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Degraded Carrageenan Causing Colitis in Rats Induces TNF Secretion and ICAM-1 Upregulation in Monocytes through NF-kappaB Activation
Carrageenan (CGN) is a high molecular weight sulphated polysaccharide derived from red seaweeds. In rodents, its degraded forms (dCGN) can induce intestinal inflammation associated with macrophage recruitment and activation. The aim of this study was: 1) to analyze the size-dependent effects of dCGN on colon inflammation in vivo, and 2) to correlate these effects with monocyte/macrophage proliferation, cytokine production and expression of various cell surface antigens including ICAM-1 adhesion molecule. Peripheral blood monocytes (PBM) and THP-1 monocytic cells were cultured in the presence of either 10 or 40 kDa, dCGN. The 40 kDa, but not the 10 kDa dCGN, induced colitis in in vivo. Degraded CGN inhibited THP-1 cell proliferation in vitro, arresting the cells in G1 phase. In addition, dCGN increased ICAM-1 expression in both PBM and THP-1 cells with a major effect seen after 40 kDa dCGN exposure. Also, dCGN stimulated monocyte aggregation in vitro that was prevented by incubation with anti-ICAM-1 antibody. Finally, dCGN stimulated TNF-alpha expression and secretion by both PBM and THP-1 cells. All these effects were linked to NF-kappaB activation. These data strongly suggest that the degraded forms of CGN have a pronounced effect on monocytes, characteristic of an inflammatory phenotype.

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Carrageenan (CGN) is a high molecular weight sulphated polysaccharide (>200 kDa) derived from red algae (Rhodophyceae). Three main forms of CGN have been identified: kappa, iota, and lambda. They differ from each other in sulphation degree and solubility [1], [2]. Native CGN is thought to be harmless and is widely used as a food additive to improve texture. It is also used in cosmetics and pharmaceuticals. However, acid treatment at high temperature (80degreesC) triggers CGN hydrolysis to lower molecular weight (<50 kDa) compounds known as poligeenan or degraded CGN (dCGN). These dCGNs induce inflammation and have been widely used as models of colitis in several species, including rats [3], rabbits [4] and guinea pigs [5]. The role of dCGN as a tumor-promoting factor remains controversial [4], [6]-[8].
Although the native form is thought to be harmless for human consumption, small amounts of dCGN are probably produced by acid hydrolysis during gastric digestion [9], [10] or interaction with intestinal bacteria [11], [12]. Whereas the effects of native and dCGN on intestinal inflammation have been extensively analyzed in animal models, only few studies have been conducted using human cell lines. Recent studies have shown a link between exposure to native form CGN and IL-8 production by the human intestinal epithelial cell line, NCM460, via Nuclear Factor-kappaB (NF-kappaB) activation [13], [14]. NF-kappaB is a transcription factor that regulates the expression of genes associated with inflammation [15], [16].
Macrophage infiltration and accumulation is a common characteristic of intestinal diseases [17]. Macrophages represent 10% of total lamina propria cells, secrete a wide range of biologically active compounds and express cell-adhesion molecules. The immune cell response to an inflammatory stimulus seems to be amplified or directly generated by cells exposed to sulphated polysaccharides such as carrageenans. Indeed, inflammation induced by dCGN was associated with recruitment of macrophages to inflammation sites [18], [19]. Also, inflammation induced by Dextran Sulphate Sodium (DSS), another sulphated compound, was directly associated with macrophages recruitment [20], since DSS still provoked inflammation after T-lymphocyte and NK cell depletion [20]. Although inflammation can be induced by dCGN, there are no data on human monocyte responses to dCGN exposure. Therefore, to investigate the effects of dCGN on human monocytes, normal Peripheral Blood Monocytes (PBM) and tumoral monocyte/macrophage THP-1 cells were exposed to 10 kDa and 40 kDa dCGN. We found that dCGN inhibited THP-1 cell proliferation in vitro, increased ICAM-1 expression, stimulated ICAM-1-dependent monocyte aggregation, and stimulated TNF-alpha expression and secretion. These responses were more pronounced after 40 kDa dCGN exposure and were linked to NF-kappaB activation. In addition, the 40 kDa dCGN, but not the 10 kDa dCGN induced in vivo colitis as shown by the inflammatory response in the rat colon. These results suggest that the degraded forms of CGN have an important effect on monocytes resulting in an inflammatory phenotype.

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Preparation of Degraded Carrageenan
Two preparations of degraded carrageenan with low, (~10 kDa; C10), and medium, (~40 kDa; C40) molecular weight were prepared from native iota-carrageenan extracted from Euchema spinosum (generously provided by Sanofi Biosystems Industry, Boulogne-Billancourt, France). Native carrageenan was dissolved in distilled water (5% w/v) under vigorous stirring and heated to 60degreesC. Then, the carrageenan solution was submitted to two different treatments to obtain both low and medium molecular weight fractions. Briefly, for the low molecular weight fraction, carrageenan solution was hydrolyzed with 0.3% (v/v) concentrated sulphuric acid for 15 min at 80degreesC. After neutralization with NaOH 4N, the solution was ultra filtered through a hollow fibre cartridge with MW cut-off 5 kDa, (Amicon Inc, Beverly, USA). For the medium molecular weight fraction, the carrageenan solution was hydrolyzed with 0.3% (v/v) concentrated sulphuric acid for 30 min at 60degreesC. After neutralization, the supernatant was ultra filtered (MW cut-off 100 kDa). The filtrate was submitted to a second ultra filtration (MW cut-off 5 kDa). Both preparations of dCGN were precipitated with 4 volumes of 95% ethanol, dried at room temperature and ground to small particles (1 mm in diameter). Using gel-permeation chromatography in combination with light scattering measurements (see Viebke et al. [21]), it was confirmed that the low fraction had an average molecular weight of 10 kDa, and the medium fraction of 40 kDa. The sulphate content of polysaccharides in both fractions was measured following the method of Quemener et al. [22]. Finally, the absence of polysaccharide structure modifications in the two fractions was confirmed using 2H-NMR spectroscopy. The absence of LPS contamination in the two fractions was confirmed using the e-Toxate(R) kit (Sigma, St Quentin Fallavier, France). Before use in cell culture, the two fractions were dissolved in complete medium during 30 min at 56degreesC.


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Animals, Chemicals and Diet
Male Wistar rats (150 g average weight) were housed under standard conditions and fed ad libitum with standard rodent laboratory chow. Degraded iota-carrageenans were administered in the drinking water (5% w/v) for 55 days to 2 groups of six animals each. The first group received the low molecular weight carrageenan (10 kDa dCGN) and the second received the medium molecular weight carrageenan (40 kDa dCGN). An additional group of four rats were maintained on regular tap water (control group). To increase palatability 0.2% sucrose was added to the drinking water of all groups (Van der Waaji et al., [23]). Fresh carrageenan solutions were prepared daily.


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Evaluation of Colitis
Body weight, liquid and food consumption, diarrhea and rectal bleeding (detected by eye inspection) were recorded throughout the feeding period. After 55 days, animals were sacrificed by cervical dislocation. The length of the colon was measured as described by Okayashu et al. [24]. Then, each colon was ligated in sections of 2 cm and 1 to 2 ml of 10% formalin was infused into the intestinal lumen. The moderately distended segment was sectioned and fixed in 10% formalin. The following day, the intestinal content was removed by vortexing. The fixed segment was kept in 10% formalin at 4degreesC until the paraffin embedding procedure. To evaluate the degree of inflammation, this segment of colon was opened longitudinally and macroscopic and histological scores of inflammation were recorded as previously described [25], [26]. The toluidine blue staining was used for identification of sulphated polysaccharides in the intestinal mucosa. On the day of sacrifice, a fresh sample of each colon (50 mg) was collected for myeloperoxidase (MPO) assay according to Krawisz et al., [27]. The level of MPO, mainly expressed by neutrophils, indicates the rate of recruitment of neutrophils to the intestinal mucosa. One unit of MPO activity corresponds to the degradation of 1 micromol of peroxide per minute at 25degreesC.

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Cell Culture
All tissue culture reagents were from Invitrogen (Cergy Pontoise, France). THP-1 human monocytic cells were maintained in RPMI-1640 supplemented with 10% FCS, 2 mM L -glutamine, 50 U/ml penicillin and 50 mg/ml streptomycin at 37degreesC in a 5% CO2 incubator. Human peripheral blood mononuclear cells were obtained from heparinized blood by Ficoll-Hypaque density gradient. Monocytes were then isolated by adherence to culture flasks as described [28]. For cell aggregation, monocytes were cultured in the presence or absence of C10 or C40 for 72 h. Cell colonies were monitored under an inverted phase contrast microscope coupled through a video camera to a computer. In some wells, neutralizing monoclonal antibody to ICAM-1 (2.5 microg/ml) (Tebu, Le Perray en Yvelines, France) was added.


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Cell Cycle Analysis
THP-1 cells in exponential growth phase were exposed to complete medium in the presence or absence of carrageenans for 24 h before being stained with propidium iodide using the DNA-Prep Coulter kit according to the manufacturer's instruction (Beckman-Coulter, Villepinte, France). Cell DNA content was then analyzed by flow cytometry using an EPICS XL2 (Beckman-Coulter). Raw data for the distribution of DNA content of 30,000 cells retrieved from the cytometer were expressed as the percentage of G0/G1 through G2/M populations. Multicycle AV software (Phoenix Flow Systems, San Diego, CA) was used to generate DNA content frequency histograms and facilitate data analysis.


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Cell Surface Antigen Expression Analysis
Peripheral Blood Monocytes or THP-1 cells were exposed to complete medium in the presence or absence of carrageenan for 36 h. After two washes in PBS without Ca2+ and Mg2+, cells were incubated in PBS containing 0.1% gelatin and 8% AB human serum to prevent binding to Fc receptors. Then, 5x105 cells were incubated with primary antibodies at 4degreesC for 30 min. Two other washes in PBS preceded incubation with FITC-conjugated goat antibody anti-mouse IgG diluted 1/1000 at 4degreesC for 30 min (Tebu). After two additional washes, analysis of stained cells was performed on an EPICS XL2 (Beckman-Coulter). The cell population was gated according to its forward and wide-angle light scattering. Data were expressed as mean relative fluorescence intensity (MFI) of 3000 cells.


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TNF Activity Bioassay
Monocytes or THP-1 cells were cultured with or without different concentrations of CGNs or LPS (Salmonella typhosa, Sigma) for 24 h or the indicated time. Biologically active TNF-alpha/beta in tissue culture supernatant was measured using the WEHI 164 clone 13-cell killing assay [29]. TNF concentrations are expressed as pg/ml.


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RT-PCR Analysis
Total RNA from monocytes was isolated using TRIzol Reagent(TM) (Invitrogen). cDNA was generated on 1 microg of total RNA in a reaction volume of 20 microl, using M-MLV reverse transcriptase (Invitrogen). PCR was done in the linear range of amplification (determined for each primer pair-cDNA combination). Standard PCR reactions were performed with 1 microl of the cDNA solution, 50 microM of each primer solution, 10 mM of each dNTP, 25 mM MgCl2, 10X Goldstar DNA polymerase reaction buffer, and 0.5 units of Goldstar DNA polymerase (Eurogentec, Seraing, Belgium). First PCR cycle consisted of 1 min at 92degreesC, 1 min at 58degreesC and 1 min at 72degreesC; then each PCR cycle consisted of 40 sec at 92degreesC, 40 sec at 58degreesC and 50 sec at 72degreesC. cDNA for beta-actin was amplified for 28 cycles using the oligos: sense 5'-GGCATCGTGATGGACTCCG-3' and antisense 5'GCTGGAAGGTGGACAGCGA-3'. cDNA for TNF-alpha was amplified for 35 cycles using the oligos: sense 5'-AAGCCTGTAGCCCATGTTGT-3' and antisense 5'-CAGATAGATGGGCTCATACC-3'. cDNA for ICAM-1 was amplified for 35 cycles using the oligos sense 5'-GTAGCAGCCGCAGTCATAATGG-3' and antisense 5'-A TGCTGTTGTATCTGACTGAGG-3'.


