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. Author manuscript; available in PMC: 2012 Jul 10.
Published in final edited form as: Arthritis Rheum. 2011 Jul;63(7):10.1002/art.30229. doi: 10.1002/art.30229

NFAT5 is a critical regulator of inflammatory arthritis

Hyung-Ju Yoon 1, Sungyong You 2, Seung-Ah Yoo 1, Nam-Hoon Kim 1, H Moo Kwon 3, Chong-Hyeon Yoon 1,4, Chul-Soo Cho 1,4, Daehee Hwang 2,5, Wan-Uk Kim 1,4
PMCID: PMC3084342  NIHMSID: NIHMS262340  PMID: 21717420

Abstract

Objective

To investigate the role of nuclear factor of activated T cells 5 (NFAT5), which is known as an osmoprotective transcription factor, in synovial hyperplasia and angiogenesis in rheumatoid arthritis (RA)

Methods

Expression of NFAT5 was examined in the synovial tissues and synoviocytes of RA patients using immunohistochemistry and Western blot analysis, respectively. The mRNAs of RA synoviocytes and human umbilical vein endothelial cells (HUVEC) transfected with dummy siRNA or NFAT5 siRNA were profiled using microarray technology. Assays to determine synoviocyte apoptosis and proliferation were performed in the presence of NFAT5 siRNA.VEGF165-induced angiogenesis was assessed by measuring the proliferation, tube formation, and wounding migration of HUVEC. Experimental arthritis was induced in mice by injection of anti-type II collagen antibody.

Results

NFAT5 was highly expressed in the rheumatoid synovium and its activity was increased by proinflammatory cytokines, such as IL-1β and TNF-α. The mRNA profiling of synoviocytes and HUVEC transfected with NFAT5-targeted siRNA revealed three major changes in cellular processes associated with the pathogenesis of RA: cell cycle and survival, angiogenesis, and cell migration. Consistent with these results, NFAT5 knock-down in RA synoviocytes and HUVEC inhibited their proliferation/survival and impeded angiogenic processes in HUVEC. Mice with NFAT5 haplo-insufficiency (NFAT5+/-) developed very limited degree of synovial proliferation in histological analysis, decreased angiogenesis, and exhibited a nearly complete suppression of experimentally induced arthritis.

Conclusion

NFAT5 regulates synovial proliferation and angiogenesis in chronic arthritis.

Key indexing terms: NFAT5, synoviocytes proliferation, angiogenesis, rheumatoid arthritis

Introduction

Rheumatoid arthritis (RA) is a chronic inflammatory disease that leads to progressive joint destruction. It is characterized by the proliferation of fibroblast-like synoviocytes (FLS) and angiogenesis, the pannus formation. Within the joint synovium, FLS actively participate in the inflammatory processes of RA (1). These cells become resistant to apoptosis, proliferate abnormally, and produce not only several matrix metalloproteinases, but also pro-inflammatory cytokines, such as IL-1 and IL-6, and angiogenic factors such as vascular endothelial growth factor (VEGF) (2,3). In addition, angiogenesis is highly active in RA, particularly at the early-onset of the disease (4). Newly formed vessels can maintain a chronic inflammatory state by transporting inflammatory cells to the sites of synovitis, as well as supplying nutrients and oxygen to the pannus (5).

Nuclear factor of activated T cells 5 (NFAT5) was originally identified as a tonicity-regulated transcription factor involved in the cellular protection from hypertonic stress (6). However, recent reports suggest that NFAT5 might have additional roles in different tissues, including those not normally exposed to hypertonicity under either physiologic or pathologic conditions. For example, NFAT5 plays a role in lymphocyte proliferation and survival (7,8), and it mediates myoblast migration during skeletal muscle myogenesis via a cysteine rich protein 61 (CYR61) dependent pathway (9). In breast cancer cells, NFAT5 is induced by α6β4 clustering, which results in enhanced cell migration (10). Despite the progress in understanding of NFAT5 functional roles, little is known about its role in the pathogenesis of inflammatory arthritis.

We report herein that NFAT5 is involved in the pathogenesis of inflammatory arthritis. NFAT5 was highly expressed in RA synoviums, and that it was strongly induced by inflammatory stimuli. To elucidate role of NFAT5 in RA pathogenesis, we performed mRNA profiling of RA FLS and endothelial cells treated with NFAT5 siRNA. The results showed that NFAT5 involves three key cellular processes associated with RA pathogenesis: 1) cell cycle and survival, 2) angiogenesis, and 3) cell migration. Based on these data, we confirmed the proliferative, angiogenic, and migratory function of NFAT5 in cultured RA FLS and endothelial cells. Moreover, heterozygous deficiency of the NFAT5 gene resulted in a very limited degree of inflammatory cell infiltration, angiogenesis and synovial hyperplasia, and prevented the progression of experimental arthritis in mice. Collectively, these data demonstrate that NFAT5 is a critical regulator of synovial proliferation and angiogenesis, and thus it could be a potential target for the treatment of RA.

