The transcription factor NFAT1 plays a pivotal role in the homeostasis of T lymphocytes. However, its functional importance in non-CD4+ T cells, especially in systemic immune disorders, is largely unknown. In this study, we report that NFAT1 regulates dendritic cell (DC) tolerance and suppresses systemic autoimmunity using the experimental autoimmune myasthenia gravis (EAMG) as a model. Myasthenia gravis and EAMG are T cell–dependent, Ab-mediated autoimmune disorders in which the acetylcholine receptor is the major autoantigen. NFAT1-knockout mice showed higher susceptibility to EAMG development with enhanced Th1/Th17 cell responses. NFAT1 deficiency led to a phenotypic alteration of DCs that show hyperactivation of NF-κB–mediated signaling pathways and enhanced binding of NF-κB (p50) to the promoters of IL-6 and IL-12. As a result, NFAT1-knockout DCs produced much higher levels of proinflammatory cytokines such as IL-1β, IL-6, IL-12, and TNF-α, which preferentially induce Th1/Th17 cell differentiation. Our data suggest that NFAT1 may limit the hyperactivation of the NF-κB–mediated proinflammatory response in DCs and suppress autoimmunity by serving as a key regulator of DC tolerance.

The NFAT family of transcription factors is comprised of five proteins (NFAT1 through NFAT5 [TonEBP]), which play crucial roles in the regulation of the immune responses of T cells, B cells, dendritic cells (DCs), and megakaryocytes through the regulation of Ca2+/calcineurin–NFAT–mediated signaling pathways (14). Given the essential role of NFAT proteins in immune regulation, NFATs have been considered key targets for treating diverse immune disorders. Calcineurin inhibitors, such as CsA and FK506, have generally been used as immunosuppressive drugs to prevent graft rejection and treat autoimmune diseases (5). However, although calcineurin inhibitors suppress T cell activation, they also induce diverse side effects mediated by NF-κB activation and the subsequent induction of proinflammatory cytokines in macrophages, kidney tubular cells, nonlymphoid fibroblastic cells, and mesangial cells (6). Moreover, the exact role of NFAT proteins in the development and pathogenesis of systemic autoimmunity and whether they have pro- or anti-inflammatory properties remain unclear.

Immune cells express NFAT1 (NFATc2), NFAT2 (NFATc1), and NFAT4 (NFATc3) (1). NFAT1 regulates T cell activation as well as the differentiation of Th cells, including Th1, Th2, Th17, follicular T helper (Tfh) cells and regulatory T (Treg) cells (7, 8). In contrast to the positive role of NFAT protein in T cell activation and differentiation, mice lacking NFAT protein(s) showed different phenotypes in response to diverse immune stimulation. NFAT1-deficient mice showed moderate lymphocyte hyperproliferation associated with splenomegaly and enhanced T cell and B cell responses with bias toward Th2 cell responses (9, 10). Mice lacking both NFAT1 and NFAT4 also developed a hyperproliferative phenotype (11). NFAT proteins play important roles in non-CD4 T cells. NFAT1 regulates CD8+ T cell activation as well as exhaustion programs, depending on the nature of its binding partners (12). Whereas NFAT1−/− DCs have been shown to facilitate the differentiation of naive CD4+ T cells to the Th1 phenotype (13), the functional importance of NFAT proteins in the pathogenesis of chronic inflammatory disorders is still unclear.

In this study, we investigated the role of NFAT1 in the development of systemic autoimmunity using an animal model of autoimmune myasthenia gravis (experimental autoimmune myasthenia gravis [EAMG]). Myasthenia gravis (MG) is a systemic autoimmune disease mainly caused by autoantibodies specific for the nicotinic acetylcholine receptor (AChR) in the neuromuscular junction (NMJ). The binding of autoantibodies to AChR leads to the failure of signal transmission at the NMJ. Rodents develop MG symptoms when injected with serum from MG patients or anti-AChR Abs from EAMG mice (14). Removal of anti-AChR Abs reduces the severity of MG symptoms (15, 16).

CD4+ T cells have crucial roles in MG development (17). MG patients have abundant T cells which mainly produce proinflammatory cytokines (18). Among the Th cell subsets, Th1 and Th17 cells play crucial roles in human MG and EAMG by producing proinflammatory cytokines, such as TNF-α, IL-1β, and IL-6 (1923). Although calcineurin inhibitors are used to treat MG, the exact role of NFAT1 in MG progression remains unclear. To address this issue, we compared the disease severity and pathogenic parameters of EAMG between wild-type (WT) and NFAT1-knockout (NFAT1KO) mice. NFAT1KO mice showed more severe EAMG symptoms accompanied by an upregulation of AChR-reactive lymphocyte proliferation as well as enhanced levels of Th1 and Th17 cells, proinflammatory cytokines (IL-1β, IL-6, IFN-γ, TNF-α, and IL-17), and anti-AChR–reactive IgG levels. DCs may play a key role in this process. NFAT1-deficient DCs produced high levels of IL-6 and IL-12 cytokines, which led to the enhancement of Th1/Th17 cells. NFAT1 may regulate DC tolerance by inhibiting the hyperactivation of NF-κB–mediated proinflammatory signaling pathways in systemic autoimmunity.

C57BL/6, TCR-transgenic OT-II, CD11c.DOG, and NFAT1-deficient mice were housed in specific pathogen-free barrier facilities and were used in accordance with protocols approved by the Pohang University of Science and Technology Institutional Animal Care and Use Committee (approval number: POSTECH-2014-0033). C57BL/6 mice and OT-II mice were purchased from SLC (Hamamatsu, Japan). NFAT1KO mice were provided by Dr. A. Rao, La Jolla Institute for Allergy and Immunology.

EAMG was induced in 8- to 10-wk-old female C57BL/6 WT and NFAT1KO mice by immunization with Torpedo AChRs (TAChRs) obtained from the electric organs of Torpedo californica via affinity chromatography, as described previously (24). Briefly, mice were immunized through multiple routes, including base of the tail, footpad, and shoulders, by injection of purified AChR (30 μg/mouse) emulsified in CFA (Difco). After the first immunization, the second and third immunizations were performed in 4-wk intervals by injecting AChR emulsified in IFA into the thighs and shoulders, near the first injection sites. Clinical symptoms of EAMG were monitored 2–3 times per week. The clinical scoring of EAMG was graded from zero to four based on observations made before and after exercise as follows: grade 0, mice with normal posture, muscle strength, and no symptoms of EAMG even after exercise (20–30 consecutive paw grips to a steel grid cage top); grade 1, normal at rest and slightly decreased activity and grip strength with hunched posture after exercise; grade 2, hunched posture at rest, weakness, and tremor; grade 3, severe weakness, dehydrated, paralyzed, and moribund; grade 4, dead. Clinical scores were assessed by double-blind evaluation for 13 wk following the first immunization. All experimental groups consisted of 10–13 mice. To measure the effect of the administration of neutralizing Abs, NFAT1KO mice were treated with anti–IL-6 (0.5 μg/mouse) or anti–IL-12 (1 μg/mouse) Abs every other day, starting from the first AChR immunization and continuing to the end of the experiments.