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NF-kB Transcription Reporter Gene Assay
The plasmid 3XMHC-luc (a generous gift from Drs. J. Westwick and D.A. Brenner, University of North Carolina, Chapel Hill) contains three copies of NF-kappaB-responsive element from the MHC class I locus, placed upstream of the luciferase gene. Human monocytic THP-1 cells were transiently transfected as previously described [30], and then cultured for 4 h alone or with increasing concentration of either C10 or C40. Luciferase activity was determined using a luminometer (Monolight 2010 Luminometer, Ann Arbor, MI).


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Western Blot Analysis
THP-1 cells were stimulated for various lengths of time with 0.1 mg/ml C10 or C40, or 10 microg/ml LPS. Cells were then pelleted, washed and homogenised in lysis buffer (10 mM Hepes, pH 7.9, 150 mM NaCl, 1 mM EDTA, 0.6% NP-40, and 0.5 mM PMSF) on ice. Homogenates were sonicated, centrifuged at 10,000 rpm to remove cellular debris, and supernatant collected. Protein concentration was determined using the DC Protein Assay (Bio-Rad). Proteins in samples (15 microg total proteins) were resolved in a denaturing 12% polyacrylamide gel and transferred to a nitrocellulose membrane. I-kappaBalpha protein was detected using a rabbit polyclonal antibody (Santa Cruz Biotechnology, CA) followed by a horseradish peroxidase-coupled goat polyclonal antibody against rabbit Ig (Caltag Laboratories). Finally, IkappaB bands were revealed using the ECL(TM) detection system (Amersham Pharmacia Biotech, Les Ullis, France) according to the manufacturers' instruction. Antibody to alpha-Tubulin (Santa Cruz) was use as loading control.
For nuclear NF-kappaB, THP-1 cells were stimulated with 1 mg/ml C10 or C40 for 30 minutes at 37degreesC. Cells were then pelleted and nuclei separated as described [31]. Nuclei were washed and homogenized directly in loading (Laemli) buffer and heated for 5 minutes at 100degreesC. Proteins in samples were resolved in a denaturing 8% polyacrylamide gel and transferred to a polyvinylidine fluoride (PVDF) membrane (Immobilon-P; Millipore, Bedford, MA). Membranes were incubated in blocking buffer (1% BSA, in PBS) for two hours at room temperature. Membranes were subsequently probed with the corresponding antibody in blocking buffer, overnight. Rabbit polyclonal antibody anti-NF-kappaB p50 subunit (# sc-114) or anti-NF-kappaB p65 subunit (# sc-109) from Santa Cruz Biotechnology were used. Membranes were washed six times in PBS with 0.05% Tween 20, 5 minutes each time, and incubated with a 1/3000 dilution of HRP-conjugated F(ab')2 goat anti-rabbit IgG in 5% nonfat dry milk and 0.05% Tween 20 in PBS for 1 hour at room temperature. After washing six more times in PBS with 0.05% Tween 20, antibody-reactive proteins were detected using a chemiluminescence substrate (SuperSignal; Pierce, Rockford, IL) according to the manufacturer's instructions. To confirm that equivalent amounts of protein were loaded in each line, membranes were also Western blotted for ERK as described [32].


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Analysis of NF-kappaB Activation by Flow Cytometry
Nuclear activation of NF-kappabeta by flow cytometry was performed as described [31].


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Statistical Analysis
The results were expressed as the mean value +/- S.E.M. of individual experiments. The statistical significance of the differences between mean values was assessed by the Student's t-test and analysis of variance (ANOVA).


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Degraded CGN Induce Colonic Inflammation
All rats developed diarrhea during degraded carrageenan administration and gross evidence of blood was frequently detected in the stools. Colon length dramatically decreased in all treated rats with a more pronounced effect being observed in the 40 kDa dCGN treated group (Fig. 1A). Furthermore, prolonged exposure to 40 kDa dCGN resulted in high macroscopic and histological scores of inflammation (Fig. 1B, C). Only weak myeloperoxidase activity was detected in both control and dCGN-treated groups (Fig. 1D), indicating that granulocytes did not play a major role in the inflammation at that stage. Histological examination revealed various degrees of mucosal inflammation. Rats treated with 10 kDa dCGN showed edema, epithelium atrophy and slight lymphocyte infiltration (data not shown). These symptoms were totally absent in the colon of control rats (Fig. 1E). More severe mucosal injuries including ulceration, hyperplastic epithelium, crypt distortion and a strong macrophage infiltration, were observed in the 40 kDa dCGN-treated rats (Fig. 1F). No sulphated polysaccharides were detected by toluidine blue staining of colon mucosa from rats treated with either the 10 or 40 kDa dCGN (not shown). Although we cannot exclude that dCGN mat not have retained in the section during the histology procedure, this indicates that these polymers may not have been phagocytosed.


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Degraded CGN Induced-TNF-alpha Production by Monocytes In Vitro
In order to study the capacity of dCGN to stimulate TNF-alpha production, peripheral blood monocytes were cultivated in the presence of dCGN (0.1 to 1 mg/ml). Very low levels of TNF-alpha were induced in PBM after stimulation with native CGN (Fig. 2A). Addition of 0.1 mg/ml 10 kDa dCGN resulted in approximately a 60-fold increase in TNF-alpha production by PBM. This was a dose-dependent effect that reached a 180-fold increase when cells were exposed to 1 mg/ml of 10 kDa dCGN (Fig. 2A). A 250-fold increase in TNF-alpha production was detected at 1 mg/ml 40 kDa dCGN (Fig. 2A). TNF-alpha production increased in time reaching a maximum level at 8 hours of culture (Figure 2B). After 24 h, the amount of secreted TNF-alpha was still one third of the total TNF-alpha. Lipopolysaccharide (LPS), a known activator of immune cells also induced TNF-alpha production with similar kinetics as dCGN (Fig. 2B). However, the amount of TNF-alpha produced by LPS was 4-fold less than the one produced by dCGN and it was not detected after 8 hours of culture (Fig. 2B). Similarly, monocytic THP-1 cells cultivated in the presence of variable concentration of dCGN showed an increase in TNF-alpha production (Fig. 2C). This increase in TNF-alpha production was significantly smaller (about 10-fold) than the one presented by PBM (Fig. 2A). No TNF-alpha was released from THP-1 cells exposed to native CGN (not shown). TNF-alpha production by THP-1 cells was not dose dependent to the amount of dCGN used. Also there was no difference between the two forms (10 and 40 kDa) of dCGN (Fig. 2C). Interestingly, TNF-alpha release from THP-1 cells stimulated with dCGN reached a maximum level at 32 h, while stimulation with LPS reached a maximum level at 56 h (Fig. 2D).

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Effect of Native and Degraded CGN on THP-1 Proliferation and Cell Cycle
Preliminary observations by enumeration of THP-1 cells exposed to different concentrations of native and dCGN (10 and 40 kDa) during 2, 5 and 7 days, showed a decline in cell number (data not shown). This suggested that dCGN might cause an alteration in the cell cycle. Cell cycle analysis using flow cytometry showed an accumulation of THP-1 cells in G0/G1 phase, which was associated with a decrease number of cells in the S phase (Fig. 3A). The percentage of cells in the G0/G1 phase was 45.2% for control cells, 62.6% for C10 dCGN (at 2 mg/ml), and 64.2% for C40 dCGN-treated cells (at 2 mg/ml) (Fig. 3B). The effect of dCGN on cell cycle was dose-dependent (Fig. 3B). Neither native nor dCGN had an effect on the number of cells in the G2/M phase (Fig. 3). This effect is not due to cytotoxicity of dCGN even at the highest concentration (i.e. 2 mg/ml) since cell viability was not affected (data not shown).


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ICAM-1 Expression Is Induced by Degraded CGN and Is Responsible for Monocytes Aggregation In Vitro
In order to study the effect of dCGN on the expression of cell surface antigens, PBM and THP-1 cells were incubated for 36 h in the presence and absence of dCGN. The expression of various cell surface molecules was analyzed by flow cytometry as described in materials and methods. Both forms of dCGN clearly stimulated expression of ICAM-1 (CD54) on PBM and THP-1 cells (Fig. 4A). The increase in ICAM-1 expression was higher on THP-1 cells treated with 40 kDa dCGN (Fig. 4B). Another surface antigen, the lymphocyte function-associated antigen 3 (CD58) was slightly reduced on PBM after treatment with 40 kDa dCGN (Fig. 4B). Interestingly, expression of major histocompatibility complex molecules of class I (HLA-ABC) and of class II (HLA-DR), as well as the monocyte marker CD14, seemed to be reduced by treatment with dCGN (Fig. 4B). However, these differences were not statistically significant.
Treatment with dCGN also induced a strong aggregation of monocytes, detected by phase contrast inverse microscopy (Fig. 5). Although this effect was easily observed in monocytes incubated with the 10 kDa dCGN (Fig. 5B), a more robust cell aggregation was observed in monocytes incubated with the 40 kDa dCGN (Fig. 5C). ICAM-1 has been proposed to be the main adhesion molecule responsible for monocyte aggregation. To confirm this, monocytes were incubated with both types of dCGN in the presence of an anti-ICAM-1 antibody, an anti-CD58 antibody and an isotype control IgG1 antibody. The anti-ICAM-1 antibody effectively blocked the cell aggregates induced by dCGN (Fig. 5D, 5E), strongly suggesting that indeed ICAM-1 is responsible for monocyte aggregation. Both the control IgG1 and the anti-CD58 antibody did not modify monocyte aggregation (data not shown).

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Degraded CGN Induce an Increase in ICAM-1 and TNF-alpha mRNA Expression
The increase in surface ICAM-1 expression and TNF-alpha production by monocytes correlated with an upregulation of mRNA for these molecules. Both 10 kDa and 40 kDa dCGN induced a robust increase in mRNA for both ICAM-1 and TNF-alpha (Fig. 6). beta-actin mRNA levels were not affected by dCGN treatment.