Materials and methods

Immunohistochemical staining for NFAT5

Immuno-staining for NFAT5 was performed in paraffin-embedded synoviums of RA and osteoarthritis (OA) patients using antibodies (Abs) to NFAT5 (6), as described previously (11).

Isolation and culture of FLS and HUVEC

The FLS from passages 3 to 6 were isolated from the synovial tissues of RA or OA patients according to a previously-described protocol (12), and incubated in DMEM supplemented with fetal bovine serum (FBS; Gibco, Carlsbad, CA). Human umbilical vein endothelial cells (HUVEC) also were isolated from normal umbilical cord veins and maintained in M 199 medium containing 20% FCS, as described previously (11,12).

Western blot analysis

The FLS and HUVEC were placed in a lysis buffer and the insoluble material was removed by centrifugation (12,000g) at 4°C for 20 minutes. Final protein concentrations were determined using the Bradford protein assay (BioRad, Hercules, CA). Electrophoresis was carried out by sodium dodecyl sulfate-10% polyacrylamide gel electrophoresis followed by a transfer to a nitrocellulose membrane. The membrane was incubated with Abs to NFAT5 (6), CCND1 (Santa Cruz Biotechnology, Santa Cruz, CA), CCNE1 (Santa Cruz Biotechnology), and CYR61 (Santa Cruz Biotechnology). The membrane was visualized using an enhanced chemiluminescent technique (ECL). The experiments were repeated three to four times using separate samples.

Immunocytochemical and Western blot analyses of nuclear translocation of NFAT5 in cultured synoviocytes

For immunostaining, the cells on glass chamber slides (1 × 103; Nunc GmbH & Co. KG, Wiesbaden, Germany) were fixed with chilled fresh methanol and permeabilized with PBS containing 0.25% Triton X-100. The slides then were treated with a blocking buffer (1% bovine serum albumin in PBS) and incubated overnight with anti-NFAT5 Ab at 4°C. After rinsing with PBS, the slides were incubated again with Alexa 488-conjugated anti-rabbit IgG (Invitrogen-Molecular Probes, Eugene, OR) for 30 minutes and were examined using a confocal laser scanning microscope.

Nuclear translocation of NFAT5 was also determined by Western blot analysis after the fractionation of nucleus and cytosol. To isolate cytoplasmic and nuclear fractions, cells were suspended in cold buffer A (10 mM HEPES; pH 7.9, 1.5 mM MgCl2, 10 mM KCl, 1 mM phenylmethylsulphonyl fluoride, 5 μg/ml leupeptin, and 10 μg/ml aprotinin) and allowed to swell on ice for 15 minutes. The cells were lysed by adding of Nonidet P-40 (final 0.6%) and centrifuged at 14000 g for 1 minute. After taking out of the supernatant which was used as a cytoplasmic fraction, we resuspended the residual nuclei in cold buffer B (20 mM HEPES; pH 7.9, 1.5 mM MgCl2, 0.42 M NaCl, 0.2 mM ethylenediaminetetraacetic acid, 25% glycerol, 1 mM phenylmethylsulphonyl fluoride, 5 μg/ml leupeptin, and 10 μg/ml aprotinin), and vigorously agitated them. After centrifuged at 14000 g for 20 minutes, the supernatant was taken and used as a nuclear fraction.

Transfection of NFAT5 siRNA

NFAT5 siRNA and scrambled control siRNA were purchased from Santa Cruz Biotechnology. The FLS and HUVEC were seeded in culture plates and grown to 60 to 80% confluence for 24 hours. The cells were transfected with 20 nM of siRNA using lipofectamine 2000 reagent in Opti-MEM medium according to the manufacturer's instructions (Invitrogen, Carlsbad, CA). After the transfection, cultures were incubated at 37 °C for 4 hours and then placed in fresh culture medium. Forty eight hours after transfection, the cells were harvested, and the NFAT5 mRNA and protein expression levels were determined by reverse transcription-polymerase chain reaction (RT-PCR) and Western blot analysis, respectively. For the detection of NFAT5 mRNA, PCR was performed using NFAT5 specific primers; sense 5′-AAGAGTGAAGATGTTACTCCA-ATGGAAG-3′, antisense 5′-AAAGTCTGTGCTTGTTCTTGTAGTGG-3′. PCR amplification was conducted for 25 cycles of denaturation at 94°C for 30 seconds, annealing at 57°C for 60 seconds, and elongation at 72°C for 30 seconds. β-actin mRNA expression was used as an internal control.