Muscles attached to the femur and tibia were extracted from nonimmunized, normal healthy mice and EAMG-induced mice in WT (WT/EAMG) or NFAT1KO (NFAT1KO/EAMG). They were then embedded in Tissue-Tek OCT compound and sectioned in the transverse direction into 10 μm using a Jung Frigout 2800E Kryostat (Leica Camera, AGVertrieb). For each experimental group, 10–15 muscle sections were analyzed. Muscle sections were fixed in cold acetone for 10 min and were washed in PBS three times for 5 min. Sections were blocked with 2% PBSA (BSA in PBS) for 1 h at room temperature, and then washed in PBS three times for 10 min. Sections were incubated for 90 min at room temperature with tetramethylrhodamine-conjugated α-bungarotoxin (Molecular Probes) or FITC-labeled goat anti-mouse C3c Ab (Nordic Immunological Laboratories, Eindhoven, the Netherlands), and then washed in PBS three times for 10 min. After the final wash, the samples were mounted with Dako mounting solution. The signal intensities of AChR (green) and C3 (red) fluorescence in the images were quantified by measuring the average pixel intensity with FluoView FV 1000 (Olympus). Relative fluorescence intensity of NTAT1KO/EAMG was calculated by comparing with that of WT/EAMG mice.

CD4+ T cells, CD19+ B cells, and CD11c+ DCs were purified from the draining lymph nodes (dLNs) and spleen by CD4-, CD19-, and CD11c-specific immunomagnetic beads according to the manufacturer’s instructions (Miltenyi Biotec). For coculture experiments, CD11c+ DCs were cocultured with CD4+ T cells in the presence of indicated amounts of AChR or OVA peptide, depending on the experimental system, in T cell medium containing RPMI 1640 medium (Invitrogen) supplemented with 10% FBS (Hyclone), 3 mM l-glutamine (Sigma-Aldrich), 10 mM HEPEs (Sigma-Aldrich), 100 U/ml penicillin (Sigma-Aldrich), 100 U/ml streptomycin (Sigma-Aldrich), and 0.05 mM 2-MΕ (Sigma-Aldrich). Proliferation was also assessed by measuring [3H]thymidine (0.5 μCi/well) incorporation during the last 18 h of a 72-h culture period. Results are expressed in δ cpm.

Three EAMG mice with representative mean clinical scores of the WT and NFAT1KO groups were sacrificed. Mixed lymphocytes from dLNs and splenocytes were cultured in the presence of AChR for 48 h and were harvested. Total RNA was extracted by using TRIzol reagent (Molecular Research Center), according to the manufacturer’s protocol. Then, it was reverse transcribed to prepare cDNA using M-MLV Reverse Transcriptase (Promega, Madison, WI). Prepared cDNA was subjected to quantitative real-time PCR (qRT-PCR) using Chromo-4 (Bio-Rad) with SYBR Premix Ex Taq (Takara). Specific primer pairs are described in Table I. The data were normalized to the expression level of hypoxanthine–guanine phosphoribosyltransferase. Results were described as relative expression levels for each gene in the NFAT1KO mice compared with that of WT mice. For the analysis of protein expression levels, culture supernatants were collected from mixed lymphocytes obtained from the dLNs, spleen, and purified CD4+ T cells stimulated in the presence of indicated amounts of AChR for 96 h. Protein expression was determined using ELISA kits specific for IL-6, IL-12, IFN-γ, TNF-α, and IL-17A according to the manufacturer’s instructions (eBioscience, San Diego, CA). For determination of AChR-reactive IgG isotypes, serums were obtained from EAMG mice 7 d after the second immunization by retro-orbital bleeding. Anti-mouse AChR Ab levels were measured using ELISA as follows: 96-well microplates were coated with human recombinant AChR Hα1-205 (100 μl; 20 μg/ml), which shows high homology in the amino acid sequences of rat AChR (24), and reacted with 100 μl of the tested serum samples. The dilution factors 1:10,000, 1:5,000, 1:1,000, and 1:5,000 were used for total IgG, IgG1, IgG2a, and IgG2b, respectively. HRP-conjugated mAbs to mouse IgG isotypes (1:10,000 for total IgG, IgG1, IgG2a, and IgG2b) were added for 1 h at room temperature and bounded Abs were detected using the activity of the peroxidase. Ab levels were evaluated by measuring the OD at 490 nm.

Cells were lysed in 1× RIPA buffer consisting of 50 mM Tris, pH 8, 150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, and were then sonicated five times on 30% amplitude for 3 s on ice. After centrifugation at 13,000 × g for 15 min, proteins in each supernatant were separated by SDS-PAGE using a 4–12% gel (Koma), transferred on a nitrocellulose membrane, and immunoblotted for indicated Abs. The following Abs were also used for immunoblotting: NFAT1 (SC-7296), FOXP-3 (SC-28705), T-bet (SC-21749), GATA3 (SC-268), p50 (SC-7178), Bcl-6 (SC-858), and NFAT2 (SC-7294) (Santa Cruz); Erk1/2 (9101), p-JNK (4686), JNK (9258), phospho-p38 (9215), p38 (9212), IκBα (9242), phospho-c-Jun (9261), and lamin B (9087) (Cell Signaling); GAPDH (in house); α-tubulin (LF-PA0146; AbFrontier); and RORγ (Ab78007; Abcam)

Chromatin immunoprecipitation (ChIP) assays were performed as previously described (25). In brief, bone marrow DCs (BMDCs) from WT and NFAT1-deficient mice were cross-linked with 1% formaldehyde after stimulation with or without Pam3 CSK (Pam3) for 1 h prior to harvest. Fixed prepared cells were lysed and sonicated to shear DNA. Sheared chromatin was immunoprecipitated with anti-p50 Ab (Ab7971; Abcam) or rabbit IgG Ab (Sigma-Aldrich). Ab–chromatin complexes were eluted and cross-linking was reversed by boiling with Chelex 100. After reversal of cross-links, the presence of enriched DNA was determined by real-time PCR using the following primers: IL-6 promoter forward, 5′-CCC ATC AAG ACA TGC TCA AG-3′; reverse, 5′-GCA CAA TGT GAC GTC GTT-3′; and IL-12 promoter forward, 5′-AGT ATC TCT GCC TCC TTC CTT-3′; reverse, 5′-GCA ACA CTG AAA ACT AGT GTC-3′ (26, 27). As a loading control, qRT-PCR was performed directly on input DNA purified from chromatin before immunoprecipitation. Data are presented as the amount of DNA recovered relative to input control.

NF-κB and IL-6 promoter reporter constructs were generated as previously described (28, 29). WT and NFAT1KO BMDCs were transfected with nucleofectin (Amaxa Nucleofector; Lonza) according to the manufacturer’s protocol. After 18 h, cells were treated with Pam3 for 12 h, and luciferase activity was assessed by the dual-luciferase assay system (Promega). Cotransfection of the HRE-luciferase vector as an internal control allowed normalization of transfection by Renilla luciferase activity.

Mouse BMDCs were generated as previously described (30, 31). Briefly, BMDCs are cultured from the bone marrow precursor cells isolated from the tibias and femurs of NFAT1KO and WT mice. Bone marrow precursor cells were plated on day 0 onto bacterial petri dishes with RPMI 1640 medium supplemented with the following: penicillin and streptomycin, 10% FBS, 2 mM l-glutamine, and 20 ng/ml recombinant murine GM-CSF (PeproTech). On day 3, an equal volume of the above media was added. On days 6–7, nonadherent cells were harvested (32). BMDCs were further sorted with anti-CD11c (purity >97%) magnetic beads by following manufacturer’s instructions (Miltenyi Biotec). The amount of contaminated bone marrow–derived macrophages (CD11c+MHCIIlowF4/80high) was <2% in the sorted CD11c+ cells. DC maturation and activation were determined by checking surface markers including CD40, CD80, CD86, and MHCII.

Before the transferring of BMDC, CD11c+ DCs were depleted by i.p. administration of diphtheria toxin (DT) (8 ng/g DT; Sigma-Aldrich, in PBS) to CD11c.DOG mice that express the human DT receptor (DTR) under control of the CD11c promoter (CD11c.DOG mice; Hochweller et al. [33]) (Supplemental Fig. 3I). BMDCs from WT and NFAT1KO mice were adoptively transferred through the footpads of DT-treated CD11c.DOG mice. Mice were also immunized with TAChR at the time of BMDC transfer. Protein levels of IFN-γ, TNF-α, IL-6, and IL-17A from mixed lymphocytes of adoptively transferred EAMG mice were measured by ELISA.

