To determine the role of leptin in modulating Th17 cell response and joint inflammation in mice with collagen-induced arthritis (CIA).
To determine the role of leptin in modulating Th17 cell response and joint inflammation in mice with collagen-induced arthritis (CIA).
Leptin receptor expression on T cells was examined by polymerase chain reaction (PCR) analysis, immunofluorescence microscopy, and flow cytometry. Effects of leptin on Th17 cell differentiation and proliferation were evaluated by quantitative PCR, carboxyfluorescein diacetate succinimidyl ester proliferation assay, and flow cytometry. Dynamic changes in leptin concentrations in the joint tissue and synovial fluid of mice with CIA were determined by immunohistochemistry analysis and enzyme-linked immunosorbent assay (ELISA). Arthritis symptoms and joint pathology in mice with CIA were assessed after injection of leptin into the knee joint. Th1 and Th17 cell populations in the spleen, draining lymph nodes, and joint tissue were analyzed by flow cytometry and enzyme-linked immunospot assay. Interleukin-17 messenger RNA and protein levels in the joint tissue were measured by PCR analysis and ELISA.
In culture, leptin treatment significantly increased Th17 cell generation from naive CD4+ T cells. During CIA development, markedly elevated levels of leptin were detected in the joint tissue and synovial fluid. Moreover, injection of leptin into the knee joint of collagen-immunized mice resulted in an early onset of arthritis and substantially increased the severity of clinical symptoms, accompanied by more pronounced synovial hyperplasia and joint damage. Further examination by immunofluorescence microscopy confirmed the presence of significantly increased numbers of Th17 cells in the joint tissue and draining lymph nodes of leptin-treated mice with CIA.
The results of this study identify a previously undescribed function of leptin in enhancing Th17 cell response and exacerbating joint inflammation in mice with CIA.
Leptin, a peptide hormone produced by adipose tissue, has been identified as a key player in regulating nutrient intake and metabolism and immune response (1–5). Early studies on leptin-deficient (ob/ob) and leptin receptor–deficient (db/db) mice have demonstrated significantly reduced numbers of both T and B lymphocytes in the peripheral lymphoid organs, with impaired immune responses (6). Accumulating data, including our recent findings that leptin enhances the proliferation and production of proinflammatory cytokines in various immune cells such as T cells, B cells, macrophages, and dendritic cells, have characterized leptin as a proinflammatory cytokine in immune responses (7–12).
There is evidence that leptin deficiency indirectly affects T cell maturation in the thymus and results in marked thymic atrophy through changes in the systemic environment, and several studies have confirmed a direct effect of leptin on T cell activation and function in the peripheral lymphoid system (13, 14). It has become clear that leptin binds to leptin receptor b (Ob-Rb) and induces activation of the STAT-3 pathway in CD4+ T cells (14). Moreover, CD4+ T cells from db/db mice display a significantly decreased proliferative response to anti-CD3/CD28 stimulation (14). Leptin can also enhance the survival of T cells via suppression of Fas-medicated and stress-induced apoptosis (14, 15). The proinflammatory activity of leptin has been clearly demonstrated by the findings that leptin drives Th1 cell polarization and suppresses Treg cell proliferation (16). Recent studies have revealed a key role of leptin in controlling the responsiveness of Treg cells via modulation of the mammalian target of rapamycin pathway (17), suggesting that leptin is critically involved in immune tolerance and autoimmune response. Although the roles of leptin in driving Th1 proinflammatory reaction and controlling Treg cell proliferation are well characterized, whether leptin exerts any direct modulating effect on Th17 cell differentiation and function has not been clear.
Apart from its prominent activities in modulating immune response, leptin has been implicated in the disease process of rheumatoid arthritis (RA) (18–20). There have been some contradictory findings regarding the correlation between arthritis activity and leptin levels in RA. For example, studies by Targonska-Stepniak et al have suggested a possible correlation between increased leptin levels and RA disease activity (21). In contrast, it has also been reported that serum leptin levels are negatively correlated with either markers of systemic inflammation or disease activity in RA patients (22). Intriguingly, no obvious change in serum levels of leptin was detected during anti–tumor necrosis factor treatment in patients with RA, indicating that serum leptin is not linked to inflammatory status in these patients (23). In both ob/ob mice and db/db mice the severity of antigen-induced arthritis has been shown to be decreased, with significantly reduced serum levels of autoantibodies and a shift toward a Th2-type cytokine response, when compared with wild-type controls (24). However, in zymosan-induced arthritis, a model of proliferative arthritis, disease is not reduced in either db/db or ob/ob mice, and resolution of acute inflammation is delayed (25).
The fact that leptin is known to exert multiple neuroendocrine and immune functions on various types of cells and tissues raises the question of whether local leptin concentrations are directly implicated in the progression of synovial inflammation and joint pathology in the context of an intact leptin signaling system. In this study, we aimed to determine the role of leptin in enhancing Th17 response and promoting disease progression in mice with collagen-induced arthritis (CIA).
DBA/1J, wild-type C57BL/6, and db/db C57BL/6 mice were purchased from The Jackson Laboratory. Mice were maintained in a specific pathogen–free animal facility at the University of Hong Kong and allowed access to food and water ad libitum. All animal experiments were approved by the Institutional Animal Care and Use Committee. Mice were 8–12 weeks of age at the time of the experiments.
CIA induction was performed as previously described (26). Briefly, male DBA/1J mice were immunized with 100 μg of bovine type II collagen (CII; Chondrex) dissolved in Freund's complete adjuvant (Difco) via intradermal injection at the base of tail, and a boost emulsion of CII dissolved in Freund's incomplete adjuvant (Difco) was administered on day 21 after the first immunization. Age- and sex-matched DBA/1J mice immunized with adjuvant only were used as controls. Leptin (PeproTech) was injected intraarticularly into the knee joint (5 μg in 10 μl phosphate buffered saline [PBS]) on days 17, 20, and 23 after the first CII immunization. Control CII-immunized DBA/1J mice were administered the same volume of PBS without leptin.