Identification of differentially expressed genes

The mRNAs of RA FLS (n=3) and HUVEC (n=4) transfected with dummy siRNA or NFAT5 siRNA were profiled using Illumina HumanRef-8 (for RA FLS) and HumanHT-12 (for HUVEC). Raw data (GSE22956) can be viewed at the Gene Expression Omnibus (online at http://www.ncbi.nlm.nih.gov/geo/). Untransfected cells were used as a control. The intensities from the arrays were normalized using quantile normalization (13). Using the normalized intensities, differentially expressed genes (DEGs) between control siRNA and NFAT5 siRNA-transfected samples were determined using the following integrated statistical hypothesis testing: 1) two independent tests, T-test and log2 median ratio test, were performed; 2) P values from each test were computed using an empirical distribution of the null hypothesis that the means of the genes are not different, which was obtained from random permutations of the samples; 3) the individual P values were combined to compute the false discovery rate (FDR) using Stouffer's method (14); and 4) the DEGs were selected as the genes with the FDR less than 0.01 and fold change larger than the cutoff of 1.5. Finally, functional enrichment analysis of DEGs was performed using DAVID software to identify cellular processes overrepresented by the DEGs governed by NFAT5.

Real-time PCR

Total RNA was reverse-transcribed to cDNA using a High Capacity cDNA RT kit (Applied Biosystems, Foster City, CA) according to the manufacturer's instruction. The 80–150 base pair long primers were designed to specifically amplify target mRNAs. The abundance of mRNA was then detected by Thermal cycler dice real-time systems (Takara, Shiga, Japan) with SYBR Green (Takara). The quantity of mRNA was calculated by using the 2-ΔΔCt method (15), and then normalized by the ACTB abundance. The quantitation of each mRNA was repeated three times. The gene-specific primers were listed in supplementary Table 1.

Cell viability: MTT assay

The synoviocyte viability was measured using MTT assay or Cell Counting Kit-8 (CCK-8; Dojindo Molecular Technology, Gaithersburg, MD) assay, as we described previously (16,17).

Detection of FLS apoptosis

For the detection of apoptotic cells, TUNEL assay was performed using the ApoTag® peroxidase in situ apoptosis detection kit (Roche Applied Science, Indianapolis, IN) at 72 hours after NFAT5 siRNA transfection.

Assay for cell proliferation

After NFAT5 siRNA transfection, the FLS and HUVEC were subjected to a proliferation assay using [3H] thymidine incorporation, as described previously (12,16,17). The proliferative responses were expressed as counter per minute (c.p.m.).

Wounding migration and capillary tube formation assays

The migration of HUVEC after in vitro wounding was measured as described previously (12,17). In brief, HUVEC plated to confluence on 60-mm culture dishes were wounded with pipette tips, and then treated with VEGF165 (20 ng/ml) in M199 medium supplemented with 1% FBS and 1 mM of thymidine. After 12 hours of incubation, migration was quantified by counting the cells that had moved beyond a reference line. For the capillary tube formation assay, HUVEC were seeded on a layer of previously polymerized Matrigel (BD Biosciences, San Jose, CA) with VEGF165 (20 ng/ml). After 12 hours of incubation, the degree of tube formation was quantified by measuring the length of tubes in 5 randomly chosen low-power fields (× 50) from each well using image-Pro Plus v4.5 (Media Cybernetics, San Diego, CA).

Assay for chemotaxis of endothelial cells

The chemotactic migration of HUVEC was assayed using a Transwell chamber with 6.5 mm diameter polycarbonate filters (8 μm pore size) (12,17). In brief, VEGF (50 ng/ml), which was prepared in DMEM that contained 1% FBS, was placed in the lower wells. In the upper wells, the HUVEC were suspended in DMEM containing 1% FBS at a final concentration of 5 × 104 cells/ml. The chamber was incubated at 37°C for 12 hours. Non-migrating cells on the filter's upper surface were removed by wiping them with a cotton swab. The migrated cells were fixed and stained utilizing a Diff-Quik kit (Microptic, Barcelona, Spain). Chemotaxis was quantified by counting the cells that migrated to the lower side of the filter with an optical microscopy at a magnification of × 200. Eight random fields were counted for each assay.