The expression levels of the costimulatory molecules CD40, CD80, CD86, and MHCII in mixed lymphocytes of dLNs and splenocytes from WT/EAMG and NFAT1KO/EAMG mice were analyzed in the presence or absence of AChR stimulation for 72 h. For the analysis of splenocyte subpopulations, CD4, CD8, B220, CD11c, CD11b, F4/80, CD25, CD44, and CD62L cell surface markers were used. The expression levels of the costimulatory molecules CD40, CD80, CD86, and MHCII in splenocytes and BMDCs were measured in the presence or absence of indicated TLR-agonist stimulation. The cells were harvested and washed twice with cold PBS. Fluorescently labeled Abs specific for cell subsets and costimulatory molecules were stained for 20 min at 4°C and then washed with cold PBS. To measure the different levels of intracellular cytokines and transcription factors, mixed lymphocytes or dLN CD4+ T cells from WT/EAMG and NFAT1KO/EAMG were stimulated with AChR. Expression levels of intracellular IFN-γ, TNF-α, IL-6, IL-17A, IL-17F, IL-21, T-bet, Rorγt, and Foxp3 were analyzed by flow cytometry. In brief, mixed lymphocytes from dLNs were cultured for 4 d in the presence of AChR. Mixed lymphocytes or in vitro–differentiated Th cells were restimulated with PMA and Ionomycin for 5 h with 1 mg/ml Brefeldin A and tracked by intracellular cytokine staining. Cells were harvested, washed with PBS, and fixed in fixation/permeabilization buffer (eBioscience) for 30 min. Cells were then washed and resuspended in 100 μl permeabilization buffer (eBioscience). For intracellular detection of IFN-γ, TNF-α, IL-6, IL-17A, IL-17F, IL-21, T-bet, Rorγt, and Foxp3; anti–IFN-γ–Alexa 488, PE (eBioscience), anti–TNF-α–PE, anti–IL-4–FITC, anti–IL-6–PE, anti–IL-17A–PE, anti–IL-17F, anti–IL-21–PE, anti–T-bet–PE, anti-Rorγt–PE, and anti-Foxp3–PE or isotype control Ab (eBioscience) was added and incubated for 30 min at 4°C. After incubation, cells were washed, resuspended in 1 ml PBS, and analyzed using flow cytometry.

RNA extracted from the BMDCs prepared from WT and NFAT1KO mice were stimulated with Pam3 (2–12 h) or left without stimulation. Total RNA from BMDCs from WT and NFAT1KO mice were extracted by using TRIzol reagent (Molecular Research Center), according to the manufacturer’s protocol. Illumina mRNA array experiments were done by Macrogen (Seoul, Korea).

For statistical analysis, all experiments were performed more than three times independently. Unless otherwise noted, statistical analyses were performed using the Student t test (two-tailed, two-sample unequal variance). A p <0.05 was considered significant.

To examine the exact role of NFAT1 in the development and progression of systemic autoimmunity, we first compared EAMG susceptibility between WT and NFAT1KO mice. Compared with WT mice, NFAT1KO mice showed higher disease susceptibility with more severe symptoms (Fig. 1A, Table II), higher complement deposits (>50% increase), and reduced AChR levels (>10% decrease) as determined by the fluorescence intensity (Fig. 1B). The sizes of dLNs and number of lymphocytes were comparable between the groups under non-EAMG conditions. However, their levels were significantly increased in NFAT1KO mice under EAMG conditions (NFAT1KO/EAMG) (Fig. 1C, 1D). NFAT1KO/EAMG mice also showed significantly higher levels of total anti-AChR IgG, IgG1, and IgG2a in serum (Fig. 1E, Supplemental Fig. 1A). Mixed lymphocytes from the dLNs (Fig. 1F), splenocytes (Fig. 1G), and CD4+ T cells (Fig. 1H) isolated from NFAT1KO/EAMG mice showed substantially higher proliferation rates after AChR stimulation. In addition, lymphocytes and splenocytes isolated from NFAT1KO/EAMG mice showed higher levels of costimulatory molecules (Supplemental Fig. 1B, 1C) regardless of AChR stimulation in vitro. Collectively, these results indicate that enhanced EAMG susceptibility in the NFAT1KO mice is associated with the hyperresponsiveness of AChR-reactive lymphocytes.

To assess whether the exacerbated disease severity in NFAT1KO mice is associated with an alteration of Th1- and Th17-related cytokines, we measured cytokine levels in EAMG mice. Compared with WT/EAMG mice, NFAT1KO/EAMG mice showed a higher expression level of Th1/Th17-type cytokines in serum (Fig. 2A), mixed lymphocytes (Supplemental Fig. 2A–C), and splenocytes (Supplemental Fig. 2D, 2E), as well as increased Th1/Th17-type, cytokine-positive, CD4+ T cells (Fig. 2B). Alteration of Il-10 and Tgf-β levels was also observed in the dLN lymphocytes (Supplemental Fig. 2A) and splenocytes (Supplemental Fig. 2D). Increased expression of Th1/Th17-type cytokines was accompanied by increased frequency of Th1- or Th17-related transcription factors such as T-bet and RORγt, positive CD4+ T cells (Fig. 2C, 2D), and mixed lymphocytes (Supplemental Fig. 2F–H). In addition, CD4+ T cells isolated from NFAT1KO/EAMG expressed higher levels of Th1/Th17 cytokines when stimulated with AChR in the presence of mitomycin C–treated splenocytes prepared from normal healthy mice (Fig. 2E, 2F). However, no significant differences were observed in Foxp3 and Bcl-6 levels (Supplemental Fig. 2I), although alterations in Treg and Tfh cells were reported in the development of MG and EAMG (34, 35). These results suggest that NFAT1 deficiency leads to an increase in Th1/Th17 cells and their effector proinflammatory cytokines in EAMG mice.

Because the differentiation of naive CD4+ T cells into Th1/Th17 cells requires a proinflammatory cytokine milieu that is provided by APCs, we determined whether NFAT1KO APCs produce higher levels of Th1/Th17-inducing cytokines. Indeed, CD19+ B cells from the NFAT1KO/EAMG mice showed substantially higher levels of proinflammatory cytokines, such as IL-1β, IL-17A, and IFN-γ (Supplemental Fig. 3A, 3B). Splenic CD11c+ DCs from NFAT1KO/EAMG displayed a more activated phenotype (Supplemental Fig. 3E, 3F) and produced substantially higher levels of proinflammatory cytokines, such as IL-1β, IL-6, IL-12, and TNF-α (Fig. 3A, 3B). Moreover, compared with CD11c+ DCs from WT/EAMG, DCs from NFAT1KO/EAMG are more potent in inducing AChR-reactive T cell proliferation (Fig. 3C) when total CD4+ T cells from WT/EAMG were cultured in the presence of AChR.

