygous mutants were chosen to optimize the signal differential. Of the approximately 39,000 transcripts interrogated, changes in only 459 transcripts (corresponding to approximately 350 independent genes) were considered statistically significant in the Ff/Ff homozygotes compared with controls (Table S1). Of the 94 down-regulated transcripts (from approximately 70 independent genes), the most reduced was Brunol4 itself, with a significant decrease in Ff/Ff homozygotes. Four genes from the down-regulated list were of obvious interest for an excitability disorder. One encodes serotonin receptor 2c (Htr2c); Htr2c-null mice are known to experience frequent spontaneous seizures, at least on a mixed background, as well as a reduced seizure threshold [21]. The second encodes synapsin II (Syn2); seizures precipitated by sensory stimuli were found previously in Syn2-knockout mice [22]. The third encodes N-ethylmaleimide-sensitive factor (Nsf), which regulates exocytosis in synaptic transmission, as well as AMPA receptor trafficking [23,24]. The fourth encodes α-synuclein (Snca), a neuron-specific presynaptic protein [25]. All four molecules have been found in the pyramidal neurons in the hippocampus where Brunol4 is highly expressed [25–28]. In particular, the expression of NSF [27] and synapsin II [28] were enriched in the CA2-CA3 region, similar to that of Brunol4.
After confirming the differential expression of all four transcripts in Ff/Ff-homozygous newborn mice (unpublished data), expression levels were assessed in adult Ff/+ heterozygous brain regions. All four RNAs had, on average, a 20%–25% reduction in the B6-Ff/+ hippocampus compared with controls (Figure 5A and 5B). Moreover, these candidates showed a significant reduction at the protein level in adult hippocampus of B6-Ff/+ mice (30%–39%; unpublished data), and on a mixed background in Ff/+ and Ff/Ff, (25%–32% Ff/+; 32%–56% Ff/Ff, Figure 5C and 5D). These region-specific decreases before the onset of seizures were consistent with the limbic-seizure phenotype and the overlapping hippocampal expression of the four genes with that of Brunol4.
Figure 5
Reduced Expression of Four Candidate Genes in Ff /+ Brains
(A) Representative images from three independent RNA blots. Sequential hybridization using Htr2c, Nsf, Snca, Syn2, Gabrb3, and β-actin probes was carried out on a blot containing poly-A RNA extracted from dissected brain regions. The synapsin II probe detected both synapsin IIa and IIb transcripts due to alternative splicing/polyadenylation [40] and there was no overt change in the synapsin 2a/2b ratio between Ff/+ and +/+ brains. In addition to the loading control actin, we examined the expression of Gabrb3, which encodes the β3 subunit of the GABA(A) receptor. Germline targeting of Gabrb3 caused perinatal lethality in homozygous mice and overt seizures in heterozygotes [41]. No significant expression difference was found between Ff/+ and +/+ brains. The position of size markers (Ambion) was shown on the left. CO, cerebral cortex; Hb, hindbrain; Hi, hippocampus. F, Ff/+; +, +/+.
(B) Histogram summarizing results from northern blots, expressed as percent of wild type after normalization to β-actin.
(C) Reduced expression at the protein level; Western blot for HTR2A, HTR2C, NSF, synapsin II, α-synuclein, and β-tubulin. HTR2A, a molecule with distribution and property similar to HTR2C, was used as a control in addition to β-tubulin. Representative images from experiments with three independent adult mice on a mixed background are shown.
(D) Histogram summarizing results from (C) in Ff/+ and Ff/Ff mutants on a mixed background, expressed as % of wild type after normalization to β-tubulin, summarizing data from three independent animals. Error bars throughout are one standard deviation.