Induction of anti-collagen antibody-induced arthritis

Eight week-old heterozygous NFAT5+/- mice, backcrossed for eight generations onto a C57BL/6 background, and their wild type littermate (NFAT5+/+) were used for this study. The mixture of anti-type II collagen Ab (5 mg; Chondrex, Seattle, WA) was injected intravenously to the two groups of mice, and 3 days later, 50 μg of lipopolysaccharide (LPS) was administered intraperitoneally (11,16). The clinical severity of arthritis was graded as described previously (11,16). During the course of arthritis, forefoot and hind foot swelling also were measured every day using a microcaliper. The diameter of the arthritic ankle at specific time points divided by the diameter at day 0 was defined as the paw thickness index, which was presented as percentage. At day 10, the paws and ankles were harvested from each mouse. The degree of inflammation, synovial hyperplasia, and joint destruction were determined using a standard scoring protocol (11,16), where the severity was scored on a scale of 0 to 3; where 0 = absent, 1 = weak, 2 = moderate, 3 = severe. The maximum total score possible was 12.

Statistical Analysis

Data are expressed as the mean ± standard deviation (SD). Comparisons of the numerical data between groups were performed by the paired or unpaired Mann-Whitney U-test. P values less than 0.05 were considered statistically significant. A permutation test strategy was used to determine the significance of overlapping of DEGs between two datasets. A total of 100,000 random permuted samples were used to compute the empirical P value of the overlapping DEG.

Results

Increased expression of NFAT5 in rheumatoid synovium and synoviocytes

Using immunohistochemical staining, NFAT5 expression was examined in joint tissues obtained from patients with RA and those with osteoarthritis (OA) as non-inflammatory controls. As shown in Figure 1A, a high level of NFAT5 expression was observed in synovial tissue sections obtained from 3 representative patients with RA (a to e), which is consistent with a previous report (18). Positive staining was observed predominantly in RA synovium, particularly in the lining layer (arrowheads in b,d,e, and g), sublining leukocytes, and endothelial cells (arrows in f). As a negative control (c), isotype antibody did not show any immunoreactivity with the NFAT5 in the same RA synovium as ‘b’. NFAT5 also was expressed in the lining layer and endothelial cells of OA synoviums (n=3; f to h), but its expression level was relatively low compared to that of RA synoviums. FLS are a major component of the lining layer where NFAT5 was highly expressed. Since NFAT5 was known to be regulated by ambient tonicity (6), we stimulated FLS by elevating medium NaCl concentration by 100 mM. As expected, the hypertonic treatment increased NFAT5 expression over the course of several hours in both RA and OA FLS (Figure 1B). However, synovial fluid in RA joints was not hypertonic: 268 ± 16 mosmol/kg for RA patients (n=11) and 268 ± 14 mosmol/kg for OA patients (n=6). We reasoned that factors other than ambient osmolarity might contribute to a high level of NFAT5 in RA synoviums. To address this question, we investigated the effect of pro-inflammatory cytokines on the expression of NFAT5 in RA FLS. We identified first that IL-1β and TNF-α induced NFAT5 expression in FLS, and these increases were greater in RA FLS than in OA FLS (Figure 1C). IL-1β and TNF-α also increased NFAT trafficking to the nucleus from the cytoplasm of FLS, as determined by both immuno-fluorescence staining and Western blot analysis of nuclear translocation of NFAT5 (Figure 1D and 1E), indicating that these cytokines enhance NFAT5 activity by at least two points: increased expression and nuclear translocation. Moreover, basal levels of NFAT5 in the nucleus over the cytoplasm were significantly higher in RA FLS (n=6) than in OA FLS (n=6) (Figure 1F). Together, these findings partly explain why RA synoviums express NFAT5 at high levels.

Figure 1. NFAT5 is highly expressed in RA synovium and induced by pro-inflammatory cytokines.

Figure 1

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(A) Immunohistochemical staining of rheumatoid arthritis (RA; a-e) and osteoarthritis (OA; f-h) synovia using anti-NFAT5 antibody (Ab); c, isotype control. The rectangular area in (a) is magnified to (b). (B,C) Induction of NFAT5 in FLS by hypertonicity and pro-inflammatory cytokines. The FLS were treated with 100 mM of NaCl (B) or stimulated with IL-1β (10 ng/ml) or TNF-α (10 ng/ml) for 12 hours (C). NFAT5 expression was determined by Western blot analysis. (D) Immuno-cytochemical staining for NFAT5 in RA FLS stimulated with NaCl (100 mM), IL-1β (10 ng/ml) or TNF-α (10 ng/ml) for 3 hours. (E) Western blot analysis of nuclear translocation of NFAT5 in RA FLS stimulated with NaCl (100 mM), IL-1β (10 ng/ml) or TNF-α (10 ng/ml) for 12 hours. C: cytosol, N: nucleus. (F) Basal distribution of NFAT5 in the nucleus over the cytosol in the FLS of RA patients and OA patients, determined by Western blot analysis. Bars in the lower panel indicate the mean level. *, P < 0.05. The data from B to F are the representative of more than three independent experiments.