Next, we examined the functional activity of CD19+ B cells and DCs to induce Ag (AChR)-specific T cell proliferation and Th1/Th17 cell differentiation. For this purpose, total CD4+ T cells, CD11c+ DCs, or CD19+ B cells were isolated from WT/EAMG mice and NFAT1KO/EAMG mice, and they then were cocultured in a diverse combination in the presence of AChR. Compared with DCs isolated from WT/EAMG, CD4+ T cells cocultured with DCs isolated from NFAT1KO/EAMG mice produced substantially higher expression levels of IFN-γ and IL-17. The highest expression levels were observed in a combination of NFAT1KO/EAMG DCs and NFAT1KO/EAMG CD4+ T cells (KO DC/KO T in Fig. 3D, 3E). Compared with other combinations, the highest level of proinflammatory cytokines was observed in a combination of NFAT1KO/EAMG CD19+ B cells and NFAT1KO/EAMG total CD4+ T cells (KO T/KO B in Supplemental Fig. 3C, 3D). However, compared with WT B cells, NFAT1KO B cells did not show an increase in IFN-γ and IL-17 levels when they were cocultured with WT CD4+ T cells (Supplemental Fig. 3D). Significant differences were observed only when NFAT1KO CD4+ T cells were used. In these ex vivo culture conditions, NFAT1-deficient CD4+ T cells might be the major cell types producing high levels of Th1/Th17 cytokines. It seems likely that already committed Th1/Th17 cells in NFAT1KO/EAMG are further activated to produce effector cytokines upon stimulation with DCs.

We also investigated whether NFAT1KO DCs could induce higher Th1/Th17-type immune responses than WT DCs under non-EAMG conditions. BMDCs prepared from healthy WT and NFAT1KO mice were cocultured with OT-II total CD4+ T cells in the presence of OVA peptide, and both T cell proliferation and cytokine production were determined. Compared with WT BMDCs, NFAT1KO BMDCs significantly enhanced T cell proliferation (Supplemental Fig. 3G) and upregulated the expression levels of proinflammatory cytokines (Supplemental Fig. 3H). In addition, NFAT1KO BMDCs prepared from healthy mice induced an enhanced AChR-reactive Th1/Th17 response when they were cultured with total CD4+ T cells isolated from WT/EAMG-induced mice (Fig. 4A, 4B). In this procedure, the addition of neutralization Abs specific for IL-1β, IL-6, and IL-12 (or a combination of Abs) significantly inhibited the expression levels of the Th1-type cytokine (IFN-γ) and Th17-type cytokine (IL-17A) (Fig. 4C).

To further verify the specific role of NFAT1 in DCs for inducing Th1/Th17 cell differentiation in EAMG development, we performed an adoptive transfer experiment. Because DC-specific conditional NFAT1KO mice are not available, we performed reconstitution experiments using DC-depleted mice. To deplete the DCs, CD11c.DTR transgenic (CD11c.DOG) mice were injected with DT 1 d ahead of DC transfer and then for an additional 7 d (Supplemental Fig. 3I). BMDCs prepared from WT and NFAT1KO mice were transferred into DC-depleted mice, and EAMG was then induced through AChR immunization on the same day as BMDCs transfer (Supplemental Fig. 3I). As reported elsewhere (36), daily administration of DT led to an ∼80% depletion of MHCII+CD11c+ DCs when analyzed 7 d after treatment (Supplemental Fig. 3J). Because depletion of DCs by DT could last <2 wk (33), we measured Th1 and Th17 cytokine levels in the dLN mixed lymphocytes stimulated with AChR. Indeed, transferring NFAT1KO BMDCs into DC-depleted mice induced a significant increase in Th1/Th17-type cytokines compared with those in WT BMDC-transferred mice (Fig. 4D). These results suggest that the absence of NFAT1 in DCs preferentially induces pathogenic Th1/Th17 cells, which mediate the exacerbation of EAMG progression.

We investigated whether the enhanced EAMG susceptibility in NFAT1KO mice could be attributed to T cell–intrinsic or –extrinsic effects. As previously reported (10), the absence of NFAT1 did not cause any significant defects in lymphocyte development (data not shown). Rather, CD4+ T cells isolated from NFAT1KO mice showed a defect in the differentiation of Th1, Th2, and Th17 cells in vitro (Supplemental Fig. 4A). No significant difference was observed in the levels of CD4+Foxp3+ Treg cells between the WT and NFAT1KO mice (Supplemental Fig. 4B). CD19+ B cells also had no significant defect in proliferation capacity or cytokine production (Supplemental Fig. 4C). No significant difference was observed in the population of conventional DCs and plasmacytoid DCs (Supplemental Fig. 4D). However, splenic DCs or BMDCs from NFAT1KO showed a slight increase in IL-1β, IL-6, and TGF-β cytokines; whereas IL-2 levels were significantly reduced (Supplemental Fig. 4E, 4F).

To further characterize the role of NFAT1 in the regulation of DC properties, we performed a microarray experiment using sorted CD11c+ BMDCs from WT and NFAT1KO mice in the absence or presence of Pam3 stimulation. Under unstimulated conditions, 242 genes were differentially expressed, but their differences were not distinctive, which suggests that the transcriptional profiles of NFAT1KO BMDCs were similar to WT BMDCs in steady status (Fig. 5A). However, Pam3 stimulation (2 and 12 h) led to significant differences between WT and NFAT1KO BMDCs (Fig. 5A). We found that most of the upregulated molecules in NFAT1KO BMDCs are known targets of NF-κB. The differentially expressed NF-κB target genes were broadly classified into four different modules: cytokine/chemokines and their modulators (Ccl22, Cx3cl1, Cxcl9, Cxcl10, Il-6, and Il-27), immune receptors (Cd40, Cd80, Cd83, Cd86, and Ccr7), transcription factors (Relb, Irf1, and Stat5a), and apoptosis (Traf1) (Fig. 5B–D). Among the NF-κB target genes, we selected a representative subset of 40 genes (Fig. 5A) and confirmed their differential mRNA or protein levels (Fig. 5B–D). NFAT1KO BMDCs also showed a higher expression of costimulatory molecules (Supplemental Fig. 4G). These results imply that a lack of NFAT1 in the DCs may lead to significant increases in proinflammatory cytokines by preferentially activating NF-κB signaling pathways.

To test this possibility, we measured the expression levels of NF-κB proteins and MAPK-regulated signaling molecules. Indeed, compared with WT BMDCs, NFAT1KO BMDCs showed increased expression of JNK, phospho-JNK, phospho-p38, and Erk as well as decreased IκBα proteins (Fig. 5E). The nuclear expression levels of NF-κB p50 and phospho-c-Jun were also substantially higher, whereas the cytosolic IκBα expression level was significantly lower in NFAT1KO BMDCs (Fig. 5F). In addition, the overexpression of NFAT1 in WT BMDCs or the reconstitution of NFAT1 in NFAT1KO BMDCs significantly downregulated the expression levels of proinflammatory cytokines, such as Il-1b, Il-6, Il-12p40, and Tnf-α (Fig. 5G). As a positive control, we confirmed that NFAT1 overexpression enhanced the expression levels of Il-2 and Il-10 cytokines in both WT and NFAT1KO BMDCs (Fig. 5H).

To confirm the role of NF-κB as a positive regulator of the proinflammatory cytokines in NFAT1KO DCs, we performed a ChIP experiment using an NF-κB p50 Ab. BMDCs prepared from WT and NFAT1KO mice were stimulated with Pam3 for 1 h, and the amount of NF-κB p50 binding to the promoters of IL-6 and IL-12 was determined. Indeed, NF-κB p50 binding was significantly enriched in NFAT1KO BMDCs compared with that in WT BMDCs after Pam3 stimulation (Fig. 5I).

Next, we assessed the possibility that NFAT1 and NF-κB may compete with each other to repress or activate the expression of proinflammatory cytokines, respectively. Indeed, NFAT1 overexpression reduced NF-κB–driven IL-6 or IL-12 reporter activity (Fig. 5J). We also verified enhanced NF-κB activity in NFAT1KO BMDCs using a reporter assay. Either NF-κB or IL-6 reporter constructs were transfected into WT BMDCs or NFAT1KO BMDCs, and the relative luciferase activity was measured. Indeed, NFAT1KO BMDCs showed substantially higher NF-κB– and IL-6–inducing activity than WT BMDCs (Fig. 5K). The results that NFAT1 reconstitution or its overexpression reduced NF-κB–driven IL-6 or IL-12 reporter activity (Fig. 5J), as well as their expression levels (Fig. 5G), collectively suggest that NFAT1 deficiency leads to a significant increase in proinflammatory cytokines through the hyperactivation of the NF-κB–mediated signaling cascade in DCs.