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Degraded CGN Induce IkappaB Degradation and NF-kappaB Activation
The expression of genes encoding for ICAM-1 and TNF-alpha is controlled by the nuclear factor NF-kappaB. Site-specific phosphorylation of the inhibitor IkappaB leads to its degradation by proteasome and to a consequential activation of the NF-kappaB pathway. Using a reporter plasmid for NF-kappaB activation, it was confirmed that dCGN induced a strong activation of NF-kappaB, as reflected by an increase in luciferase activity (Fig. 7A). Both forms of dCGN used induced NF-kappaB activation in a dose dependent manner. However, the effect was more strongly induced by the 40 kDa dCGN (Fig. 7A). These results were further confirmed by directly detecting NF-kappaB in the cell nucleus by Western blotting (Fig. 7C) and by FACS (Fig. 7D). These assays also allowed us to determine what NF-kappaB subunits were activated by dCGN. Both forms (10 or 40 kDa) of dCGN induced activation of the p50 and p65 subunits of NF-kappaB. This nuclear factor was present in low levels in the cell nucleus and increased considerably after treatment with dCGN. Western blots suggested the the 40 kDa form of dCGN induced a stronger activation of NF-kappaB (Fig. 7C). A more sentive assay for nuclear factor activation is flow cytometry of nuclei stained with specific antibodies for the nuclear factor of interest. In agreement with the previous data, FACS analysis of nuclei from THP-1 cells showed that there was a basal level of nuclear NF-kappaB (Fig. 7D). Again, both forms (10 or 40 kDa) of dCGN induced an increase of the p50 and p65 subunits of NF-kappaB in the nucleus of these cells. The 40 kDa degraded CGN gave a stronger increase of NF-kappaB (Fig. 7D). These data strongly suggest that the heterodimer p50/p65 is the NF-kappaB isoform activated by degraded CGN in monocytes. In addition, degradation of the inhibitor IkappaBalpha was also observed in cells treated with dCGN (Fig. 7B). No significant IkappaBalpha degradation was detected within two hours of dCGN treatment, but IkappaBalpha was markedly degraded by four hours of dCGN treatment (Fig. 7B). We focused on IkappaBalpha subunit, since it masks the nuclear localisation sequence of p65, it is the most rapidly degraded subunit and the most studied one.

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Inflammation of the intestinal tract is usually associated with infiltration and activation of intestinal macrophages [17]. These macrophages are able to initiate immune responses and can be induced to differentiate into cells that either exacerbate or inhibit the inflammation. Accumulation of different types of leukocytes, including monocytes/macrophages, neutrophils, and lymphocytes in the intestinal mucosa during inflammation, is normally followed by secretion of pro-inflammatory cytokines [33]. Several stimuli can induce these leukocytes to produce and secrete cytokines during inflammation. One of the most potent and known stimuli for leukocyte activation is LPS from Gram-negative bacteria [34]. In addition, other factors are also able to stimulate cytokine secretion from various leukocytes [35], [36]. One such factor is CGN, a high molecular weight sulphated polysaccharide (>200 kDa) derived from red algae (Rhodophyceae) [1], [2]. Native CGN is widely used as a food additive (E 407) to improve texture. It is also used in cosmetics and pharmaceuticals. Although native form CGN (200-800 kDa) has been declared harmless to humans [8], its degraded forms (<50 kDa), also known as poligeenan, are widely used to induce colitis in rodents [3]-[5]. These degraded CGN may also have a possible carcinogenic effect [4], [6]-[8]; however this is still controversial.
Although acid treatment at high temperature (80degreesC) is required for CGN hydrolysis in vitro to lower molecular weight dCGN, it is probable that some dCGN are produced by acid hydrolysis during gastric digestion [9], [10] or interaction with intestinal bacteria [11], [12]. Thus, understanding the mechanisms of dCGN-induced bowel inflammation is of great importance. In this report, we have analyzed the role of human monocytes (PBM and THP-1) in dCGN-induced inflammation.
Preliminary in vivo studies in rats treated with dCGN revealed significant shortening of the large intestine associated with an inflammatory state, i.e. strong infiltration of macrophages to the intestinal mucosa similar to DSS-induced inflammation [20]. Using two fractions of dCGN (10 and 40 kDa), we observed a strong correlation between the severity of the inflammation and the dCGN molecular size, thus confirming the size related inflammation in vivo. This macrophage accumulation was not due to cell proliferation because dCGN inhibited THP-1 monocytes proliferation in vitro. These results are similar to those obtained with human colonic epithelial cells (NCM460 cell line) exposed to native CGN for 1-8 days [37]. Thus, it seems that dCGN promote macrophage infiltration by recruiting new cells to the inflamed intestinal mucosa and not by inducing cell proliferation. In addition, very few polymorphonuclear cells were detected in the mucosa at the time point analyzed as demonstrated by a very low level of MPO in the intestinal tissue.
These results suggest that monocytes might produce cytokines associated with activation into macrophages in response to dCGN. Thus, we analyzed the production of TNF by both PBM and THP-1 cells in response to dCGN. Degraded CGN induced a robust production of TNF by monocytes. The 40 kDa form of dCGN was more potent for monocyte stimulation than the 10 kDa or the native ones. Surprisingly, monocyte activation by dCGN to produce TNF was much stronger than the activation induced by LPS, an inflammatory factor considered to be among the most potent stimuli for leukocyte activation. These results underline the fact that partially degraded forms of CGN have important cellular effects. The amount of TNF secreted by PBM induced with LPS was much larger than the one secreted by THP-1 monocytes. Since monocyte activation by LPS is associated with the presence of the CD14 and TLR4 receptors [38]-[40], the different response observed could be due to the different expression of these receptors in PBM and THP-1 cells. CD14 is not expressed by THP-1 monocytes, but it is expressed by PBM. This fact could explain why THP-1 cells produced a much smaller response to LPS than PBM and also the difference of kinetics in the LPS induced TNF secretion. The peak of TNF response was observed after 8 h stimulation followed by a rapid decrease to baseline at 10 h, whereas on THP-1 cells the peak was not reached until after 56 h. Moreover, the amount of TNF secreted by monocytes induced with dCGN was much larger than the one induced by LPS. On the other hand, LPS and dCGN displayed a very different TNF secretion curve in THP-1 cells. These differences suggest that dCGN and LPS could use different activation mechanisms. It is noteworthy that neutralizing antibody to CD14 only partially (<40%) inhibited dCGN-induced TNF secretion (not shown). TLR4 has been recently identified as a surface membrane receptor for CGN in human colonic epithelial cells [41]. Thus, it is possible that TLR4 is activated by dCGN to induce cytokine secretion by monocytes. We can only speculate that TLR4 may have a higher affinity-binding site for dCGN than for LPS, however, this hypothesis remains to be tested.
Another indicator that dCGN stimulate monocytes leading to a more active phenotype is the fact that surface expression of the adhesion molecule ICAM-1 was enhanced in both PBM and THP-1 cells. The over expression of ICAM-1 caused the activated monocytes to form cell aggregates that were more abundant among cells treated with the 40 kDa dCGN. This correlates with the higher expression of ICAM-1 induced by the 40 kDa dCGN, suggesting that the partially degraded CGN is more biologically active. In addition, the cell aggregates are reminiscent of monocyte aggregates, which form multinucleated giant cells (MGC) in patients with Crohn's disease [42]. These giant cells were not observed in healthy individuals or in patients with ulcerative colitis [42]. Moreover, increased expression levels of ICAM-1 and LFA-3 (CD58) were also detected in monocytes from patients with Crohn's disease [43], [44]. Thus, degraded CGN clearly can activate monocytes to express an increased number of ICAM-1 adhesion molecules, therefore being capable of creating the conditions characteristic of Crohn's disease symptomatology, i.e. PBM accumulation and MGC formation [45], [46].
The NF-kappaB pathway regulates genes responsible for ICAM-1 and TNF-alpha expression. NF-kappaB activation is associated with the degradation of the inhibitor protein IkappaB [47]. Indeed, dCGN induced NF-kappaB activation as shown by degradation of IkappaBalpha, translocation of p65 and p50 sub-units to the nucleus and by activation of an NF-kappaB-responsive luciferase reporter plasmid. Again, a stronger activation of NF-kappaB was induced by the 40 kDa dCGN compared to 10 kDa dCGN, suggesting that the partially degraded CGN is more biologically active. Our data are in agreement with a previous study showing that native CGN also induced activation of NF-kappaB in human colonic epithelial cells [48].
NF-kappaB pathway is often associated to promote cell survival and cancer cell growth, however it has been sometimes reported to behave as a tumor suppressor, arresting cell proliferation [49]. Such relationship between NF-kB activation and cell cycle regulation has been reported in normal human epidermal cells [50]. Indeed, it has been shown that NF-kappaB activation suppressed cdk4 expression, which is necessary for the transition to S phase. Finally, NF-kappaB activation was reported to induce growth arrest in normal human keratinocytes by a mechanims involving the cdk inhibitor p21 [51]. The effects of dCGN on p21 and cdk4 expression remain to be studied.
In these studies, we have demonstrated a direct action of dCGN on monocytes. Monocytes exposed to dCGN acquired an inflammatory phenotype that included higher expression of the adhesion molecule ICAM-1 and TNF-alpha production, via the NF-kappaB pathway. This higher expression of ICAM-1 resulted in formation of cell aggregates similar to those observed in patients with Crohn's disease. We presume that the differential effects of 10 and 40 kDa dCGN to induce these effects on monocytes are tightly linked to their capacity to induce inflammation in vivo. However, in vivo, macrophages do not come in direct contact with the intestinal lumen and are separated by the epithelial barrier. The way by which dCGN may leave the intestinal lumen and cross the epithelial barrier to reach the macrophages is an intriguing open question. One possible explanation resides in the potential of dCGN to "induce" cellular and paracellular injurious effects at the intestinal epithelial cell monolayer [48].
In conclusion, dCGN inhibited THP-1 cell proliferation in vitro, accumulating the cells in the G1 phase of the cell cycle, increased ICAM-1 expression, stimulated ICAM-1-dependent monocyte aggregation in vitro, and stimulated TNF-alpha expression and secretion. These responses were more pronounced following 40 kDa dCGN, and were all linked to NF-kappaB activation. These results suggest that, although CGN is widely used as a food additive, its degraded forms have an important effect on monocytes characteristic of an inflammatory phenotype.

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Input:
Degraded CGN induced colon inflammation in rats.
Histograms showing the effect of degraded CGN on: colon length (A); macroscopic (B) and histological (C) inflammation score of colon; Myeloperoxidase (MPO) activity (D). Control rats (white bars); 10 kDa degraded CGN-treated rats (grey bars); 40 kDa degraded CGN-treated rats (black bars). * p<0.05 from control. ** p<0.01 from control. Histological analysis of colon from control rats (E), and from 40 kDa dCGN-treated rats (F).


Input:
Degraded CGN stimulated TNF secretion from monocytes.
Levels of TNF released from peripheral blood monocytes (A-B) and THP-1 cells (C-D) after stimulation with dCGN. A: TNF release induced by native CGN (open bars), 10 kDa dCGN (grey bars), or 40 kDa dCGN (black bars). B: Kinetics of TNF release induced by nothing (control; black diamonds), 0.1 mg/ml 10 kDa dCGN (black triangles), 0.1 mg/ml 40 kDa dCGN (black squares), or 10 microg/ml LPS (open squares). C: TNF release induced by nothing (control; open bars), 10 microg/ml LPS (hatched bars), or increasing concentrations of 10 kDa dCGN (grey bars), or 40 kDa dCGN (black bars). D: Kinetics of TNF release induced by nothing (control; black diamonds), 0.1 mg/ml 10 kDa dCGN (black triangles), 0.1 mg/ml 40 kDa dCGN (black squares), or 10 microg/ml LPS (open squares).

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Input:
Degraded CGN induced THP1 cell cycle arrest in G1 phase.
THP-1 cells in exponential growth phase were incubated in the presence or absence of carrageenan for 24 h before being stained with propidium iodide. Cell DNA content was then analyzed by flow cytometry. A: Histograms of cells treated with medium only (control), 10 kDa dCGN (C10), or 40 kDa dCGN (C40). B: Percentage of cells in each phase of the cell cycle when treated with medium only (control), different concentrations of 10 kDa dCGN (C10), of 40 kDa dCGN (C40), or of native CGN (Native).