Transcriptomes regulated by NFAT5 in RA FLS and endothelial cells

To explore cellular processes regulated by NFAT5 associated with RA pathogenesis, we performed mRNA profiling of RA FLS and HUVEC as FLS and endothelial cells are key components of invasive pannus. RA FLS (n=3) and HUVEC (n=4) were transfected with NFAT5-targeted siRNA or non-specific scrambled (control) siRNA, cultured in the presence of 10% FBS for 48 hours, and their gene expression was profiled using microarray analysis. We first confirmed NFAT5 knockdown in RA FLS and HUVEC (Figure 2A and data not shown). DEGs between those cells transfected with NFAT5-targeted siRNA versus control siRNA were then investigated using an integrative statistical testing (see METHODS). We identified 682 (310 upregulated and 372 downregulated genes in NFAT5 siRNA treated samples) and 910 (457 upregulated and 453 downregulated genes) DEGs with a FDR < 0.01 and fold change > 1.5 in RA FLS and HUVEC datasets, respectively (Figure 2B and 2C). Among the total of 1373 DEGs, 219 DEGs were shared by RA FLS and HUVEC (P=0.00001) while 463 and 691 genes were specific to RA FLS and HUVEC, respectively (Figure 2B and 2C). Functional enrichment analysis of the shared and cell type-dependent DEGs was performed using DAVID software (19), which showed that these DEGs are involved mainly in three cellular processes closely associated with RA pathogenesis: 1) cell cycle and survival, 2) angiogenesis, and 3) cell migration (Figure 2D). For example, 41 of 219 DEGs are involved in the cell cycle (P=2.20×10-14), out of which 38 DEGs are downregulated and only 3 are upregulated, suggesting that NFAT5 promotes the proliferation of RA FLS and endothelial cells. Of note, 39 of the 463 DEGs dominant in RA FLS are involved in cell death, while 25 and 21 of 691 DEGs dominant in HUVEC are associated with cell migration (P=1.90×10-4) and angiogenesis (P=1.10×10-6), respectively (Figure 2D). Quantitative real time PCR assays of selected 19 DEGs, which associated with three pathways intrinsic for the development of rheumatoid pannus, confirmed the changes in their expression levels in independent RA FLS and HUVEC samples (Figure 2E and 2F).

Figure 2. Molecular signatures regulated by NFAT5 in FLS and HUVEC.

Figure 2

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(A) Decrease in NFAT5 expression level after the transfection of NFAT5 siRNA (lane 3). The mRNA and protein expression levels of NFAT5 in RA FLS were determined by RT-PCR (upper panel) and Western blot analysis (lower panel), respectively; lane 1: no transfection, lane 2: control siRNA transfection. (B) Venn diagram of DEGs depicts the overlap between RA FLS and HUVEC microarray data sets. The P value indicates the significance of 219 overlapping DEGs. (C) Heatmap displaying the expression patterns of 1373 DEGs in RA FLS or HUVEC 48 hours after NFAF5 siRNA transfection (lane 2); lane 1: control siRNA transfection. (D) Biological processes perturbed by NFAT5 siRNA in RA FLS and HUVEC. Functional enrichment analysis of common and cell type-dependent DEGs was performed using DAVID software. (E and F) Quantitative real time PCR assays for a subset of DEGs associated with three key cellular processes, including cell cycle and survival, angiogenesis, and cell migration in RA FLS (E) and HUVEC (F). Data represent the mean ± SD of the three replicates.