The results described above collectively suggest that NFAT1-deficient DCs facilitate the differentiation of Th1 and Th17 cells, which exacerbates EAMG progression in NFAT1KO mice. Because the important role of IL-6 and IL-12 cytokines in the development of EAMG progression was already reported (17, 22), we determined whether blocking Th1 (IL-12)– or Th17 (IL-6)–inducing cytokines could suppress EAMG progression in NFAT1KO mice. Anti–IL-6 or anti–IL-12 neutralization Abs were administered i.p. three times a week in the course of EAMG development. Administration of neutralization Abs for IL-6 or IL-12 in NFAT1KO/EAMG mice significantly reduced the clinical score to a level similar to that of WT/EAMG mice (Fig. 6A, Table III), and the levels of Th1/Th17-type proinflammatory cytokines were reduced in CD4+ T cells, dLNs, and the spleen (Fig. 6B–D). These results suggest that IL-6 and IL-12 cytokines are key mediators of the exacerbation of EAMG progression in NFAT1KO mice.

In this study, we uncovered a novel function of transcription factor NFAT1 as a key regulator of DC tolerance in systemic autoimmunity. Through a loss-of-gene-function study, we demonstrated that the exacerbated EAMG progression in NFAT1KO mice is closely associated with the upregulation of Th1/Th17-type immune responses. The lack of NFAT1 in DCs induced the hyperactivation of NF-κB–mediated proinflammatory cytokine production, which enhanced pathogenic Th1/Th17 differentiation. Treatment with anti–IL-12 or anti–IL-6 Abs ameliorated EAMG progression in NFAT1KO mice by inhibiting Th1/Th17 immune responses. Our studies suggest that NFAT1 may regulate DC tolerance by suppressing the production of proinflammatory cytokines through the inhibition of NF-κB–mediated systemic autoimmunity.

Previous studies have reported that NFAT1 plays a crucial role in activation and differentiation of CD4+ T cells (2, 8), T cell anergy (37), as well as in regulation of CD8+ T cell activation and exhaustion (12). Functional diversity of NFAT1 in the regulation of immunity and tolerance depends on its associated partners and their availability (12). However, the role of NFAT1 in innate immune cells has been poorly investigated, especially in systemic autoimmunity. In this study, we found that NFAT1KO mice were more susceptible to EAMG induction because of the upregulation of Th1- and Th17-type cytokines. We questioned whether the enhancement of Th1/Th17 cells was a T cell–intrinsic or DC-mediated process. We found that, as also previously reported, NFAT1KO mice showed only moderately enhanced T cell and B cell responses, with a bias toward Th2 cell responses (9, 10) and did not show any significant increase in Th1/Th17-type cells under homeostatic conditions. Instead, NFAT1KO CD4+ T cells were defective in the in vitro differentiation of Th1/Th17 cells (Supplemental Fig. 4A).

We evaluated whether the enhanced EAMG development in the NFAT1KO mice was because of any defects in the Foxp3+ Treg population, because Treg cells play crucial roles in MG development (34). Although Foxp3 expression was comparable (Supplemental Fig. 4B), NFAT1-deficient Treg cells showed a reduced mean fluorescence intensity of Foxp3 expression in the steady state (38). However, the role of NFAT1 for peripherally induced Treg differentiation is still unclear. Vaeth et al. (39) reported that peripherally induced Treg differentiation was reduced in NFAT2fl/fl-CD4-cre and NFAT1−/−/NFAT2fl/fl-CD4-cre but not in NFAT1−/− mice. NFAT1−/− CD4+ T cells showed only a marginal defect to differentiate into inducible Tregs under TGF-β and IL-2 conditions, which suggests that the level of Foxp3 expression in induced Tregs depends on the threshold value of NFAT rather than on the individual members that are present (39). In addition, the natural Treg frequency remained unaltered in mice lacking NFAT1, NFAT2, or NFAT4 alone or in combination (39). Treg cells derived from NFAT1−/−NFAT4−/− double knockout cells appear to be functional (40). NFAT1 may play an important role in mediating Treg-mediated suppression of Th cells by inducing the transcriptional repressor Ikaros. NFAT1-deficient Th cells showed resistance to Treg-mediated suppression activation (41). These results suggest that exacerbated Th1/Th17 immune responses in the NFAT1KO/EAMG mice may not be due to a significant defect in Treg cells, although we cannot completely rule out the possibility.

Recently, the role of Tfh cells in MG development was reported (35, 42, 43). To assess whether NFAT1 deficiency affects Tfh cell development, we measured the transcription factor Bcl-6, which is known as a major Tfh cell marker (44). We could not detect any significant differences in the Bcl-6 levels between the T cells of WT and NFAT1KO mice (Supplemental Fig. 2I). In addition, NFAT1KO CD19+ B cells also showed a normal proliferation capacity and cytokine production after BCR or TLR stimulation. These results indicate that the exacerbated EAMG progression in NFAT1KO mice may not be caused by an alteration in the intrinsic properties of CD4+ T and B cells.

Next, we tested the possibility that NFAT1 deficiency in DCs may lead to an alteration in the functional properties of DCs to enhance Th1/Th17 cell differentiation. DCs regulate the adaptive immune response by controlling the activation and differentiation of naive T cells into diverse effector T cells, such as Th1, Th2, Th17, Tfh cells, or Treg cells (45). Previous studies showed that uncontrolled activated DC phenotypes can cause autoimmune diseases (46), and NFAT1-deficient DCs preferentially induced the Th1 phenotype by enhancing IL-12 production (13). However, the role of DCs in the pathogenesis of EAMG has been poorly explored. We found that compared with WT DCs, NFAT1KO DCs showed more activated phenotypes and produced higher levels of proinflammatory cytokines which can preferentially induce Th1/Th17 cell differentiation (Fig. 3, Supplemental Fig. 3E, 3F). Comparative analysis of the gene expression profile between WT and NFAT1KO BMDCs showed that NFAT1 deficiency leads to a significant upregulation of proinflammatory mediators, which are well-known NF-κB target genes, and hyperactivation of TLR downstream signaling pathways (Fig. 5A–F). In addition, in the absence of NFAT1, physical binding of NF-κB (p50) to the promoters of IL-6 and IL-12 was significantly increased and enhanced their activity (Fig. 5I, 5J). A previous report also suggested a role for NF-κB activation in DCs and macrophages. The adoptive transfer of NF-κB knockdown DCs ameliorates EAMG development (47), whereas the absence of NFAT1 in DCs increases IL-12p35–mediated IFN-γ production in CD4+ T cells (13). However, the role of NFAT1 in regulation of macrophage properties is still unclear. We found that, unlike BMDCs, bone marrow–derived macrophages from NFAT1KO mice did not show any increase in the level of proinflammatory cytokines (data not shown). However, further studies are required to define the exact role of NFAT1 in regulation of activation and differentiation of macrophages.