Input:
Degraded CGN stimulated ICAM-1 expression in monocytes.
Peripheral blood monocytes (PBM) or THP-1 cells were incubated in the presence or absence of carrageenan for 24 h before being stained for various cell surface antigens. Antigen expression was then analyzed by flow cytometry. A: Histograms of ICAM-1 expression in cells treated with medium only (control), 10 kDa dCGN (C10), or 40 kDa dCGN (C40). B: Fluorescence intensity for expression of the antigens HLA-ABC, HLA-DR, CD14, ICAM-1, and CD58 in cells treated with medium only (control), 10 kDa dCGN (C10), or 40 kDa dCGN (C40). Data are mean +/- SEM.

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Input:
Degraded CGN induced monocytes aggregation in vitro.
Monocytes were incubated in the absence (A) or the presence of 1 mg/ml 10 kDa dCGN (B), 1 mg/ml 40 kDa dCGN (C), or 1 mg/ml 40 kDa dCGN plus 2.5 microg/ml anti-ICAM-1 antibody (D). Cells were observed by phase contrast inverse microscopy at 150X magnification. Inserts in A and B show a close up of cells at 300X magnification. E: Number of monocyte aggregates in 24 wells plate of PBM cell cultured with nothing (control), with 10 kDa degraded CGN (C10), or with 40 kDa degraded CGN (C40). Some cultures had also 2.5 microg/ml anti-ICAM-1 antibody. Data are mean +/- SEM.


Input:
Degraded CGN stimulated ICAM-1 and TNF-alpha gene expression in monocytes.
Representative samples of RT-PCR analysis showing over expression of ICAM-1 and TNF-alpha after stimulation of monocytes with 1 g/l of degraded CGN. beta-actin expression was used as normalization gene.

expression

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Input:
Degraded CGN activated the NF-kB pathway in monocytes.
A: THP-1 cells were transfected with a NF-kappaB reporter plasmid driving expression of luciferase. Cells were then treated with various concentrations of 10 kDa (triangles), or 40 kDa dCGN (squares). B: THP-1 cells treated with 1 mg/ml of 10 kDa dCGN (C10), or with 1 mg/ml of 40 kDa dCGN (C40) were lysed after various periods of time. Proteins in cell extracts were resolved by SDS-PAGE and then Western blotted for IkappaBalpha or alpha-tubulin as loading control. C: Degraded carrageenans (dCGN) induced activation of NF-kappaB. THP-1 cells were treated with nothing (control), or with 1 mg/ml of 10 kDa dCGN (C10), or with 1 mg/ml of 40 kDa dCGN (C40) for 30 minutes at 37degreesC. Nuclei were isolated and lysed. Proteins in nuclear extracts were resolved by SDS-PAGE and then Western blotted for NF-kappaB p50 subunit (p50) or NF-kappaB p65 subunit (p65). Lower panels show Western blots of nuclear ERK revealing equivalent amount of protein in each sample. Data are representative of three separate experiments. D: Degraded carrageenan (dCGN) induced activation of NF-kappaB. Nuclei isolated from THP-1 cells were fluorescence-stained for NF-kappaB p50 subunit or NF-kappaB p65 subunit before (filled area) or after cells were treated with 1 mg/ml of 10 kDa dCGN (C10), or with 1 mg/ml of 40 kDa dCGN (C40) for 30 minutes at 37degreesC. Dashed line corresponds to nuclei stained only with secondary fluorescence antibody. Fluorescence intensity was analyzed by flow cytometry as described.

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Input:
Transcription factors RUNX1 and RUNX3 in the induction and suppressive function of Foxp3+inducible regulatory T cells
Forkhead box P3 (FOXP3)+CD4+CD25+ inducible regulatory T (iT reg) cells play an important role in immune tolerance and homeostasis. In this study, we show that the transforming growth factor-beta (TGF-beta) induces the expression of the Runt-related transcription factors RUNX1 and RUNX3 in CD4+ T cells. This induction seems to be a prerequisite for the binding of RUNX1 and RUNX3 to three putative RUNX binding sites in the FOXP3 promoter. Inactivation of the gene encoding RUNX cofactor core-binding factor-beta (CBFbeta) in mice and small interfering RNA (siRNA)-mediated suppression of RUNX1 and RUNX3 in human T cells resulted in reduced expression of Foxp3. The in vivo conversion of naive CD4+ T cells into Foxp3+ iT reg cells was significantly decreased in adoptively transferred CbfbF/F CD4-cre naive T cells into Rag2-/- mice. Both RUNX1 and RUNX3 siRNA silenced human T reg cells and CbfbF/F CD4-cre mouse T reg cells showed diminished suppressive function in vitro. Circulating human CD4+ CD25high CD127- T reg cells significantly expressed higher levels of RUNX3, FOXP3, and TGF-beta mRNA compared with CD4+CD25- cells. Furthermore, FOXP3 and RUNX3 were colocalized in human tonsil T reg cells. These data demonstrate Runx transcription factors as a molecular link in TGF-beta-induced Foxp3 expression in iT reg cell differentiation and function.

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Input: Regulatory T (T reg) cells expressing the transcription factor forkhead box P3 (FOXP3, human; Foxp3, mouse) play an essential role in controlling immune responses to autoantigens, allergens, tumor antigens, transplantation antigens, and infectious agents (Hori et al., 2003;Akdis, 2006). Foxp3 is a member of the forkhead/winged-helix family of transcriptional regulators, and its expression in T reg cells is essential for their development and function (Fontenot et al., 2003;Williams and Rudensky, 2007). A spontaneous mutation of the X-linkedFoxp3gene inscurfymice causes an autoimmune-like disease, whereas the mutation in humans leads to immunodysregulation, polyendocrinopathy, enteropathy, and X-linked syndrome that is also a severe multiorgan autoimmune disease with hyper-IgE (Ziegler, 2006).Although the essential role of Foxp3 in central and peripheral tolerance has been extensively studied, its regulation, cooperation with other transcription factors, and how it functions in inducible T reg (iT reg) cells to suppress various target genes is mostly not yet understood. It is known that Foxp3 cooperates with the nuclear factor of activated T cells (NFAT) or nuclear factor-kappa B (NF-kappaB) to regulate the transcription of different target genes (Schubert et al., 2001;Bettelli et al., 2005;Wu et al., 2006). The Th2 cytokine IL-4 inhibits FOXP3 expression during T cell priming. GATA3 binds to theFOXP3promoter and can repress the FOXP3 trans-activation process directly in Th2 cells (Mantel et al., 2007). It was further demonstrated that both Th1 and Th2 transcription factors T-bet and GATA3 oppose peripheral induction of Foxp3+T reg cells in mice through STAT1-, STAT4-, and STAT6-dependent pathways (Wei et al., 2007). Although natural T reg (nT reg) cells that differentiate in the thymus are characterized by their stable Foxp3 expression, the generation of iT reg cells specific for allergens, alloantigens, and autoantigens in the periphery has been associated with a transient Foxp3+phenotype (Fontenot et al., 2003;Hori et al., 2003). The crucial role of TGF-beta in their generation has been demonstrated.The RUNX gene family (Runt-related transcription factor, acute myeloid leukemia [AML], core-binding factor-alpha [CBFalpha], and polyoma enhancer-binding protein-2alpha [PEBP2alpha]) contains three members, RUNX1 (AML1/CBFA2/PEBP2alphaB), RUNX2 (AML3/CBFA1/ PEBP2alphaA), and RUNX3 (AML2/CBFA3/PEBP2alphaC). They are essential transcriptional regulators of different developmental pathways. RUNX2 is mostly important for bone development and osteoblast differentiation (Komori et al., 1997). RUNX1 plays an important role in hematopoiesis during development, and RUNX3 has important functions in thymogenesis and neurogenesis (Wang et al., 1996;Inoue et al., 2002;Levanon et al., 2002). RUNX1 and RUNX3 also work together in the establishment of lineage specification of T lymphocytes (Taniuchi et al., 2002;Egawa et al., 2007).RUNX1is a frequent target for chromosomal translocations associated with leukemias (Look, 1997), andRUNX3methylation and silencing is observed in various human epithelial cancers (Blyth et al., 2005).RUNX family members share the Runt domain, which is responsible for DNA binding (Ito, 1999). The Runt domain-containing protein constitutes the alpha-chain partner of the heterodimeric CBF complex. RUNX proteins heterodimerize with the non-DNA-binding partner, CBFbeta, which increases the affinity for DNA binding and stabilizes the complex by preventing ubiquitin-dependent degradation (Wang et al., 1993). The CBF complexes regulate the expression of cellular genes through binding to promoters or enhancer elements. The effects of the RUNX-CBFbeta complex regulation are clearly cell lineage and stage specific. They include the crucial choices between cell-cycle exit and continued proliferation, as well as between cell differentiation and self-renewal (Blyth et al., 2005).Because of the involvement of RUNX mutations in different autoimmune diseases and the known interaction with TGF-beta, we investigated the impact of RUNX1 and RUNX3 on the expression of FOXP3 and subsequently on the development and function of iT reg cells. This study demonstrates that RUNX1 and RUNX3 induced by TGF-beta are involved in the development and suppressive function of Foxp3+iT reg cells.

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Input:
The role of RUNX1 and RUNX3 transcription factors in TGF-beta-mediated iT reg cell generation
To investigate the role of RUNX transcription factors in the development of iT reg cells, we cultured naive CD4+ T cells, isolated from human PBMCs, in conditions that enable the development of iT reg cells. Stimulation with anti-CD2/3/28 mAbs or TGF-beta alone resulted in a minimal up-regulation of RUNX1 and RUNX3 mRNA (Fig. 1 A). In contrast, the combination of both TGF-beta and anti-CD2/3/28 mAbs induced RUNX1 and RUNX3 mRNAs, as well as FOXP3 mRNA, within 48 h in naive CD4+ T cells. This result suggested further experiments to investigate whether the up-regulation of RUNX1 and RUNX3 might be a feature of iT reg cells during their development or even a prerequisite for their induction.
To test this hypothesis, RUNX1 and RUNX3 expression was knocked down in human naive CD4+ T cells by transfection of small interfering RNAs (siRNAs; Fig. 1 B). Deficiency of RUNX1 or RUNX3 resulted in markedly reduced TGF-beta-mediated induction of FOXP3 mRNA in naive CD4+ T cells compared with control cells transfected with scrambled siRNA. The level of FOXP3 mRNA was further reduced when both RUNX1 and RUNX3 were knocked down in naive CD4+ T cells during their differentiation to iT reg cells (Fig. 1 B).
The influence of RUNX1 and RUNX3 on the development of other T cell subsets and their specific transcription factor expression was further investigated. Naive CD4+ T cells were cultured under Th1, Th2, T reg cell, and Th17 differentiation conditions and the mRNA expression of the predominant transcription factor for each cell type was subsequently analyzed. We observed no change in GATA3 expression in Th2 cells, T-bet expression in Th1 cells, or RORC2 mRNA expression in Th17 cells in which RUNX1 and RUNX3 were knocked down compared with control cells. On the contrary, FOXP3 mRNA was significantly decreased in RUNX1- and RUNX3-deficient T reg cells compared with control cells (Fig. 1 C).
The effect of RUNX silencing on the expression level of intracellular FOXP3 during naive CD4+ T cell differentiation to iT reg was evaluated by flow cytometry. FOXP3 was only slightly reduced after RUNX1 silencing. Transfection of siRNA for RUNX3 had a stronger effect. The most striking FOXP3 reduction was observed when RUNX1 and RUNX3 were silenced together (Fig. 1 D). Similar results were obtained in total CD4+ T cells (Fig. S1 and Fig. S2). The increased impact of combined RUNX1 and RUNX3 knockdown implies that RUNX1 and RUNX3 might have redundant functions in the induction of FOXP3. In addition, the levels of IL-4, IL-5, IL-10, IL-13, and IFN-gamma in control siRNA-transfected or RUNX1 and RUNX3 siRNA-transfected CD4+ T cells that were cultured with or without anti-CD2/3/28 mAb and TGF-beta did not show any significant difference (Fig. S3).
To determine whether RUNX1 and RUNX3 are also expressed in human T reg cells in vivo, we isolated peripheral blood CD4+ CD127- CD25high T reg cells and compared them with CD4+ CD127+ CD25- T cells. Circulating T reg cells expressed significantly higher levels of RUNX3 mRNA compared with CD4+CD25- cells. As expected, IL-10, TGF-beta, and FOXP3 mRNAs are also expressed in circulating T reg cells (Fig. 2 A). There was no difference in RUNX1 mRNA expression between these two cell subsets. We also performed an analysis of human tonsils, which contain high numbers of FOXP3+ T reg cells (Verhagen et al., 2006). Staining of tonsil sections for FOXP3 and RUNX3 demonstrated in vivo coexpression of these two molecules in a subset of T reg cells, whereas there was low RUNX1 expression in all tonsil cells (Fig. 2 B).