NFAT5 promotes survival and proliferation of synoviocytes

Based on the analysis of the DEGs described above, we next sought to establish if NFAT5 functionally modulates cell survival and proliferation of RA FLS since alterations in mRNA associated with survival and proliferation were most prominent in NFAT5-deficient FLS. As expected, NFAT5 knockdown decreased RA FLS viability significantly, which was determined by the Cell Counting Kit-8 (CCK-8) assay, at 72 hours after transfection (Figure 3A); Under phase-contrast microscopy (upper panel), the FLS transfected with NFAT5 siRNA for 72 hours became spherical, shrunken, and detached from the bottom of the culture plate (right), whereas the untreated cells retained a bipolar appearance (left). The number of apoptotic cells, determined by TUNEL staining, a sensitive tool for the detection of apoptosis, also was markedly increased in FLS transfected with NFAT5-targeted siRNA, but not in those with control siRNA or untransfected cells (Figure 3B), indicating a role of NFAT5 in preventing the apoptotic death of FLS. Moreover, NFAT5 siRNA also completely abrogated the TNF-α- and TGF-β-induced increases in [3H]-thymidine incorporation into RA FLS (Figure 3C), suggesting that NFAT5 promotes FLS proliferation. Reduced proliferation was similarly noted in mouse embryonic fibroblasts (MEF) of NFAT5-/- mice when compared to NFAT5+/+ MEF (data not shown), suggesting that the regulation of cell growth by NFAT5 is not limited to inflammatory RA FLS. Cyclin expression and cell cycle progression were reported to be regulated by NFAT5 in T cells exposed to hypertonic stress (20). Consistently, the levels of cyclin D1 (CCND1) and cyclin E1 (CCNE1) were increased in RA FLS in response to TNF-α and TGF-β (Figure 3D), and downregulation of NFAT5 by siRNA resulted in a marked decrease in TNF-α and TGF-β-stimulated expression of CCND1 and CCNE1. Taken together, NFAT5 is an important mediator for survival and proliferation of FLS.

Figure 3. Effect of NFAT5 on the survival and proliferation of FLS.

Figure 3

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(A) Effect of NFAT5 siRNA on survival of FLS. The degree of cell viability was determined by CCK-8 assay after the transfection of NFAT5 siRNA for indicated times (lower panel). Data are presented as a relative value to untransfected cells. The symbol of * represents P < 0.05 versus control siRNA-transfected cells (same in the following Figure 3A to 3C). (B) Increase in FLS apoptosis by NFAT5 siRNA. The extent of apoptosis was determined by the TUNEL staining of RA FLS at 72 hours after the NFAT5 siRNA transfection. Black arrows indicate TUNEL positive cells. (C) Decrease in FLS proliferation by NFAT5 siRNA. At 24 hours after NFAT5 siRNA transfection, RA FLS were treated with TGF-β (10 ng/ml) or TNF-α (10 ng/ml). The FLS proliferation rate was determined by [3H] thymidine incorporation assays at 48 hours after the treatment. †, P < 0.05 versus untransfected cells in the absence of TGF-β or TNF-α (D) Cyclin D1 and E1 expression determined by Western blot analysis in RA FLS at 24 hours after the stimulation of RA FLS with TNF-α (10 ng/ml) or TGF-β (10 ng/ml).

NFAT5 regulates the proliferation and migration of endothelial cells

In addition to abnormal proliferation of FLS, angiogenesis is a critical step for the initiation and progression of RA (4,5). Since the gene expression data from NFAT5-deficient HUVEC exhibited significant changes in cell survival and proliferation, similar to RA FLS, we further tested whether NFAT5 can modulate survival and proliferation of HUVEC as well. As shown in Figure 4A, NFAT5 siRNA decreased the viability of HUVEC at 48 hours after transfection, as determined by the MTT and CCK-8 assays (Figure 4A and data not shown). Furthermore, it decreased the proliferation of HUVEC in the presence or absence of VEGF165 (Figure 4B). The gene expression data also suggested that NFAT5 regulates cell migration predominantly in HUVEC (Figure 2D). Thus, we tested whether NFAT5 can regulate the endothelial tube formation, migration, and chemotaxis under the non-lethal time condition of 24 hours after transfection. As expected, the formation of tube-like structures induced by VEGF165 was markedly decreased by NFAT5 siRNA (Figure 4C). Additionally, NFAT5 knock-down suppressed the VEGF165-induced increase in the migration of HUVEC in response to wounding (Figure 4D). The VEGF165-stimulated chemotaxis of HUVEC in Boyden chambers also was completely impeded by NFAT5 siRNA (Figure 4E). In support of this observation, the protein expression level of CYR61, a key mediator of cell migration and angiogenesis (21), was reduced in NFAT5 siRNA-transfected HUVEC (Figure 4F), which is in accordance with our real-time PCR data performed in FLS and HUVEC (Figure 2E and 2F). Collectively, our data demonstrate that NFAT5 promotes proliferation, migration, and chemotaxis of endothelial cells in vitro.

Figure 4. NFAT5 knock down reduces angiogenic activity of HUVEC.