Although it is still unclear how NFAT1 regulates DC tolerance, diverse mechanisms, which are not mutually exclusive, might be involved in this process. These include hyperactivation of TLR-mediated MAPK signaling or competing with NF-κB to regulate common target genes. Microarray data showed that, among the diverse TLRs, only the TLR2 level was significantly reduced; whereas levels of costimulatory factors were increased in the CD11c+ BMDCs of NFAT1KO mice. However, Western blot analysis showed that NFAT1 deficiency led to more activated MAPK signaling, suggesting that NFAT1 deficiency may lead to hyperactivation of TLR-induced MAPK signaling through an unknown mechanism. Another possibility is that NFAT1 may repress the production of proinflammatory cytokines by inhibiting NF-κB activity. A previously published report also supports the possibility that calcineurin activity is important to prevent the hyperactivation of NF-κB and MAPK-activation pathways (48). Compared with resting T cells, resting macrophages have higher levels of intracellular free calcium, which leads to the basal activation of NFAT proteins (49, 50). This activation could occur in the DCs, which could support a basal level of NFAT1 activity to silence NF-κB–mediated proinflammatory signaling pathways. Indeed, we found that the overexpression of NFAT1 in WT DCs or the reconstitution of NFAT1 expression in NFAT1KO DCs reduced the expression levels of proinflammatory cytokines (Fig. 5G). NF-κB (p50) –mediated activation of IL-6 and IL-12 promoters was significantly inhibited by NFAT1 expression (Fig. 5J). These results suggest that NFAT1 may compete with NF-κB for binding sites as observed in several target genes, such as the HIV-1 long terminal repeat (51, 52), GM-CSF (53), and Th cells (54). This competitive binding may be due to the high similarity of NFAT1 and NF-κB at the sequence and structural levels (55, 56). NFAT1 ChIP-sequencing data also suggest that NFAT1 binds to multiple proinflammatory cytokine genes, such as Il-1a, Il-1b, Il-17d, Il-22, Il-12b, and Il-23a, which have been previously identified as targets of NF-κB (57).

In addition to these possibilities, the proinflammatory properties of NFAT1KO DCs may be due to the reduced production of IL-2 (Supplemental Fig. 4F), because IL-2 suppresses the differentiation of Th17 cells while supporting Treg survival (58). In addition, because NFAT1KO T cells are more resistant to activation-induced cell death (38), accumulation of pathological Th1 and Th17 cells may also contribute to the exacerbated EAMG progression in NFAT1KO mice.

In summary, our data suggest that NFAT1 may act as a key regulator of DC tolerance under the pathogenesis of systemic autoimmunity by controlling the hyperactivation of the NF-κB–mediated proinflammatory immune response. Proper controlling of NF-κB activity when calcineurin inhibitors are employed might therefore be more effective in treating inflammatory immune disorders.

We thank Dr. Dipayan Rudra and Dr. Hye-Ran Kim for editing the manuscript and for technical support for imaging analysis.

This work was supported by Grant IBS-R005-G1 (to S.-H.I.) from the Institute for Basic Science, Korean Ministry of Science, Information/Communication Technology and Future Planning.

The RNA-sequencing data presented in this article have been submitted to the Gene Expression Omnibus (http://www.ncbi.nlm.nih.gov/geo/) under accession number GSE92593.

The online version of this article contains supplemental material.

Abbreviations used in this article:

AChR

acetylcholine receptor

BMDC

bone marrow DC

ChIP

chromatin immunoprecipitation

DC

dendritic cell

dLN

draining lymph node

DT

diphtheria toxin

DTR

DT receptor

EAMG

experimental autoimmune MG

KO

knockout

MG

myasthenia gravis

NFAT1KO

NFAT1 knockout

NFAT1KO/EAMG

EAMG induced in NFAT1KO

NMJ

neuromuscular junction

Pam3

Pam3 CSK

qRT-PCR

quantitative real-time PCR

TAChR

Torpedo AChR

Tfh

follicular T helper cell

Treg

regulatory T cell

WT

wild type

WT/EAMG

EAMG induced in WT.