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Input:
RUNX1 and RUNX3 bind to the FOXP3 promoter
Transcription element search system analysis of the human FOXP3 promoter predicted 3 putative RUNX binding sites at 333, 287, and 53 bp upstream of the transcription start site (TSS). All three binding sites are conserved between human, mouse, and rat (Fig. S4). To verify the putative binding sites in the FOXP3 promoter, we transiently transfected HEK293T cells with RUNX1 and RUNX3. After the pull-down with oligonucleotides containing the wild-type binding sequences, but not mutant sequences, RUNX binding to the FOXP3 promoter oligonucleotides was detected by Western blot (Fig. 3 A). To confirm these results and test the ability of single binding site sequences to bind either RUNX1 or RUNX3, we used the promoter enzyme immunoassay. Cell lysates were obtained from HEK293T cells that had been transiently transfected with RUNX1 or RUNX3 expression vectors. The biotinylated FOXP3 promoter oligonucleotides were linked to a streptavidin-coated microtiter plate, and bound RUNX1 or RUNX3 was detected by using anti-RUNX antibodies and a peroxidase-labeled secondary antibody. We showed binding of RUNX1 and RUNX3 to the mixture of all three oligonucleotides containing the binding sites, whereas there was no binding detectable when a combination of the mutated oligonucleotides was used in the assay (Fig. 3 B). Although there was a similar and high degree of binding to the -333 and -287 sites, a lower degree of binding was detected when the oligonucleotide containing the -53 site in the Foxp3 promoter was used. This effect was observed both for binding to RUNX1 and RUNX3 (Fig. 3 B). The binding to the two single binding sites at -333 and -287 was comparable to the mixture of all three oligonucleotides. Chromatin immunoprecipitation (ChIP) assay results confirmed the binding of RUNX1 and RUNX3 complexes containing CBFbeta to FOXP3 promoter during the differentiation of naive T cells toward T reg cells. Here, naive CD4+ T cells were cultured with IL-2, anti-CD2/3/28 mAb, and TGF-beta as a Foxp3-inducing stimulation. Amplification of PCR products from the FOXP3 promoter region with the predicted RUNX binding sites showed that RUNX1, RUNX3, and CBFbeta were immunoprecipitated together with the FOXP3 promoter (Fig. 3 C). Negative control primer targeting open reading frame-free intergenic DNA, IGX1A did not show any significant change in site occupancy.

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Input:
Regulation of FOXP3 promoter activity and FOXP3 protein expression by RUNX1 and RUNX3
To investigate the effect of RUNX1 and RUNX3 binding to the RUNX binding sites in the FOXP3 promoter, we transfected human peripheral blood CD4+ T cells with a FOXP3 promoter luciferase reporter vector and RUNX1 or RUNX3 expression vectors. An increase in luciferase activity was observed only when the FOXP3 promoter (-511 to +176) luciferase construct was cotransfected with RUNX1 or RUNX3 expression vectors (Fig. 4 A). The increase in promoter activity was greater upon cotransfection of RUNX3 compared with RUNX1. Luciferase expression was abrogated when the Runx binding sites in the FOXP3 promoter (-511 to +176) luciferase construct were mutated (Fig. 4 A). In these experiments, the overexpression of RUNX1 and RUNX3 eliminated the need of TGF-beta for FOXP3 promoter activation and PMA/ionomycin stimulation was sufficient.
To examine the role of each of the three RUNX binding sites for the FOXP3 promoter activity, we mutated each individually or in combination. No reduction in luciferase activity was observed when the -53 site was mutated and only a slight reduction when either the -287 or -333 site was mutated (Fig. 4 B). However, mutating the -53 site in combination with one of the other two sites led to a significant decrease in luciferase activity, with the greatest reduction observed when all three binding sites were mutated (Fig. 4 B), suggesting that the identified binding sites have redundant functions and RUNX binding to more than one site is necessary for the full activation of the FOXP3 promoter. Supporting these findings, the overexpression of RUNX1 in human primary CD4+ T cells resulted in significantly elevated levels of FOXP3 protein measured by flow cytometry after 48 h. This was achieved without any requirement for anti-CD3, anti-CD28 stimulation, or the presence of TGF-beta. Although there was a trend, the transfection of CD4+ T cells with RUNX3 did not lead to statistically significant increase in FOXP3 (Fig. S5).

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Input:
Role of CBFbeta in the induction of Foxp3
CBFbeta, a common cofactor of all RUNX proteins, stabilizes and increases the binding of the runt domain to target DNA sequences. To target all Runx proteins that might be involved in the induction of Foxp3, we used mice in which loxP-flanked Cbfb alleles were inactivated in T cells through expression of a CD4-cre transgene. Retinoic acid and TGF-beta synergize in the induction of Foxp3 in naive T cells (Kang et al., 2007). To investigate whether Runx-mediated induction of Foxp3 is dependent on the expression of CBFbeta, naive CD4+ CD8- T cells from CbfbF/F CD4-cre and control CbfbF/+ CD4-cre mice were stimulated with anti-CD3/28 mAbs, retinoic acid, and increasing concentrations of TGF-beta. After 3 d in culture, the cells were restimulated with PMA and ionomycin and analyzed for intracellular Foxp3 and IFN-gamma expression. TGF-beta induced Foxp3 in CbfbF/+ CD4-cre cells in a dose dependent manner, and this was significantly reduced in CbfbF/F CD4-cre cells. Retinoic acid enhanced Foxp3 expression even in 20 pg/ml of TGF-beta and more than 95% of the CD4+ T cells from CbfbF/+ CD4-cre mice became Foxp3+ in 100 and 500 pg/ml TGF-beta doses. The induction of Foxp3 was again significantly lower in CbfbF/F CD4-cre CD4+ T cells even in the presence of retinoic acid, demonstrating that deficiency in Runx binding to DNA affects the TGF-beta induction of Foxp3 in T reg cells (Fig. 5 A). There was no difference in the induction of Foxp3 when endogenous IL-4 and IFN-gamma were neutralized (Fig. S6).
The importance of RUNX transcription factors for the in vivo conversion of naive CD4+ T cells into iT reg cells was examined. Control CbfbF/F or CbfbF/F CD4-cre naive T cells, harboring a Foxp3-IRES-GFP allele were adoptively transferred into Rag2-/- mice. 6 wk later, CD4+ T cells in spleen, mesenteric lymph node, and lamina propria of the small intestine were analyzed for Foxp3-GFP expression (Fig. 5 B). There was a consistently lower percentage of CD4+ T cells that had developed Foxp3 expression in the mesenteric lymph node and lamina propria of mice transferred with CbfbF/F CD4-cre cells compared with control cells (Fig. 5, B and C). These data affirm the significance of RUNX proteins for the in vivo generation of CD4+ Foxp3+ T cells.

binding

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Role

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Input:
CBFbeta, RUNX1, and RUNX3 are important for the suppressive activity of CD25+ Foxp3+ T reg cells
Even though Cbfb-deficient CD4+ T cells had impaired induction of Foxp3 after stimulation with anti-CD3/28 mAbs and TGF-beta, sufficient numbers of Foxp3+ cells could be generated to permit analysis of their suppressive activity. Purified naive CD4+ T cells from CbfbF/F CD4-cre and control CbfbF/+ CD4-cre mice (Cd45.2) harboring a Foxp3-ires-GFP allele (Bettelli et al., 2006) were stimulated in vitro with anti-CD3/28 mAbs, IL-2, and TGF-beta. After 3 d, Foxp3+-GFP+ cells were sorted by flow cytometry and mixed with CFSE-labeled naive CD45.1+ CD4+ cells at ratios of 1:4, 1:2, and 1:1. The cells were then incubated with inactivated splenocytes and stimulated with anti-CD3 mAb for four more days. CD45.1+ cells were analyzed for CFSE dilution (Fig. 6 A). CbfbF/+ CD4-cre CD4+ T cells activated in the presence of TGF-beta showed a clear suppression of T cell proliferation that became even more apparent when an increased ratio of FOXP3+/CD25- cells was used (Fig. 6 B). The suppression was significantly reduced when cells from CbfbF/F CD4-cre mice were used, demonstrating that TGF-beta-induced Runx complexes are important for the suppressive activity of Foxp3+ T reg cells (Fig. 6 B). As a control, CbfbF/+ CD4-cre CD4+ T cells activated in absence of TGF-beta were mixed with CFSE-labeled naive CD45.1+ CD4+ cells. No suppression could be observed in all control groups without TGF-beta at all tested ratios (Fig. 6 C). The decreased suppression capacity of CbfbF/F CD4-cre iT reg cells was unlikely to be caused by decreased survival or proliferation. Foxp3+ and Foxp3- cells generated from both CbfbF/F CD4-cre and control cells all displayed similar proliferation rates and cell death as measured by CFSE dilution and annexin V staining, respectively (Fig. S7, A and B).
In addition, we tested the requirement of RUNX1 and RUNX3 for the development of the suppressive capacity in human iT reg cells. We isolated human naive CD4+ T cells and transfected them with a combination of RUNX1 and RUNX3 siRNA, or with a scrambled control siRNA. Cells were cultured under T reg conditions, and then mixed with CFSE-labeled autologous CD4+ T cells and stimulated with anti-CD3 mAb. Cells in which RUNX1 and RUNX3 were knocked down showed markedly lower suppressive activity compared with control iT reg cells at a T reg/CD4+ T responder cell ratio of 1:20, but not when the T reg/CD4+ T responder cell ratio was increased to 1:5 (Fig. 6 D). These results demonstrate the important role of RUNX1 and RUNX3 not only for the induction of FOXP3, but also for the suppressive capacity of iT reg cells both in humans and in mice. The data suggest both quantity and quality of T reg cells are hampered. Reduced intrinsic suppressive capacity of iT reg cells was demonstrated in mice, because Foxp3-GFP+ cells were FACS sorted and same numbers of iT reg cells are included in control experiments. In the suppression experiment with human cells, reduced suppressive activity was caused by reduced FOXP3 expression in T cells by siRNA inhibition of RUNX1 and RUNX3.