Figure 4

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(A) Effect of NFAT5 siRNA on survival of HUVEC as determined by CCK-8 assay. Data are normalized with respect to control siRNA-transfected cells. (B) Decrease in proliferation of HUVEC by NFAT5 siRNA. The HUVEC were transfected with NFAT5 siRNA in the presence or absence of VEGF165 (20 ng/ml). Cell proliferation rate was determined by [3H] thymidine incorporation assays at 48 hours after the transfection. The symbol of * represents P < 0.05 versus control siRNA-transfected cells (same in all the following figures). (C) Inhibition of tube formation of HUVEC by NFAT5 siRNA. The total length of the tube network was calculated using Image-Pro Plus software at 24 hours after the NFAT5 siRNA transfection when HUVEC are still viable. (D) NFAT5 siRNA suppresses the wounding migration of HUVEC induced by VEGF165. (E) Inhibition of chemotaxis of HUVEC in a Boyden chamber by NFAT5 siRNA, as determined at 24 hours after the transfection. (F) Reduction of CYR61 expression in HUVEC transfected with NFAT5 siRNA, compared to control siRNA-transfected HUVEC and untransfected cells, as determined by Western blot analysis.

Decrease in arthritis severity in NFAT5 deficient mice

We next attempted to define the role of NFAT5 in an in vivo animal model of arthritis. To this end, experimental arthritis was induced in wild type and NFAT5 deficient mice by administration of arthritogenic, anti-type II collagen Ab. In this passive model of arthritis, the innate immune system, including FLS and endothelial cells, plays a central role in the progression of joint disease independent of adaptive immunity (22). Since complete NFAT5 deficiency was lethal, we used a line of mice heterozygous (NFAT5+/-) for an inactive allele (7). The heterozygotes displayed haplo-insufficiency of NFAT5 mRNA and protein expression in many tissues we examined, including the joints (data not shown and ref. [7]). As shown in Figure 5A, the severity of arthritis was significantly lower in the NFAT5+/- mice when compared with wild type, NFAT5+/+ littermates. Paw swelling, assessed by the diameter of arthritic ankle, also was much lower in the NFAT+/- mice than in NFAT5+/+ mice (Figure 5B). In comparison with the NFAT5+/+ mice, the NFAT5+/- mice also developed a very limited degree of inflammatory cell infiltration, joint destruction, and synovial hyperplasia on histological analysis (Figure 5C and 5D). Moreover, NFAT5 haplo-insufficiency mitigated the extent of angiogenesis in the synovia of mice with antibody-induced arthritis, as determined by immunofluorescence staining for von Willebrand factor (Figure 5E), suggesting that NFAT5 is essential for pathologic angiogenesis in vivo. Together, these findings indicate that partial ablation of NFAT5 gene sufficiently blocks the progression of experimental arthritis, suppressing synovial proliferation and angiogenesis in the arthritic joints.

Figure 5. Arthritis induction in NFAT5+/- mice and NFAT5+/+ mice.

Figure 5

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(A) Severity of anti-type II collagen antibody-induced arthritis in NFAT5+/- (n=8) and wild type littermates (NFAT5+/+, n=8). The symbol of * represents P < 0.001 versus NFAT5+/+ mice (same in all the following figures). (B) Effect of NFAT5 deficiency on paw edema in mice with antibody-induced arthritis. Data are presented using the paw thickness index of forefoot (FF) and hind foot (HF). (C) Hematoxylin and eosin staining of joints from NFAT5+/- versus NFAT5+/+ mouse. The rectangular areas in the upper panel (× 40) are magnified to the lower panel (× 100). (D) Mean histological scores of inflammatory cell infiltration (IFLM), synovial proliferation (SP), and joint destruction (JD) in NFAT5+/- (n=8) versus NFAT5+/+ mice (n=8) as determined at day 10 after antibody administration. (E) Expression of von Willebrand factor (vWF) in the joints of mice. Ten days after the injection of anti-type II collagen antibody, ankle joints of NFAT5+/- (KO, n=8) and wild type NFAT5+/+ mice (WT, n=8) were subjected to immuno-fluorescence staining using anti-vWF Ab. A representative of vWF (+) cells (white arrows) in the joints of NFAT5+/- mice is shown. Mean fluorescence intensity (MFI) was compared between the two groups.

Discussion

Hypertonicity has been recognized as the most prominent signal for NFAT5 (23). NFAT5 stimulated by local hypertonicity via signaling pathways, including p38 mitogen-activated protein (MAP) kinase (24), is essential for the survival and proliferation of lymphocytes (7,8). Likewise, local hypertonicity in the subcutaneous tissue drives the NFAT5-mediated VEGF-C expression and lymphatic capillary hyperplasia leading to normalization of sodium balance and blood pressure (25). In renal epithelial cells, hypertonicity-activated NFAT5 promotes inflammation by stimulating nuclear factor-κB bound to the promoters of its target genes (26). In this study, we found that proinflammatory cytokines, including TNFα and IL-1β, stimulated both expression and nuclear localization of NFAT5 in FLS. This is the first to demonstrate that NFAT can be activated by factors other than hypertonicity, and that the proinflammatory stimuli represent a distinct modality of signal for NFAT5.