1
Hogan
,
P. G.
,
L.
Chen
,
J.
Nardone
,
A.
Rao
.
2003
.
Transcriptional regulation by calcium, calcineurin, and NFAT.
Genes Dev.
17
:
2205
2232
.
2
Macian
,
F.
2005
.
NFAT proteins: key regulators of T-cell development and function.
Nat. Rev. Immunol.
5
:
472
484
.
3
Zanoni
,
I.
,
R.
Ostuni
,
G.
Capuano
,
M.
Collini
,
M.
Caccia
,
A. E.
Ronchi
,
M.
Rocchetti
,
F.
Mingozzi
,
M.
Foti
,
G.
Chirico
, et al
.
2009
.
CD14 regulates the dendritic cell life cycle after LPS exposure through NFAT activation.
Nature
460
:
264
268
.
4
Müller
,
M. R.
,
A.
Rao
.
2010
.
NFAT, immunity and cancer: a transcription factor comes of age.
Nat. Rev. Immunol.
10
:
645
656
.
5
Azzi
,
J. R.
,
M. H.
Sayegh
,
S. G.
Mallat
.
2013
.
Calcineurin inhibitors: 40 years later, can’t live without ...
.
J. Immunol.
191
:
5785
5791
.
6
Muraoka
,
K.
,
K.
Fujimoto
,
X.
Sun
,
K.
Yoshioka
,
K.
Shimizu
,
M.
Yagi
,
H.
Bose
Jr.
,
I.
Miyazaki
,
K.
Yamamoto
.
1996
.
Immunosuppressant FK506 induces interleukin-6 production through the activation of transcription factor nuclear factor (NF)-kappa(B). Implications for FK506 nephropathy.
J. Clin. Invest.
97
:
2433
2439
.
7
Martinez
,
G. J.
,
J. K.
Hu
,
R. M.
Pereira
,
J. S.
Crampton
,
S.
Togher
,
N.
Bild
,
S.
Crotty
,
A.
Rao
.
2016
.
Cutting edge: NFAT transcription factors promote the generation of follicular helper T cells in response to acute viral infection.
J. Immunol.
196
:
2015
2019
.
8
Hermann-Kleiter
,
N.
,
G.
Baier
.
2010
.
NFAT pulls the strings during CD4+ T helper cell effector functions.
Blood
115
:
2989
2997
.
9
Hodge
,
M. R.
,
A. M.
Ranger
,
F.
Charles de la Brousse
,
T.
Hoey
,
M. J.
Grusby
,
L. H.
Glimcher
.
1996
.
Hyperproliferation and dysregulation of IL-4 expression in NF-ATp-deficient mice.
Immunity
4
:
397
405
.
10
Xanthoudakis
,
S.
,
J. P.
Viola
,
K. T.
Shaw
,
C.
Luo
,
J. D.
Wallace
,
P. T.
Bozza
,
D. C.
Luk
,
T.
Curran
,
A.
Rao
.
1996
.
An enhanced immune response in mice lacking the transcription factor NFAT1.
Science
272
:
892
895
.
11
Rengarajan
,
J.
,
B.
Tang
,
L. H.
Glimcher
.
2002
.
NFATc2 and NFATc3 regulate T(H)2 differentiation and modulate TCR-responsiveness of naïve T(H)cells.
Nat. Immunol.
3
:
48
54
.
12
Martinez
,
G. J.
,
R. M.
Pereira
,
T.
Äijö
,
E. Y.
Kim
,
F.
Marangoni
,
M. E.
Pipkin
,
S.
Togher
,
V.
Heissmeyer
,
Y. C.
Zhang
,
S.
Crotty
, et al
.
2015
.
The transcription factor NFAT promotes exhaustion of activated CD8+ T cells.
Immunity
42
:
265
278
.
13
Barboza
,
B. A.
,
B. P.
Fonseca
,
J. P.
Viola
.
2014
.
NFAT1 transcription factor in dendritic cells is required to modulate T helper cell differentiation.
Immunobiology
219
:
704
712
.
14
Toyka
,
K. V.
,
D. B.
Brachman
,
A.
Pestronk
,
I.
Kao
.
1975
.
Myasthenia gravis: passive transfer from man to mouse.
Science
190
:
397
399
.
15
Pinching
,
A. J.
,
D. K.
Peters
.
1976
.
Remission of myasthenia gravis following plasma-exchange.
Lancet
2
:
1373
1376
.
16
Newsom-Davis
,
J.
,
A. J.
Pinching
,
A.
Vincent
,
S. G.
Wilson
.
1978
.
Function of circulating antibody to acetylcholine receptor in myasthenia gravis: investigation by plasma exchange.
Neurology
28
:
266
272
.
17
Kaul
,
R.
,
M.
Shenoy
,
E.
Goluszko
,
P.
Christadoss
.
1994
.
Major histocompatibility complex class II gene disruption prevents experimental autoimmune myasthenia gravis.
J. Immunol.
152
:
3152
3157
.
18
Tüzün
,
E.
,
R.
Huda
,
P.
Christadoss
.
2011
.
Complement and cytokine based therapeutic strategies in myasthenia gravis.
J. Autoimmun.
37
:
136
143
.
19
Moiola
,
L.
,
F.
Galbiati
,
G.
Martino
,
S.
Amadio
,
E.
Brambilla
,
G.
Comi
,
A.
Vincent
,
L. M.
Grimaldi
,
L.
Adorini
.
1998
.
IL-12 is involved in the induction of experimental autoimmune myasthenia gravis, an antibody-mediated disease.
Eur. J. Immunol.
28
:
2487
2497
.
20
Duan
,
R. S.
,
H. B.
Wang
,
J. S.
Yang
,
B.
Scallon
,
H.
Link
,
B. G.
Xiao
.
2002
.
Anti-TNF-alpha antibodies suppress the development of experimental autoimmune myasthenia gravis.
J. Autoimmun.
19
:
169
174
.
21
Wang
,
W.
,
M.
Milani
,
N.
Ostlie
,
D.
Okita
,
R. K.
Agarwal
,
R. R.
Caspi
,
B. M.
Conti-Fine
.
2007
.
C57BL/6 mice genetically deficient in IL-12/IL-23 and IFN-gamma are susceptible to experimental autoimmune myasthenia gravis, suggesting a pathogenic role of non-Th1 cells. [Published erratum appears in 2007 J. Immunol. 179: 7184.]
J. Immunol.
178
:
7072
7080
.
22
Aricha
,
R.
,
K.
Mizrachi
,
S.
Fuchs
,
M. C.
Souroujon
.
2011
.
Blocking of IL-6 suppresses experimental autoimmune myasthenia gravis.
J. Autoimmun.
36
:
135
141
.
23
Schaffert
,
H.
,
A.
Pelz
,
A.
Saxena
,
M.
Losen
,
A.
Meisel
,
A.
Thiel
,
S.
Kohler
.
2015
.
IL-17-producing CD4(+) T cells contribute to the loss of B-cell tolerance in experimental autoimmune myasthenia gravis.
Eur. J. Immunol.
45
:
1339
1347
.
24
Yi
,
H. J.
,
C. S.
Chae
,
J. S.
So
,
S. J.
Tzartos
,
M. C.
Souroujon
,
S.
Fuchs
,
S. H.
Im
.
2008
.
Suppression of experimental myasthenia gravis by a B-cell epitope-free recombinant acetylcholine receptor.
Mol. Immunol.
46
:
192
201
.
25
Nelson
,
J. D.
,
O.
Denisenko
,
K.
Bomsztyk
.
2006
.
Protocol for the fast chromatin immunoprecipitation (ChIP) method.
Nat. Protoc.
1
:
179
185
.
26
Turroni
,
F.
,
E.
Foroni
,
P.
Pizzetti
,
V.
Giubellini
,
A.
Ribbera
,
P.
Merusi
,
P.
Cagnasso
,
B.
Bizzarri
,
G. L.
de’Angelis
,
F.
Shanahan
, et al
.
2009
.
Exploring the diversity of the bifidobacterial population in the human intestinal tract.
Appl. Environ. Microbiol.
75
:
1534
1545
.
27
Liu
,
J.
,
D. I.
Beller
.
2003
.
Distinct pathways for NF-kappa B regulation are associated with aberrant macrophage IL-12 production in lupus- and diabetes-prone mouse strains.
J. Immunol.
170
:
4489
4496
.
28
Kang
,
J. A.
,
S. P.
Jeong
,
D.
Park
,
M. S.
Hayden
,
S.
Ghosh
,
S. G.
Park
.
2013
.
Transition from heterotypic to homotypic PDK1 homodimerization is essential for TCR-mediated NF-κB activation.
J. Immunol.
190
:
4508
4515
.
29
Ryu
,
J. H.
,
S.
Yang
,
Y.
Shin
,
J.
Rhee
,
C. H.
Chun
,
J. S.
Chun
.
2011
.
Interleukin-6 plays an essential role in hypoxia-inducible factor 2α-induced experimental osteoarthritic cartilage destruction in mice.
Arthritis Rheum.
63
:
2732
2743
.
30
Lutz
,
M. B.
,
N.
Kukutsch
,
A. L.
Ogilvie
,
S.
Rössner
,
F.
Koch
,
N.
Romani
,
G.
Schuler
.
1999
.
An advanced culture method for generating large quantities of highly pure dendritic cells from mouse bone marrow.
J. Immunol. Methods
223
:
77
92
.
31
Datta
,
S. K.
,
S.
Okamoto
,
T.
Hayashi
,
S. S.
Shin
,
I.
Mihajlov
,
A.
Fermin
,
D. G.
Guiney
,
J.
Fierer
,
E.
Raz
.
2006
.
Vaccination with irradiated Listeria induces protective T cell immunity.