induction

expression

deficient

Cbfb

impaired

deficient

induction

siRNA

siRNA

knocked down

knocked down

reduced

expression

inhibition

inhibition

inhibition

induction

important

induction

important

induction


Input:
This study demonstrates that RUNX transcription factors 1 and 3 play an important role in the generation of FOXP3+ iT reg cells by TGF-beta. TGF-beta mediates RUNX induction and FOXP3 is efficiently up-regulated by RUNX1 and RUNX3 in human CD4+ T cells. There are three putative RUNX binding sites in the proximal FOXP3 promoter. One binding site was predicted as a binding site for RUNX2. Promoter enzyme immunoassay results showed that binding of RUNX1 and RUNX3 also occurred at this site (as well as at the other two), which were initially identified as RUNX1 binding sites. This finding is not surprising because RUNX proteins bind to promoter or enhancer elements of their target genes via the runt domain, which is conserved between members of the RUNX family. The RUNX protein that actually induces the expression of FOXP3 might therefore be dependent on the availability of the specific RUNX family member at certain stages of T cell development. RUNX proteins are able to increase or inhibit transcriptional activity of their target genes depending on the cell type and the target gene (Otto et al., 2003). Mutation of only one of the three binding sites had only a little effect on the promoter activity; however, when two binding sites were mutated, the FOXP3 promoter activity dropped to a greater extent. The most striking effect was observed when all three binding sites were mutated. We therefore assume that these binding sites have partially redundant functions, but binding to at least two sites seems to be necessary for full promoter activation.
TGF-beta promotes or inhibits the proliferation, differentiation, and survival of a wide array of different cells. It is also produced in activated T cells and it inhibits T cell proliferation (Kehrl et al., 1986; Siegel and Massague, 2003). It was shown that TGF-beta is mandatory for the maintenance of peripheral T reg cells and their expression of Foxp3 (Marie et al., 2005; Rubtsov and Rudensky, 2007). RUNX transcription factors are targets of the TGF-beta superfamily and they are involved in the TGF-beta pathway. They interact directly with regulatory SMADs (Miyazawa et al., 2002; Ito and Miyazono, 2003). TGF-beta can activate RUNX genes at the transcriptional level, and at the posttranscriptional level through activation or stabilization of RUNX proteins (Jin et al., 2004). It was shown that RUNX2 regulates the expression of TGF-beta type I receptor (Ji et al., 2001), suggesting that other mechanisms for their function could be involved. The fusion proteins RUNX1-EVI1 and RUNX1-ETO block TGF-beta inhibition of leukemic cell growth. RUNX3 plays an important role in TGF-beta-mediated growth control in epithelial cells, as loss of RUNX3 leads to decreased sensitivity to TGF-beta and hyperproliferation of the gastric mucosa (Blyth et al., 2005). The present study demonstrates that RUNX3 expression is more dominant in circulating human T reg cells and tonsil T reg cells compared with RUNX1. This could be dependent on the stage of the cells and organ from which they were isolated.
We observed that single siRNA interference of either RUNX1 or RUNX3 alone shows a slight decrease in Foxp3+ T reg cell induction, which could be caused by redundancy of these proteins. For this reason, we decided to use CbfbetaF/F CD4-cre mice. Foxp3 induction by TGF-beta is reduced in CD4+ T cells of CbfbF/F CD4-cre mice compared with CbfbF/+ CD4-cre mice. Retinoic acid is secreted by a subset of dendritic cells in the gut-associated lymphoid tissue. It inhibits the IL-6-driven induction of Th17 cells and facilitates the differentiation of naive T cells to Foxp3+ T reg cells (Mucida et al., 2007). We observed an increased number of Foxp3+ cells by retinoic acid and TGF-beta compared with TGF-beta treatment alone in CbfbF/+ CD4-cre mice and CbfbF/F CD4-cre mice. In addition, we showed a defective in vivo generation of T reg cells from Cbfb-deficient CD4+ T cells in Rag2-/- mice. These data in mice confirm the human data that RUNX proteins play an important role for TGF-beta-dependent FOXP3 induction, as well as in the suppressive capacity of iT reg cells. As an additional support for this concept, the overexpression of RUNX1 induced increased FOXP3 protein expression without any requirement of TGF-beta and anti-CD3 and anti-CD28 stimulation in human primary CD4+ cells. In both human and mouse systems, reduced Foxp3 expression was associated with reduced T reg cell suppressive activity.
In a recent study, the role of Runx-CBFbeta was investigated in nT reg cell development in the thymus (Rudra et al., 2009). It was reported that Foxp3 expression in nT reg cells is unstable in the absence of Runx-CBFbeta complexes. Cbfb-deficient nT reg cells progressively lose Foxp3 upon division, and there is no evidence of increased death of Cbfb-deficient nT reg cells in that study. The experiments in Cbfb-deficient CD4-cre T cells in mice and the knockdown experiments in humans in this study suggest that the induction of Foxp3 expression is a major contributing factor in the in vivo conversion experiment. Here, we observed that there is a twofold increased Foxp3+ iT reg cell generation in vivo. This is in the same range with previously published studies targeting different mechanisms in Foxp3 induction (Maynard et al., 2007; Sun et al., 2007). Whether the diminished capacity of Cbfb-deficient CD4-cre T cells in the generation of Foxp3 may be caused by peripheral expansion of Cbfb-deficient non-T reg cells or survival problems faced by Cbfb-deficient iT reg cells after Foxp3 induction remains to be elucidated.
The involvement of RUNX proteins in autoimmune diseases has been previously suggested (Alarcon-Riquelme, 2003). A mutation in the RUNX1 binding site in the promoter of programmed cell death 1 gene (PDCD-1) has been implicated in systemic lupus erythematosus pathogenesis (Prokunina et al., 2002). Polymorphisms that alter RUNX1 binding to other genes have also been described in rheumatoid arthritis linkage at 5q31 in Japanese patients (Tokuhiro et al., 2003) and in a psoriasis linkage at 17q25 (Prokunina et al., 2002; Helms et al., 2003). RUNX3-deficient mice spontaneously develop inflammatory bowel disease and hyperplastic gastritis-like lesions (Brenner et al., 2004). These disease symptoms resemble those occurring after depletion of Foxp3-expressing T reg cells (Sakaguchi, 2004). Derepression of Th2 cytokines might also account for some of the observed disease symptoms, as it was shown that T-bet first induces Runx3 in Th1 cells and then partners with Runx3 to direct lineage-specific gene activation. Runx3/Cbfbeta are both required for the activation of the Ifng gene and silencing of the Il4 gene in Th1 cells (Djuretic et al., 2007; Naoe et al., 2007). Runx proteins also play an essential role during T lymphocyte differentiation in the thymus (Taniuchi et al., 2002). Runx1 regulates the transitions of developing thymocytes from the CD4- CD8- double-negative stage to the CD4+ CD8+ double-positive stage and from the DP stage to the mature single-positive stage (Egawa et al., 2007). Runx1 and Runx3 deficiencies caused marked reductions in mature thymocytes and T cells of the CD4+ helper and CD8+ cytotoxic T cell lineages. In addition, inactivation of both Runx1 and Runx3 at the double-positive stages resulted in a severe blockage in the development of CD8+ mature thymocytes. These results indicate that Runx proteins have important roles at multiple stages of T cell development and in the homeostasis of mature T cells, and suggest that they may play a role in nT reg cell development, which remains to be elucidated. Furthermore, it was shown that Runx1 activates IL-2 and IFN-gamma gene expression in conventional CD4+ T cells by binding to their respective promoter. RUNX1 interacts physically with Foxp3 protein, and it was demonstrated that this interaction might be responsible for the suppression of IL-2 and IFN-gamma production and up-regulation of T reg cell-associated molecules (Ono et al., 2007).
It has been shown that Foxp3 also influences Th17 differentiation. Specifically, Foxp3 physically interacts with RORgammat, and this interaction inhibits RORgammat function (Zhou et al., 2008). This relationship of RORgammat and Foxp3 and probably yet unknown mechanisms might be the basis of the observation that the differentiation of Th17 cells and T reg cells is often reciprocal (Bettelli et al., 2006). Recently, data suggests that Runx1 may also be involved in regulating Il17 transcription, functioning in complex with RORgammat to activate transcription (Zhang et al., 2008).
The Runx3-deficient mice develop spontaneous Th2-dominated autoimmune colitis and asthma (Brenner et al., 2004; Fainaru et al., 2005). Cbfbf/f Cd4 mice also show a spontaneous Th2 dominated disease, with increased serum IgA, IgG1, and IgE titers and lymphocyte and eosinophil infiltration of the lung (Naoe et al., 2007). All these phenotypes were previously attributed to a loss of Th2 silencing whereas our findings additionally suggest that loss of T reg function plays a role. We have shown a link between Foxp3 induction in iT reg cells and RUNX1 and RUNX3. RUNX proteins play a central role in pathways regulating cell growth and differentiation, and their interaction with the TGF-beta pathway is of particular interest.
Foxp3 protein interacts not only with RUNX proteins but also with several other transcriptional partners, such as NFAT and possibly NF-kappaB; with histone acetyl transferases, such as TIP60; and histone deacetyl transferase (HDAC) complexes, such as HDAC7 and HDAC9 (Wu et al., 2006; Sakaguchi et al., 2008). NFAT forms a complex with AP-1 and NF-kappaB and regulates the expression of IL-2, IL-4, IFN-gamma, and CTLA4 in conventional T cells, which leads to the activation and differentiation to effector T cells (Dolganov et al., 1996; Hu et al., 2007). The NFAT-AP-1 complex also binds to the Foxp3 promoter after TCR triggering and regulates its gene expression positively (Mantel et al., 2006). It was shown that NFAT and Smad3 cooperate to induce Foxp3 expression through its enhancer (Tone et al., 2008), but no TGF-beta response element was identified in the Foxp3 gene or in the surrounding regions. The initial induction of RUNX1 and RUNX3 and the subsequent binding of these transcription factors to the Foxp3 promoter that we showed here might explain the relatively late induction of Foxp3 mRNA that peaks 24-48 h after stimulation. The interaction of Foxp3 and NFAT is dependent on their cooperative binding to DNA (Wu et al., 2006). RUNX1 alone, or together with its interacting partners p300 and CREB-binding protein, may cooperate with the NFAT transcription complex to activate the IL-2 promoter (Sakaguchi et al., 2008). Similar to this interaction, NFAT may also cooperate with RUNX1 or RUNX3 to activate Foxp3, but further studies are necessary to elaborate on this concept.
In conclusion, our findings elucidate the role of RUNX proteins in iT reg cell development and function. The induction of the transcription factors RUNX1 and RUNX3 by TGF-beta and the subsequent up-regulation of Foxp3 play a role in iT reg cell generation and its suppressive capacity.