Htr2c Deficiency Contributes to, but Is Not Sufficient for, Seizure Phenotypes in Brunol4Ff Mutants
Null mutants of the Htr2c receptor gene have several phenotypic similarities with Brunol4Ff/+ heterozygotes, including reduced seizure threshold, hyperactivity, and late-onset weight gain [21,29]. However, the fact that Brunol4Ff/+ mice on the B6 strain background experience frequent handling-provoked convulsive seizures after 3 mo of age, whereas Htr2c null mutants do not (Y. Yang, W. Frankel, unpublished data), suggests that down-regulating Htr2c receptor expression is not sufficient to account for the handling-induced seizures. To determine whether Htr2c deficiency is sufficient to account for the ECT of Brunol4Ff/+ mice, we compared seizure threshold in wild-type, single-mutant, and double-mutant mice (Figure 6). These studies were all done on the B6 strain, to avoid the confounding effect of genetic background. Although the average seizure threshold of Htr2c null mutants was lower than that of Brunol4Ff/+ mice by approximately 2 mA, the threshold of double mutants was 1 mA lower still (p = 0.0003; |t|-test). This suggests that factors in addition to reduced expression of Htr2c contribute to seizure threshold in Brunol4Ff/+ mice. Another difference between Htr2c and Brunol4 mutant mice is that Htr2c-null mutants do not have SWD ([21]; our unpublished observations). However, we found that blocking serotonin reuptake in Brunol4Ff/Ff homozygotes by fluoxetine (Prozac) treatment lowers the SWD incidence by about 50% (Figure 1E, right)—again suggesting that Htr2c down-regulation combines with other Brunol4-downstream deficiencies to cause the seizure disorder of frequent-flyer mutants. Although further tests are required to determine whether the other causative factors are Syn2, Nsf, or Snca specifically, it is clear that the seizure phenotypes of Brunol4Ff are determined in a genetically complex manner.
Figure 6
Genetic Interaction between Brunol4 and Htr2c for ECT
Shown is a histogram of average threshold (in mA) to a minimal clonic or maximal tonic hindlimb extension seizure, determined as described in Materials and Methods, for wild-type (+/+, +/+), single mutant (Ff/+, +/+ or B6-Htr2ctm1Jul/Y), or double mutant (Ff/+; Htr2ctm1Jul/Y), littermates from crosses between B6-Htr2ctm1Jul/Y and B6-Ff/+ mice.
Discussion
Here, we report on the causal association between Brunol4, encoding a brain-specific RNA-binding protein, and the seizure disorder of frequent-flyer mouse mutants. The origin of the disorder is a transgenic insertion in the Brunol4 gene, resulting in very little Brunol4 transcript in homozygotes and accordingly reduced amount in heterozygotes—suggesting haploinsufficiency. We do not know why the transcript levels were very low, but one possibility is due to the inverted repeat in the transgene cluster, creating a potential hairpin structure that may prevent read-through transcription (Figure 2). Brunol4Ff mutant mice have several different kinds of seizures, depending upon genotype, age, and strain background. In heterozygotes on a B6 inbred strain background, these include recurrent limbic and tonic–clonic seizures—observed readily following routine animal handling after 3 mo of age—and a significantly lower ECT at an earlier age. Although homozygotes do not usually survive on a C57BL/6J strain background, on F2 hybrid backgrounds homozygotes (and some heterozygotes) that survived also displayed spike-wave discharges, the hallmark of absence epilepsy.
The prospect of a defective RNA-binding protein such as BRUNOL4 causing a complex seizure disorder suggests a way in which a single gene defect can mimic a complex genetic disease, by impairing the function of multiple molecules simultaneously. This could happen either as a direct consequence of its absence, or secondarily, e.g., a cascade of effects. Microarray analysis between homozygous mutants and coisogenic control brain yielded a small number of down-regulated transcripts that were statistically significant, two of which (Htr2c and Syn2) are already known to cause seizure-related phenotypes when knocked out in mice [21,22], and two other (Nsf and Snca) have obvious functions that relate to synaptic transmission. The down-regulation of each was confirmed in adults, and was greatest in the hippocampus, where Brunol4 expression is high. Interestingly, in addition to seizure susceptibility, Ff/+ heterozygotes display two nonseizure phenotypes like those of Htr2c null mutants: mild hyperactivity and late-onset obesity [21,30,31]. This might suggest that compromised serotonergic transmission is the major factor of the frequent-flyer phenotype. However, because Htr2c expression is reduced only modestly (∼25% RNA, ∼35% protein) in Ff/+ heterozygotes, compared with complete loss in Htr2c-null mice, it seems more likely that a combination of compromised serotonergic transmission and other defects is responsible for the disorder. Further evidence for this idea was obtained in the ECT paradigm (additive phenotypic effects of double mutants), and by observing that SWD phenotype of Brunol4 homozygotes was partially mitigated by up-regulation of serotonergic transmission through blocking the reuptake of serotonin. With any of the seizure paradigms, it is plausible that reduced synaptic efficacy, e.g., due to reduced expression of the other three candidates singled out—Syn2, Nsf, or Snca—is the other contributing variable. However, we cannot ignore other genes misregulated in Brunol4 mutants, some with unknown function (Table S1), since many also expressed selectively within the hippocampus in B6 mice (Allen Brain Atlas [32]).