We also demonstrate a novel pro-inflammatory function for NFAT5 and show that its absence reduces arthritis severity in mice. Using gene expression profiling and in vitro assays, we further show that NFAT5 regulates several cellular processes closely related to RA pathogenesis, including 1) the survival and proliferation of synoviocytes and endothelial cells, and 2) the angiogenesis and migration of endothelial cells. These results collectively suggest that the NFAT5 regulation of such cellular processes in the major types of cells implicated in RA, including FLS and endothelial cells, is likely responsible for the suppression of experimentally-induced arthritis. It is remarkable that a 50% reduction in the NFAT5 activity results in almost complete suppression of arthritis in vivo.

Our work demonstrates that NFAT5 regulates both shared and dominant cellular processes in RA FLS and HUVEC. Gene expression data indicate that NFAT5 promotes proliferation/survival in both FLS and HUVEC, but NFAT5 inhibits apoptosis predominantly in FLS, while it promotes angiogenesis/cell migration predominantly in HUVEC (Figure 2D). Consistent with this, analysis of DEGs between NFAT5-/- and NFAT5+/+ MEF also reveals that NFAT5 controls cell proliferation and migration, as well as angiogenic process (data not shown). Thus, it would appear that the core molecular processes governed by NFAT5 are shared, while certain processes are particular to cell context. In this regard, it is important to characterize functions of NFAT5 in other cell types involved in RA pathogenesis, such as T cells, B cells, and mast cells. Additionally, it would be interesting to investigate shared versus distinct NFAT5-dependent processes in individual cell types at the molecular level.

To explore clinical relevance of our findings, we compared the NFAT5-dependent gene signatures in FLS and HUVEC to the previously published data in RA synovial tissues treated with an anti-TNF-α Ab, Adalimumab (27). Among the 1373 NFAT5-dependent DEGs identified in RA FLS or HUVEC (Figure 2B and 2C), 233 DEGs overlapped with 1565 DEGs (P < 0.05 and fold change >1.5) found in RA synovial tissues of poor versus good responders to Adalimumab (Figure 6A and 6B). To elucidate potential links between NFAT5 and responses to Adalimumab, we focused on 93 out of 233 shared DEGs that were downregulated by NFAT5 knockdown but upregulated in poor responders to Adalimumab (Figure 6B). Functional enrichment analysis revealed that the 39 genes among the 93 genes are involved in cell cycle and regulation of apoptosis (P < 0.05; Figure 6C). These results raise the possibility that dysregulated NFAT5 signatures in RA FLS and HUVEC, specifically those related to cell death and cell cycle, might mimic treatment responsiveness to anti-TNF-α antibody and indicate a subset of immunopathology unique to RA.

Figure 6. NFAT5 signature in FLS and HUVEC reflects the responsiveness to anti-TNF-α treatment.

Figure 6

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(A) Heatmap displaying the expression of 93 genes upregulated (red) in poor responders compared to good responders, but downregulated (green) in either RA FLS or HUVEC transfected with NFAT5 siRNA. (B) Venn diagram depicting the overlap between the genes perturbed by NFAT5 in this study and the DEGs in the RA synovial tissues of poor versus good responders to the anti-TNF-α antibody, Adalimumab, as described in a previous study (ref. 27). This overlap is highly significant (P < 0.00001). (C) Biological processes enriched by the 93 genes.

In summary, the results presented here demonstrate for the first time the functional role of NFAT5 in promoting synovial hyperplasia and angiogenesis, key pathological features of invasive pannus in arthritis. This study also offers the first evidence on the functional and molecular links of NFAT5 to cellular process contributing to arthritis development in major cell types composing the pannus. In this respect, NFAT5 might be an effective therapeutic target not only in inflammatory arthritis but also in some other inflammatory and angiogenesis-dependent disorders because NFAT5 is a hub of multiple pathways.

Supplementary Material

Supp Table S1

Acknowledgments

We thank all members of the Institute of Bone and Joint Diseases at the Catholic University of Korea.

This work was supported by grants from National Research Foundation of Korea (NRF) funded by the Ministry of Education, Science and Technology (R33-2008-000-10064-0), the Korea Healthcare technology R&D Project, Ministry for Health, Welfare and Family Affairs (No. A092258 and A080768), and in part by NIH grant (RO1-DK42479).

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Supplementary Materials

Supp Table S1
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