binding

binding

bind

RUNX

runt domain

binding

binding sites

interact

RUNX

binding

partners

T-bet

binding

interacts

interacts

RORgammat

interaction

RUNX

TGF-beta

interacts

RUNX

interacts

forms a complex

binds

promoter

binding

promoter

binding

promoter

interaction

binding

interacting

binding

CREB

induction

RUNX

expression

produced

TGF-beta

expression

expression

TGF-beta type I receptor

expression

expression

induction

overexpression

expression

expression

expression

expression

generation

expressing

expression

expression

IFN-gamma

production

production

IFN-gamma

induction

expression

expression

expression

IFN-gamma

expression

expression

expression

loss

interference

interference

decrease

interference

induction

reduced

induction

deficient

Cbfb

reduced

expression

deficient

Cbfb

lose

deficient

deficient

Cbfb

deficient

Cbfb

diminished

generation

deficient

Cbfb

deficient

Cbfb

deficient

Cbfb

deficient

depletion

expressing

silencing

deficiencies

deficiencies

inactivation

inactivation

suppression

production

inhibits

interacts

RORgammat

deficient

mediates

TGF-beta

induction

up-regulated

up-regulated

induces

RUNX

expression

necessary

binding

activation

activation

promoter

mandatory

TGF-beta

expression

activate

TGF-beta

transcriptional

activation

TGF-beta

RUNX

induction

induction

TGF-beta

increased

TGF-beta

increased

TGF-beta

important role

RUNX

induction

dependent

TGF-beta

induction

induced

increased

increased

expression

requirement

induced

TGF-beta

induction

expression

increased

induction

caused

diminished

induction

induces

T-bet

required

activation

required

silencing

required

Cbfbeta

activation

required

Cbfbeta

silencing

activation

Ifng

activates

expression

binding

responsible

interacts

suppression

regulates

binds

expression

induce

expression

induction

induction

activate

promoter

activate

promoter

activate

induction

TGF-beta

induction

TGF-beta

up-regulation

dependent

induces

RUNX

targets

RUNX

involved

RUNX

regulates

expression

involvement

RUNX

alter

binding

regulating

transcription

regulates

forms a complex

expression

dependent

interaction

binding

transcriptional

RUNX

transcription

induction


Input:
Mice.
CbfbF/F CD4-cre and Foxp3GFP mice have previously been described (Bettelli et al., 2006; Naoe et al., 2007). Cd45.1 and Rag2-/- mice were purchased from Jackson ImmunoResearch Laboratories and Taconic, respectively. For the in vivo Foxp3 conversion assay, naive CD4+ T cells from Cbfb CD4-cre or control mice (harboring a Foxp3-IRES-GFP allele) were adoptively transferred into Rag-deficient mice. 5 x 106 cells were used per transfer. 6 wk later, TCRbeta+CD4+ gated cells from the spleen, mesenteric lymph node (MLN), and lamina propria of the small intestine were analyzed for Foxp3-GFP expression. All analyses and experiments were performed on animals at 6-8 wk of age. Animals were housed under specific pathogen-free conditions at the animal facility of the Skirball Institute, and experiments were performed in accordance with approved protocols for the New York University Institutional Animal Care and Usage Committee.

expression

deficient

Rag


Input:
Isolation of PBMCs, CD4+ T cells, and culture conditions.
Human PBMCs were isolated by Ficoll (Biochrom) density gradient centrifugation and CD4+ T cells were then isolated using the Dynal CD4+ Isolation kit (Invitrogen) according to the manufacturer's instructions. The purity of CD4+ T cells was initially tested by flow cytometry and was >=95%. Cells were stimulated with the following combination of mAbs to T cell surface molecules (Meiler et al., 2008): anti-CD2 (clone 4B2 and 6G4; 0.5 microg/ml), anti-CD3 (clone OKT3; 0.5 microg/ml), and anti-CD28 mAb (clone B7G5; 0.5 microg/ml; all from Sanquin) and cultured in serum-free AIM-V medium (Life Technologies) with the addition of 1 nmol/liter IL-2 (Roche). TGF-beta (R&D Systems) was used at 5 ng/ml, if not stated otherwise. A combination of PMA (25 ng/ml) and ionomycin (1 mg/ml; Sigma-Aldrich) was used.
Human CD4+ CD127- CD25high and CD4+ CD127+ CD25neg cells were purified by flow cytometry using anti-CD127, anti-CD25-PC5 (Beckman Coulter), and anti-CD4-FITC antibodies (Dako).
Mouse naive (CD62Lhi44lo25-) CD4+ T cells were purified by flow cytometry and activated in vitro with 5 microg/ml plate-bound anti-CD3 and 1 microg/ml soluble anti-CD28 antibodies (eBioscience) in RPMI supplemented with 10% FCS, 5 mM beta-mercaptoethanol, and antibiotics. Neutralizing anti-IFN-gamma and anti-IL-4 mAbs (BD) were used at 1 microg/ml concentrations when indicated.


Input:
In vitro T cell differentiation.
CD4+ CD45RA+ magnetically sorted (CD45RO depletion with AutoMACS; Miltenyi Biotec) cells were stimulated with immobilized plate-bound anti-CD3 (1 microg/ml; OKT3; IgG1) and anti-CD28 (2 microg/ml). For Th1 differentiation conditions, cells were stimulated with the following: 40 ng/ml IL-2, 5 microg/ml anti-IL-4, and 25 ng/ml IL-12 (R&D Systems). For Th2 conditions, cells were stimulated with the following: 40 ng/ml IL-2, 25 ng/ml IL-4, and 5 microg/ml anti-IL-12 (R&D Systems). For T reg cell conditions, cells were stimulated with the following: 40 ng/ml IL-2, 5 ng/ml TGF-beta, 5 microg/ml anti-IL-12, 5 microg/ml anti-IL-4. For Th17 conditions, cells were stimulated with the following: 40 ng/ml IL-2, 20 ng/ml IL-6, 5 ng/ml TGF-beta, 20 ng/ml IL-23 (Alexis Biochemicals Corp.), 10 ng/ml IL-1beta, 5 microg/ml anti-IL-4, and 5 microg/ml anti-IL-12 were used. Proliferating cells were expanded in medium containing IL-2. The cytokine profile of these cells demonstrated that IFN-gamma is the predominant cytokine in Th1 cells, IL-4 and IL-13 in Th2 cells, and IL-17 in Th17 cells (Akdis et al., 2000; Burgler et al., 2009).

depletion


Input:
Immunohistochemistry.
Human tonsils were obtained from tonsillectomy samples of hypertrophic and obstructive tonsils without a current infection. Ethical permission was obtained from Cantonal Ethics Commission, and informed consent was obtained from patients. Paraformaldehyde-fixed tonsil cryosections were stained with unconjugated rabbit IgG polyclonal antibody to human RUNX1 (Santa Cruz Biotechnology, Inc.) or unconjugated mouse IgG1 mAb to human RUNX3 (Abcam). After a washing step, the sections were stained with the corresponding secondary antibodies. RUNX1-binding antibodies were detected by using Alexa Fluor 633-conjugated goat anti-rabbit IgG and RUNX3-binding antibodies were detected by using Alexa Fluor 532-conjugated goat anti-mouse IgG1. Afterward, the sections were washed and in the case of RUNX3 staining a blocking step with an unconjugated mouse IgG1 mAb was used. Finally, the sections were stained with Alexa Fluor 488-conjugated mouse IgG1 mAb to human FOXP3 (eBioscience) or the corresponding isotype control. Tissue sections were stained with DAPI for the demonstration of nuclei and mounted with Prolong antifade (Invitrogen). Images were acquired and analyzed using the confocal microscope DMI 4000B and the TCS SPE system (both from Leica).

binding

binding


Input:
In vitro suppression assays.
Mouse Foxp3+ CD4+ CD8- T cells were FACS purified based on GFP expressed from a Foxp3-IRES-GFP knock-in allele (Foxp3GFP). Naive (CD62Lhi44lo25-) CD4+ T cells (effectors) were FACS-purified from Cd45.1 mice, then loaded with 5 microM CFSE (Invitrogen). Total splenocytes from C57BL/6 mice inactivated with 50 microg/ml mitomycin C (Sigma-Aldrich) for 45 min were used as APCs. A total of 4 x 105 CD4+ cells (CFSE-loaded CD25- plus Foxp3-GFP+) was mixed with 105 APCs + 1 microg/ml anti-CD3 mAb per well of a 96 well round bottom plate. Proliferation of the effector cells was analyzed by CFSE dilution. Apoptosis of the cells was investigated by annexin V staining and flow cytometry. Positive and negative control gates were made according to T cells cultured only in the presence of IL-2 without anti-CD3/28 stimulation.
Human naive CD4+ T cells were isolated by negative selection by MACS from PBMCs and either transfected with a scrambled siRNA or with a combination of RUNX1 and RUNX3 siRNA. Cells were then cultured under iT reg cell differentiating conditions and mixed with 2 x 105 autologous irradiated PBMCs that were used as APCs and autologous CFSE-labeled CD4+ T cells. T reg cell to responder cell ratio was 1:20, 1:10, and 1:5. To check the proliferation of the CD4+ T cells without suppression, no T reg cells were added in a control group. Cells were stimulated with 2.5 microg/ml anti-CD3 mAb, cultured in a 96-well plate and the proliferation of the effector cells was determined by analyzing the CFSE dilution by flow cytometry after 5 d of culture. Gating on the CD4+CFSE+ T cells enabled the exclusion of APCs and T reg cells.

expressed

GFP

siRNA

siRNA


Input:
Cloning of the FOXP3 promoter, construction of mutant FOXP3 promoter, and RUNX expression plasmids.
The FOXP3 promoter was cloned into the pGL3 basic vector (Promega Biotech) to generate pGL3 FOXP3 -511/+176 (Mantel et al., 2006). Site-directed mutagenesis for the three putative RUNX binding sites in the FOXP3 promoter region was introduced using the QuickChange kit (Stratagene), according to the manufacturer's instructions and confirmed by sequencing the DNA. The following primers and their complementary strands were used: foxp runx-333, forward 5'-CACTTTTGTTTTAAAAACTGTCCTTTCTCATGAGCCCTATTATC-3'; foxp runx-333 reverse 5'-GATAATAGGGCTCATGAGAAAGGACAGTTTTTAAAACAAAAGTG-3'; foxp runx-287 forward 5'-CCTCTCACCTCTGTCCTGAGGGGAAGAAATC-3'; foxp runx-287 reverse 5'-GATTTCTTCCCCTCAGGACAGAGGTGAGAGG-3'; foxp runx-53 forward 5'-GCTTCCACACCGTACAGCGTCCTTTTTCTTCTCGGTATAAAAG-3'; foxp runx-53 reverse 5'-CTTTTATACCGAGAAGAAAAAGGACGCTGTACGGTGTGGAAGC-3'.
The human RUNX1 fragment from the Addgene plasmid 12504 (Biggs et al., 2006) pFlagCMV2-AML1B was sub-cloned in the pEGFPN1 vector (Clontech Laboratories). The RUNX3 vector pCMV human RUNX3, which was a gift from K. Ito (Institute of Molecular and Cell Biology, Proteos, Singapore) was subcloned into pEGFPN1 vector (Clontech Laboratories; Yamamura et al., 2006).

expression

RUNX


Input:
Transfections and reporter gene assays.
T cells were rested in serum-free AIM-V medium overnight. 3.5 microg of the FOXP3 promoter luciferase reporter vector or a combination together with the RUNX1, RUNX3 pEGFPN1 vector, and 0.5 microg phRL-TK were added to 3 x 106 CD4+ T cells resuspended in 100 microl of Nucleofector solution (Lonza) and electroporated using the program U-15. After a 24-h culture in serum-free conditions and stimuli as indicated in the figures, luciferase activity was measured by the dual luciferase assay system (Promega) according to the manufacturer's instructions. PMA/ionomycin was used to stimulate the cells, because the transfection was only transient and the luciferase assay required a strong and fast stimulation of the cells. To evaluate the effect of overexpression of RUNX1 or RUNX3 on FOXP3 protein levels, CD4+ T cells were preactivated with 2 microg/ml phytohemagglutinin (Sigma-Aldrich) in serum-free AIM-V medium in the presence of 1 nmol/liter IL-2 (Roche) for 12 h, and then transfected with the vector pEGFPN1 containing the RUNX1 or RUNX3 fragment using the Nucleofector system (Amaxa Biosystems) and the program T-23. FOXP3 expression was evaluated by flow cytometry after 48 h of culture in AIM-V medium containing 1 nmol/liter IL-2.

overexpression

overexpression

transfected

transfected

expression

effect

overexpression


Input:
RNA interference.
CD4+ or naive CD4+ T cells were resuspended in 100 microl of Nucleofector solution (Lonza) and electroporated with 2 microM siRNA using the Nucleofector technology program U-14 (Lonza). Five different Silencer or Silencer Select Pre-designed siRNAs for RUNX1 (Applied Biosystems) and three Silencer Pre-designed siRNAs for RUNX3 (Applied Biosystems) were tested, and the best was selected for all further experiments. The Silencer Negative Control #1 siRNA (Applied Biosystems) was used for normalization. Cells were then left unstimulated or were stimulated after 12 h with anti-CD2, anti-CD3, and anti-CD28. Cells were cultured in serum-free AIM-V medium with the addition of 1 nmol/l IL-2 (Roche). Cells were harvested for mRNA detection of the target genes after 24 h and for protein detection after 48 h.

siRNAs

siRNAs


Input:
RNA isolation and cDNA synthesis.
RNA was isolated using the RNeasy Mini kit (QIAGEN) according to the manufacturer's protocol. Reverse transcription of human samples was performed with reverse-transcription reagents (Fermentas) with random hexamers according to the manufacturer's protocol.