We do not know why Brunol4 deficiency results in the decreased expression of these and other transcript RNAs. BRUNOL4 has been shown to regulate alternative splicing in cells and tissues without endogenous BRUNOL4 [33–35], but we did not detect aberrant splice variants in mutant brains, at least in the subset of transcripts that we analyzed (our unpublished results). Since members of the Bruno gene family are involved in RNA editing, stability, and translation, the possibility exists that Brunol4 is involved in other aspects of RNA metabolism, for example, in stabilizing transcripts for translation, a possibility that is supported, in part, by the fact that the degree of reduction at the RNA level and at the protein level was not 100% concordant in the mutant hippocampus (∼20%–25% and ∼31%–39%, respectively). However, since many RNA processing steps are believed to be coupled with transcription [36], BRUNOL4 may indeed serve multiple roles in RNA metabolism.
Most genes known to cause idiopathic epilepsy encode ion channels. Brunol4 joins a growing list of non-ion-channel epilepsy genes in both humans and mice [3–5,7]. It is noteworthy that the human BRUNOL4 gene is in a region on human Chromosome 18 showing strong evidence for linkage with adolescent-onset idiopathic generalized epilepsy [37], suggesting that BRUNOL4 may be a candidate gene for these seizure disorders.
Materials and Methods
Mice.
Origin of Brunol4Ff mice. C57BL/6J (B6) transgenic mice were generated at The Jackson Laboratory (http://www.jax.org/) using a construct where the expression of murine Ighmbp2 cDNA and enhanced green fluorescent protein (EGFP) was driven by a bidirectional tetracycline-responsive promoter. Briefly, the coding region of the Ighmbp2 cDNA was cloned in the pBI-EGFP vector (Clontech, http://www.clontech.com/). The PvuI linearized transgene (∼8.3 kb) was microinjected into pronuclei of single cell B6 mouse embryos, which were subsequently implanted into pseudopregnant mice. B6.129-Htr2ctm1Jul mice, derived from mice published in 1995 [21], were obtained from The Jackson Laboratory's Induced Mutant Resource and are now fully congenic on the B6 background after backcrossing for ten generations. All animals were fed standard National Institutes of Health diet containing 6% fat and acidified water ad libitum. All animal procedures followed Association for Assessment and Accreditation of Laboratory Animal Care guidelines and were approved by institutional Animal Care and Use Committee.
ECT.
As previously described [38], mice were restrained, a drop of anesthetic containing 0.5% tetracaine and 0.9% NaCl was placed onto each eye, and a preset current was applied via silver transcorneal electrodes using a electroconvulsive stimulator (Ugo Basile model 7801; http://www.ugobasile.com). The stimulator was set to produce rectangular wave pulses with the following parameters: 299 Hz, 0.2 s duration, 1.6 ms width. Sixty Ff/+ and 57 littermate +/+ male mice (ages 6–9 wk) were tested for ECT over a range of electric current settings for minimal clonic forebrain seizure and each ECT response was recorded. The data were analyzed in the computer program MiniTab (Minitab, http://www.minitab.com/) and a response curve was generated using the log-Probit procedure. To determine ECT in double mutants, male mice (ages 6–9 wk) were tested by increasing the stimulus once daily until at least a minimal clonic seizure was observed, and the average threshold determined for each genotype.
Electroencephalogram analysis.
Mice were anesthetized with tribromoethanol (400mg/kg i.p.) Small burr holes were drilled (1 mm anterior to the bregma and 2 mm posterior to the bregma) on both sides of the skull 2 mm lateral to the midline. Electroencephalogram (EEG) activity was measured by four Teflon-coated silver wires soldered onto a microconnector. The wires were placed between the dura and the brain and a dental cap was then applied. The mice were given a post-operative analgesic of carprofen (5 mg/kg subcutaneous) and were given a 48-h recovery period before recordings were made. The mice were recorded for a 2-h period on each of the following two days using the Grass EEG Model 12 Neurodata Acquisition System and PolyViewPro software program (Grass-Telefactor, http://www.grasstechnologies.com/). For mice that were treated with ethosuximide (200 mg/kg; Sigma-Aldrich, http://www.sigmaaldrich.com) or fluoxetine (20 mg/kg, Sigma-Aldrich), on the day following their second standard EEG recording, mice were recorded for 90 min and then injected intraperitoneally. They were then recorded for a minimum of one additional hour. The control mice were injected intraperitoneally with saline and recorded in the same manner. Matched pairs tests were done using the program JMP 6.0.3 (SAS, http://www.sas.com/).
Identification of the transgene insertion site.