Immunity
25
:
143
152
.
32
Helft
,
J.
,
J.
Böttcher
,
P.
Chakravarty
,
S.
Zelenay
,
J.
Huotari
,
B. U.
Schraml
,
D.
Goubau
,
C.
Reis e Sousa
.
2015
.
GM-CSF mouse bone marrow cultures comprise a heterogeneous population of CD11c(+)MHCII(+) macrophages and dendritic cells.
Immunity
42
:
1197
1211
.
33
Hochweller
,
K.
,
J.
Striegler
,
G. J.
Hämmerling
,
N.
Garbi
.
2008
.
A novel CD11c.DTR transgenic mouse for depletion of dendritic cells reveals their requirement for homeostatic proliferation of natural killer cells.
Eur. J. Immunol.
38
:
2776
2783
.
34
Gertel-Lapter
,
S.
,
K.
Mizrachi
,
S.
Berrih-Aknin
,
S.
Fuchs
,
M. C.
Souroujon
.
2013
.
Impairment of regulatory T cells in myasthenia gravis: studies in an experimental model.
Autoimmun. Rev.
12
:
894
903
.
35
Luo
,
C.
,
Y.
Li
,
W.
Liu
,
H.
Feng
,
H.
Wang
,
X.
Huang
,
L.
Qiu
,
J.
Ouyang
.
2013
.
Expansion of circulating counterparts of follicular helper T cells in patients with myasthenia gravis.
J. Neuroimmunol.
256
:
55
61
.
36
Phythian-Adams
,
A. T.
,
P. C.
Cook
,
R. J.
Lundie
,
L. H.
Jones
,
K. A.
Smith
,
T. A.
Barr
,
K.
Hochweller
,
S. M.
Anderton
,
G. J.
Hämmerling
,
R. M.
Maizels
,
A. S.
MacDonald
.
2010
.
CD11c depletion severely disrupts Th2 induction and development in vivo.
J. Exp. Med.
207
:
2089
2096
.
37
Macián
,
F.
,
F.
García-Cózar
,
S. H.
Im
,
H. F.
Horton
,
M. C.
Byrne
,
A.
Rao
.
2002
.
Transcriptional mechanisms underlying lymphocyte tolerance.
Cell
109
:
719
731
.
38
Kwon
,
H. K.
,
G. C.
Kim
,
J. S.
Hwang
,
Y.
Kim
,
C. S.
Chae
,
J. H.
Nam
,
C. D.
Jun
,
D.
Rudra
,
C. D.
Surh
,
S. H.
Im
.
2016
.
Transcription factor NFAT1 controls allergic contact hypersensitivity through regulation of activation induced cell death program.
Sci. Rep.
6
:
19453
.
39
Vaeth
,
M.
,
U.
Schliesser
,
G.
Müller
,
S.
Reissig
,
K.
Satoh
,
A.
Tuettenberg
,
H.
Jonuleit
,
A.
Waisman
,
M. R.
Müller
,
E.
Serfling
, et al
.
2012
.
Dependence on nuclear factor of activated T-cells (NFAT) levels discriminates conventional T cells from Foxp3+ regulatory T cells.
Proc. Natl. Acad. Sci. USA
109
:
16258
16263
.
40
Bopp
,
T.
,
A.
Palmetshofer
,
E.
Serfling
,
V.
Heib
,
S.
Schmitt
,
C.
Richter
,
M.
Klein
,
H.
Schild
,
E.
Schmitt
,
M.
Stassen
.
2005
.
NFATc2 and NFATc3 transcription factors play a crucial role in suppression of CD4+ T lymphocytes by CD4+ CD25+ regulatory T cells.
J. Exp. Med.
201
:
181
187
.
41
Shin
,
D. S.
,
A.
Jordan
,
S.
Basu
,
R. M.
Thomas
,
S.
Bandyopadhyay
,
E. F.
de Zoeten
,
A. D.
Wells
,
F.
Macian
.
2014
.
Regulatory T cells suppress CD4+ T cells through NFAT-dependent transcriptional mechanisms.
EMBO Rep.
15
:
991
999
.
42
Xie
,
X.
,
L.
Mu
,
X.
Yao
,
N.
Li
,
B.
Sun
,
Y.
Li
,
X.
Zhan
,
X.
Wang
,
X.
Kang
,
J.
Wang
, et al
.
2013
.
ATRA alters humoral responses associated with amelioration of EAMG symptoms by balancing Tfh/Tfr helper cell profiles.
Clin. Immunol.
148
:
162
176
.
43
Zhang
,
C. J.
,
Y.
Gong
,
W.
Zhu
,
Y.
Qi
,
C. S.
Yang
,
Y.
Fu
,
G.
Chang
,
Y.
Li
,
S.
Shi
,
K.
Wood
, et al
.
2016
.
Augmentation of circulating follicular helper T cells and their impact on autoreactive B cells in myasthenia gravis.
J. Immunol.
197
:
2610
2617
.
44
Vinuesa
,
C. G.
,
M. A.
Linterman
,
D.
Yu
,
I. C.
MacLennan
.
2016
.
Follicular helper T cells.
Annu. Rev. Immunol.
34
:
335
368
.
45
Walsh
,
K. P.
,
K. H.
Mills
.
2013
.
Dendritic cells and other innate determinants of T helper cell polarisation.
Trends Immunol.
34
:
521
530
.
46
Ganguly
,
D.
,
S.
Haak
,
V.
Sisirak
,
B.
Reizis
.
2013
.
The role of dendritic cells in autoimmunity.
Nat. Rev. Immunol.
13
:
566
577
.
47
Zhang
,
Y.
,
H.
Yang
,
B.
Xiao
,
M.
Wu
,
W.
Zhou
,
J.
Li
,
G.
Li
,
P.
Christadoss
.
2009
.
Dendritic cells transduced with lentiviral-mediated RelB-specific ShRNAs inhibit the development of experimental autoimmune myasthenia gravis.
Mol. Immunol.
46
:
657
667
.
48
Kang
,
Y. J.
,
B.
Kusler
,
M.
Otsuka
,
M.
Hughes
,
N.
Suzuki
,
S.
Suzuki
,
W. C.
Yeh
,
S.
Akira
,
J.
Han
,
P. P.
Jones
.
2007
.
Calcineurin negatively regulates TLR-mediated activation pathways.
J. Immunol.
179
:
4598
4607
.
49
Beppu
,
M.
,
M.
Hora
,
T.
Watanabe
,
M.
Watanabe
,
H.
Kawachi
,
E.
Mishima
,
M.
Makino
,
K.
Kikugawa
.
2001
.
Substrate-bound fibronectin enhances scavenger receptor activity of macrophages by calcium signaling.
Arch. Biochem. Biophys.
390
:
243
252
.
50
Korhonen
,
R.
,
H.
Kankaanranta
,
A.
Lahti
,
M.
Lähde
,
R. G.
Knowles
,
E.
Moilanen
.
2001
.
Bi-directional effects of the elevation of intracellular calcium on the expression of inducible nitric oxide synthase in J774 macrophages exposed to low and to high concentrations of endotoxin.
Biochem. J.
354
:
351
358
.
51
Macián
,
F.
,
A.
Rao
.
1999
.
Reciprocal modulatory interaction between human immunodeficiency virus type 1 Tat and transcription factor NFAT1.
Mol. Cell. Biol.
19
:
3645
3653
.
52
Falvo
,
J. V.
,
C. H.
Lin
,
A. V.
Tsytsykova
,
P. K.
Hwang
,
D.
Thanos
,
A. E.
Goldfeld
,
T.
Maniatis
.
2008
.
A dimer-specific function of the transcription factor NFATp.
Proc. Natl. Acad. Sci. USA
105
:
19637
19642
.
53
Shang
,
C.
,
J.
Attema
,
D.
Cakouros
,
P. N.
Cockerill
,
M. F.
Shannon
.
1999
.
Nuclear factor of activated T cells contributes to the function of the CD28 response region of the granulocyte macrophage-colony stimulating factor promoter.
Int. Immunol.
11
:
1945
1956
.
54
Casolaro
,
V.
,
S. N.
Georas
,
Z.
Song
,
I. D.
Zubkoff
,
S. A.
Abdulkadir
,
D.
Thanos
,
S. J.
Ono
.
1995
.
Inhibition of NF-AT-dependent transcription by NF-kappa B: implications for differential gene expression in T helper cell subsets.
Proc. Natl. Acad. Sci. USA
92
:
11623
11627
.
55
Chen
,
L.
,
J. N.
Glover
,
P. G.
Hogan
,
A.
Rao
,
S. C.
Harrison
.
1998
.
Structure of the DNA-binding domains from NFAT, Fos and Jun bound specifically to DNA.
Nature
392
:
42
48
.
56
Jain
,
J.
,
E.
Burgeon
,
T. M.
Badalian
,
P. G.
Hogan
,
A.
Rao
.
1995
.
A similar DNA-binding motif in NFAT family proteins and the Rel homology region.
J. Biol. Chem.
270
:
4138
4145
.
57
Yu
,
H. B.
,
M.
Yurieva
,
A.
Balachander
,
I.
Foo
,
X.
Leong
,
T.
Zelante
,
F.
Zolezzi
,
M.
Poidinger
,
P.
Ricciardi-Castagnoli
.
2015
.
NFATc2 mediates epigenetic modification of dendritic cell cytokine and chemokine responses to dectin-1 stimulation.
Nucleic Acids Res.
43
:
836
847
.
58
Boyman
,
O.
,
J.
Sprent
.
2012
.
The role of interleukin-2 during homeostasis and activation of the immune system.
Nat. Rev. Immunol.
12
:
180
190
.

The authors have no financial conflicts of interest.

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