Input:
Real-time PCR.
PCR primers and probes were designed based on the sequences reported in GenBank with the Primer Express software version 1.2 (Applied Biosystems) as follows: FOXP3 forward primer, 5'-GAAACAGCACATTCCCAGAGTTC-3'; FOXP3 reverse primer, 5'-ATGGCCCAGCGGATGAG-3'; EF-1a forward primer, 5'-CTGAACCATCCAGGCCAAAT-3'; and EF-1a reverse primer, 5'-GCCGTGTGGCAATCCAAT-3', as previously described (Mantel et al., 2007). GATA3 forward primer, 5'-GCGGGCTCTATCACAAAATGA-3'; and GATA3 reverse primer 5'-GCTCTCCTGGCTGCAGACAGC-3' (Mantel et al., 2007). T-bet forward primer, 5'-GATGCGCCAGGAAGTTTCAT-3'; T-bet reverse primer, 5'-GCACAATCATCTGGGTCACATT-3'; RORC2 forward primer, 5'-CAGTCATGAGAACACAAATTGAAGTG-3'; and RORC2 reverse primer 5'-CAGGTGATAACCCCGTAGTGGAT-3'. The prepared cDNAs were amplified using SYBR green PCR master mix (Fermentas) according to the recommendations of the manufacturer in an ABI PRISM 7000 Sequence Detection System (Applied Biosystems).
RUNX1 and RUNX3 mRNA was detected by using TaqMan Gene Expression Assays from Applied Biosystems and used according to the manufacturer's instruction using TaqMan master mix using a 7000 real-time PCR system (Applied Biosystems). PCR amplification of the housekeeping gene encoding elongation factor (EF)-1alpha or by using the 18S rRNA Gene Expression Assay (Applied Biosystems) was performed to allow normalization between samples. Relative quantification and calculation of the range of confidence was performed using the comparative deltadeltaCT method (Applied Biosystems). The percentage of FOXP3 mRNA in siRNA-mediated RUNX knockdown cells was calculated in relation to cells, which were transfected with scrambled control siRNA. Arbitrary units show the 2-(deltact) values multiplied by 10,000 incorporating the ct values of the gene of interest and the housekeeping gene.

knockdown

RUNX

mediated

knockdown

mRNA

mRNA

mRNA


Input:
Flow cytometry.
For analysis of human FOXP3 expression on the single-cell level, cells were first stained with the monoclonal CD4 mAb (Beckman Coulter), and after fixation and permeabilization, they were incubated with anti-human Foxp3-Alexa Fluor 488 antibody (BioLegend) based on the manufacturer's recommendations and subjected to FACS (EPICS XL-MCL; Beckman Coulter). A mouse IgG1 antibody (BioLegend) was used as an isotype control. Data were analyzed with the CXP software (Beckman Coulter). Cells were cultured with IL-2, and then left unstimulated or stimulated with anti-CD2/-CD3/-CD28 mAb.
Flow cytometry analyses of the mouse cells were performed on an LSRII (BD) and cell sorting was performed on a FACSAria (BD). All antibodies for these experiments were purchased from eBioscience or BD.

expression


Input:
Western blotting.
For human RUNX1 and RUNX3 analysis on the protein level, 106 cells were lysed and loaded next to a protein-mass ladder (Invitrogen) on a NuPAGE 4-12% Bis-Tris gel (Invitrogen). The proteins were electroblotted onto a PVDF membrane (GE Healthcare). Unspecific binding was blocked with 3% milk in TBS Tween, and the membranes were subsequently incubated with a 1:1,000 dilution of rabbit anti-RUNX1 (ab11903; Abcam) or 1:200 dilution of rabbit anti-RUNX3 (H-50; Santa Cruz Biotechnology, Inc.) in blocking buffer containing 3% milk in TBS Tween overnight at 4degreesC. The blots were developed using an anti-rabbit IgG HRP-labeled mAb (Cell Signaling Technology) and visualized with a LAS-1000 gel documentation system (Fujifilm). To confirm sample loading and transfer membranes were incubated in stripping buffer and reblocked for 1 h and reprobed using anti-GAPDH (6C5; Ambion) and developed using an anti-mouse IgG HRP-labeled mAb (Cell Signaling Technology).

protein level

protein level


Input:
Pull-down assay.
HEK293T cells were transfected with RUNX1 or RUNX3 using the Lipofectamine 2000 reagent (Invitrogen) according to the manufacturer's instruction. Cells were lysed by sonication in HKMG buffer (10 mM Hepes, pH 7.9, 100 mM KCl, 5 mM MgCl2, 10% glycerol, 1 mM DTT, 0.5% Nonidet P-40) containing a protease inhibitor cocktail (Roche Diagnostics). The cell lysate was precleared using streptavidin-agarose beads (GE Healthcare), incubated with biotinylated double-stranded oligonucleotides containing the wild-type or mutated RUNX binding sites, and polydeoxyinosinicdeoxycytidylic acid (Sigma-Aldrich). A combination of all three oligonucleotides containing the mutated or the wild-type binding sites was used in the assay. DNA-bound proteins were collected with streptavidin-agarose beads, washed with HKMG buffer, and finally resuspended in NuPAGE loading buffer (Invitrogen Life Technologies), heated to 70degreesC for 10 min, and separated on a NuPAGE 4-12% Bis-Tris gel (Invitrogen Life Technologies). The proteins were electroblotted onto a PVDF membrane (GE Healthcare) and detected using RUNX1 or RUNX3 antibodies described in the previous section.

transfected

transfected


Input:
Promoter enzyme immuno assay.
As performed in the pull-down assay, HEK293T cells were transfected with RUNX1 or RUNX3 and subsequently lysed. Insoluble material was removed by centrifugation. 384-well plates, precoated with streptavidin (Thermo Fisher Scientific) were washed 3 times with washing buffer (PBS and 0.05% Tween 20). Biotinylated FOXP3 promoter/oligonucleotides probes containing the RUNX binding sites were added (1 pmol per well; 50 fmol/microl) and incubated for 1 h at room temperature. Either a combination of all three oligonucleotides containing the mutated or the wild-type binding sites was used or single oligonucleotides were used in the assay. After 3 washing steps with washing buffer, the nuclear extract was added (concentration > 0.2 microg/microl) and incubated overnight at 4degreesC. The lysates were incubated with 10 microg of poly-deoxyinosinic-deoxycytidylic acid (Sigma-Aldrich). The plate was washed with HKMG buffer and incubated with a 1:1000 dilution of rabbit anti-RUNX1 (ab11903, Abcam) or 1:200 dilution of rabbit anti-RUNX3 (H-50, Santa Cruz Biotechnology, Inc.) at 4degreesC for 2 h. After three washing steps with HKMG buffer, a secondary antibody (anti-rabbit IgG-HRP, 1:3,000 in HKMG buffer, Cell Signaling Technology) was added, and the plate was incubated for 1 h at 4degreesC. The wells were washed 4 times with HKMG buffer before adding the substrate reagent (R&D Systems). The colorimetric reaction was stopped by adding 2 M H2SO4. Absorbance at 450 nm was measured using a microplate reader (Berthold Technologies).

transfected

transfected


Input:
ChIP.
Human naive CD4+ T cells were cultured either with IL-2 only or with IL-2, anti-CD2/3/28, and TGF-beta for 72 h, and protein-DNA complexes were fixed by cross-linking with formaldehyde in a final concentration of 1.42% for 15 min. Formaldehyde was quenched with 125 mM glycine for 5 min, and cells were subsequently harvested. The ChIP assay was performed as described in the fast chromatin immunoprecipitation method (Nelson et al., 2006). Cells were lysed with immunoprecipitation buffer (150 mM NaCl, 50 mM Tris-HCl, pH 7.5, 5 mM EDTA, NP-40 [0.5% vol/vol]) containing phosphatase (Roche) and protease inhibitors cocktails (Roche), the nuclear pellet was washed, the chromatin was sheared by sonication and incubated with antibodies for RUNX1 (H-65 X; Santa Cruz Biotechnology, Inc.), RUNX3 (H-50 X; Santa Cruz Biotechnology, Inc.), CBFbeta (PEBP2beta; FL-182 X; Santa Cruz Biotechnology, Inc.), and as controls normal rabbit IgG (Santa Cruz Biotechnology, Inc.), anti-human RNA polymerase II antibody, and mouse control IgG (both from SA Biosciences). The cleared chromatin was incubated with protein A agarose beads and, after several washing steps, DNA was isolated with 10% (wt/vol) Chelex 100 resin. Samples were treated with proteinase K at 55degreesC for 30 min. The proteinase K was then inactivated by boiling the samples for 10 min. The purified DNA was used in a real-time PCR reaction. Specific primers for the FOXP3 promoter, spanning the region from -87 to -3, FOXP3 promoter forward primer 5'-AGAGGTCTGCGGCTTCCA-3', FOXP3 promoter reverse primer 5'-GGAAACTGTCACGTATCAAAAACAA-3', or control GAPDH primer (SA Biosciences) for the RNA polymerase II were used. A negative control PCR for each immunoprecipitation using IGX1A negative control primer targeting ORF-free intergenic DNA (SA Biosciences) was used. The fold enrichment in site occupancy was calculated incorporating IgG control values and input DNA values using the ChampionChIP qPCR data analysis file (SA Biosciences).


Input:
Quantification of cytokine levels.
IL-4, IL-5, IL-6, IL-10, IL-13, IL-17, and IFN-gamma secretion was assessed using fluorescent bead-based technology. The Bio-Plex-hu Cytokine Panel, 17-Plex Group 1 was used according to the manufacturer's instructions (Bio-Rad Laboratories). Fluorescent signals were read and analyzed using the Bio-Plex 200 System (Bio-Rad Laboratories).

secretion

secretion

secretion

secretion

secretion

secretion

secretion

IFN-gamma