Genomic DNA from transgenic mice and control mice was digested with BclI, SpeI, BglII, SphI, and StuI and electrophoresed and blotted onto a Nytron Plus membrane (Schleicher & Schuell, http://www.whatman.com/). The blot was probed with an EGFP probe and a unique transgene-genomic junction fragment was present in StuI-digested DNA at about 3 kb. The StuI-digested fragment of ∼3 kb was cloned into the pBluescript II-SK vector (Stratagene, http://www.stratagene.com). A vector-specific primer and an EGFP primer were used to amplify the junction fragment. Automated sequencing confirmed the presence of the EGFP cDNA as well as other vector sequence. A 652-nt fragment did not match with the pBluescript II-SK vector sequence and was used as a query to BLAST search the mouse genome. A single perfect match to intron 1 of the Brunol4 gene was found on mouse Chromosome 18. The other breakpoint of the transgene insertion was cloned by a PCR strategy using a transgene-specific primer and a Brunol4 intron 1 primer. Based on the sequence information around the insertion breakpoints in the Ff allele, a 3-primer PCR assay was designed to detect the Ff allele and wild-type allele. Primers for this assay are: s3gtf, 5′-CTCTTCATCCCTTCTGGCAAGTAG-3′; s3gtr, 5′-GTATTCAACAATTCCGTGTCGCCC-3′; and s3gtr2, 5′-CCACACAGAGACCAAGAAGATTCC-3′. At 55 °C annealing temperature, 35 cycles, standard PCR conditions, the s3gtf/s3gtr2 primers produce a 672-bp wild-type allele, and the s3gtf/s3gtr primers produce a 464-bp mutant allele.
Northern blot analysis and quantification.
Total RNA was prepared from newborn brain, adult brain and dissected brain regions using TRIzol reagent (Invitrogen, http://www.invitrogen.com). Two probes were generated for Brunol4, p1 containing 5′ UTR and exon1 5′ to the insertion site and p2 containing the region between BRUNOL4′s second and third RNA recognition motif. Both probes detected a single transcript on northern blots and p2 was also used for in situ analysis. Hybridization was carried out in formamide-based solution at 42 °C overnight and the blot was washed and exposed to an X-ray film at −80 °C. The same blot was stripped and reprobed with a mouse β-actin probe. Films were imaged by Fuji Luminescent Image Analyzer LAS-1000 Plus (http://fujifilmlifescienceusa.com) and subsequently quantified by Fuji Image Gauge Ver. 3.4.
Real-time reverse-transcription PCR.
Total RNA was prepared from the cerebral cortex of adult B6-Ff/+ and and B6-+/+ /littermates with Trizol (Invitrogen, http://www.invitrogen.com) and treated with DNase I (Promega, http://www.promega.com) under the manufacturer's suggested conditions. RNA (2 μg) was reverse transcribed with AMV reverse transcriptase (Promega). The cDNA was diluted 20-fold, and 1.5 μl was added to qPCR Mastermix Plus for SYBR Green I (Eurogentec, http://www.eurogentec.be) with pairs of the following primers: beta-actinF (5′-CATTGCTGACAGGATGCAGAA-3′) and beta-actinR (5′-GCCACCGATCCACACAGAGT-3′), Be1u (5′-TCGCAGTAGGTGAGGAAAGCGCAG-3′) and Be2d (5′-TCGCAGTAGGTGAGGAAAGCGCAG-3′), corresponding to Brunol4 exon 1 forward and exon 2 reverse, respectively. The PCR reactions were analyzed on an ABI Prism 7000 Sequence Detection System (PerkinElmer, http://www.perkinelmer.com/). The PCR amplifications from three pairs of age-matched mice were run in triplicate. Amplification of the correct size products was confirmed by agarose gel electrophoresis.
RNA in situ hybridization.
Thin brain sections (7 μm) from 10-wk-old B6 male mice were hybridized with 33P probes overnight at 50 °C and washed, RNase A treated, dehydrated, and air dried. Slides were dipped in liquid emulsion (Kodak, http://www.kodak.com/) and images were developed 5 d afterwards. For DIG-based in situ analysis, 15-μm cryosections were hybridized with DIG probes overnight at 65 °C and washed extensively before an overnight incubation of alkaline phosphatase–conjugated anti-DIG antibody (1:2,000, Roche, http://www.roche.com/). Staining signal was developed using BM purple (Roche) at room temperature for 12 h.
Microarray analysis.
Total RNA was prepared from six male newborn heads (3 Ff/Ff and 3 +/+). 10 μg of total RNA was used to generate 15 μg of cRNA for hybridization to the Affymetrix 430 v2.0 Gene Chip (Affymetrix, http://www.affymetrix.com/) according to manufacturer's recommendation. Using the R/maanova package [39], an analysis of variance (ANOVA) model was applied to the data, and F1, F2, F3, and Fs test statistics were constructed along with their permutation p-values. Changes in 459 transcripts were considered statistically significant among the 39,000 transcripts interrogated.
Antibodies for Western blotting.
The primary antibodies and the dilutions were: α-HTR2A, 1:500 (BD Pharmingen, http://www.bdbiosciences.com/); α-HTR2C, 1:500 (Immunostar, http://www.immunostar.com/); α-NSF, 1:1,000 (H-300, Santa Cruz Biotechnology, http://www.scbt.com/); α-Synapsin 2, 1:5,000 (Stressgen, http://www.nventacorp.com/); α-alpha-synuclein, 1:250 (BD Transduction, http://www.bdbiosciences.com/); and α-beta-tubulin, 1:1,000 (Sigma). The secondary antibodies and the dilutions were: HRP α-mouse, 1:6,000 (Zymed, http://www.invitrogen.com/) and HRP α-rabbit, 1:2,000 (PerkinElmer).
Northern blot probes.
The following probes were designed to detect expression differences among the putative BRUNOL4 target RNAs between Ff/+ and +/+ brains: (1) Htr2c, 437-nt probe in exon 6, the last nucleotide is 43 nt 3′ of the TAA stop codon. (2) Nsf, 470-nt probe as described in [27]. (3) Syn2, 234-nt probe in exon 1, the first nucleotide is 90 nt 3′ of the ATG start codon. This probe was able to detect the two alternatively polyadenylated Syn2 transcripts. (4) Snca, 416-nt probe in exon 6, the first nucleotide is 37 nt 3′ of the TAA stop codon. (5) Gabrb3, 412-nt probe specific to the 3′ end of the coding sequence, the last nucleotide is 14 nt 5′ of the TGA stop codon.
Western blot.
Hippocampi were dissected from three male Ff/+ mice (before the onset of handling-provoked seizures) and three +/+ littermates. Protein extracts were made using RIPA buffer with proteinase inhibitors (Roche) and subsequently quantified using the Bradford reagent (Bio-Rad, http://www.bio-rad.com/). Protein (50 μg) from each sample was loaded and probed with the primary antibody and a secondary peroxidase-conjugated antibody and visualized with the ECL plus kit (Amersham, http://www.amersham.com/). The nitrocellulose membrane was incubated with Restore Western blot stripping buffer (Pierce, http://www.piercenet.com/) at 37 °C for 30 min to remove all the antibodies. The membrane was washed and subsequently reprobed with a different set of antibodies. Signals were quantified using the method described above.
Supporting Information
Table S1
List of Genes with Differential Expression between Ff/Ff Homozygous and Wild-Type Newborn Mice
Microarray analysis was carried out as described in Materials and Methods. Down-regulated transcripts (94) are listed first, followed by 365 up-regulated transcripts.
(416 KB XLS)
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Accession Numbers
The National Center for Biotechnology Information (NCBI) Nucleotide database (http://www.ncbi.nlm.nih.gov/sites/entrez?db=Nucleotide) accession number for intron 1 of the Brunol4 gene on mouse Chr 18 is EF639873.
Acknowledgements
We thank Jason Affourtit, Sandy Daigle, Barbara Beyer, Carolyne Dunbar, Jonette Gilley, Neena Haider, Sonya Kamdar, Yong Woo, and Weidong Zhang for technical assistance. We also thank Susan Ackerman, Albert Becker, Robert Burgess, and Verity Letts for comments and advice.
Author contributions. YY and WNF conceived and designed the experiments and wrote the paper. YY, CLM, NB, and WNF performed the experiments and analyzed the data. TPM and GAC contributed reagents/materials/analysis tools.
Funding. The Jackson Laboratory's Gene Expression and DNA Sequencing services were subsidized by an NCI core grant CA34196. This work was supported by a grant from the National Institutes of Health (NS31348 to WNF). YY was supported by a TJL Postdoctoral Fellowship and a research award from Citizens United for Research in Epilepsy (CURE).
Abbreviations
Brunol4, - Bruno-like 4
ECT - electroconvulsive threshold
EEG - electoencephalogram
Ff, - frequent-flyer
RT-PCR - reverse-transcriptase PCR
SWD - spike-wave discharge
Footnotes
Competing interests. The authors have declared that no competing interests exist. | [
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