After removal of skin, muscle, and bone under a dissecting microscope, joint samples were minced and incubated with collagenase for 1 hour at 37°C. Cell suspensions were filtered with a cell strainer after red blood cell lysis. Synovial fluid was collected according to a previously described protocol (27), with minor modifications. Briefly, after excision of the skin and patellar ligament under a dissecting microscope to expose the synovial membrane, a 30-gauge needle (BD Biosciences) was carefully inserted into the membrane, and the synovial cavity was washed by repetitive injections and aspirations with 20 μl/PBS to obtain synovial lavage material. The procedure was repeated 5 times, and a total volume of 100 μl of synovial lavage fluid was obtained. Fluid samples were stored at −80°C prior to assay.
Naive CD4+CD62L+ T cells from C57BL/6 and db/db mice were isolated using a CD4+CD62L+ T Cell Isolation Kit II (Miltenyi Biotec) as previously described (28); purity was routinely >95%. Purified naive T cells were cultured in anti-CD3– and anti-CD28–precoated 24-well plates (5 × 105 cells/well) with 3 ng/ml transforming growth factor β, 20 ng/ml interleukin-6 (IL-6), and 20 ng/ml IL-23 (all from R&D Systems) in the presence or absence of leptin (50, 250, or 500 ng/ml) with or without pretreatment with Ob-R:Fc, a soluble blocker of leptin. After 72 hours of culture, cells and supernatants were collected for further analysis. In separate experiments, purified naive CD4+ T cells were labeled with carboxyfluorescein diacetate succinimidyl ester (CFSE) and cultured for 72 hours in the presence or absence of leptin (250 ng/ml) under Th17 polarization conditions as described above, before intracellular staining for IL-17 and flow cytometric analysis of Th17 proliferation. Recombinant murine leptin with an endotoxin level of <0.1 ng/μg (1 endotoxin unit/μg) was purchased from PeproTech.
RNA samples were extracted with TRIzol reagent (Invitrogen), and complementary DNA was prepared using SuperScript III First-Strand Synthesis SuperMix (Invitrogen). PCR primers used for qPCR were as follows: for Leptin (113 bp) sense 5′-AGTCCAGGATGACACCAA-3′, antisense 5′-GAATGAAGTCCAAGCCAGT-3′; for Il17 (142 bp) sense 5′-AGCTTTCCCTCCGCATTGA-3′, anti-sense 5′-GCTCCAGAAGGCCCTCAGA-3′; for Rorc (93 bp, encoding retinoic acid–related orphan nuclear receptor γt [RORγt]) sense 5′-CTGAGGGGCTGTCAAAGTG-3′, antisense 5′-AAGGCTGGGTGAAGGGA-3′; for β-Actin (195 bp) sense 5′-GCGTGACATCAAAGAGAAGCT-3′, antisense 5′-ATGCCACAGGATTCCATACC-3′. Leptin, IL-17, and RORγt transcript levels were measured using an ABI 7900 system with a SYBR Green Two-Step qRT-PCR Kit according to the instructions of the manufacturer (Applied Biosystems) and as previously described (29). Relative expression levels of target genes were calculated with normalization to β-actin values using the Ct method.
Single-cell suspensions were prepared and filtered with a cell strainer. Surface staining was performed using the following monoclonal antibodies: anti-CD4 (clone GK1.5), anti-CD3 (clone 145-2C11), anti–interferon-γ (anti-IFNγ) (clone XMG1.2), and anti–IL-17 (clone TC11-18H10.1) (all from BioLegend). Anti-FoxP3 (clone FHK-16s) was purchased from eBioscience. For intracellular staining of IFNγ and IL-17, phorbol myristate acetate (50 ng/ml; Sigma-Aldrich), ionomycin (500 ng/ml; Sigma-Aldrich), and monensin (2 μM; BioLegend) were added and cultured for the last 5 hours before flow cytometric analysis, as previously described (28).
IL-17–producing CD4+ cells in the inflamed tissue of mice with CIA were identified by ELISpot assay as previously described (26). Briefly, 96-well plates (Millipore) were precoated with rat anti-mouse IL-17 (2 μg/ml; R&D Systems) overnight at 4°C. Purified CD4+ T cells were seeded into wells and incubated at 37°C. After 6 hours of incubation, biotinylated goat anti-mouse IL-17 (200 ng/ml; R&D Systems) was added and incubated for 2 hours, and then alkaline phosphatase–conjugated streptavidin (Invitrogen) was added and incubated for 1 hour at room temperature. Finally, plates were washed and developed by adding BCIP/nitroblue tetrazolium solution (Sigma-Aldrich).
Levels of IL-17 and leptin in the synovial fluid and culture supernatants were measured by colorimetric sandwich ELISA as previously described (29). Briefly, 96-well MaxiSorp plates (Nunc) were coated with goat anti-mouse IL-17 (0.4 μg/ml) and leptin (100 μg/ml) overnight at 4°C. Standards and samples were incubated for 2 hours at room temperature. Then, 50 μl of streptavidin–horseradish peroxidase conjugate was added and incubated for 30 minutes. Freshly prepared 3,3′,5,5′-tetramethylbenzidine was added, and optical density at 405 nm was measured with a Sunrise microplate reader (Tecan). IL-17 levels were measured with a DuoSet ELISA Development System (R&D Systems), and leptin levels were determined with an ELISA Development Kit according to the instructions of the manufacturer (PeproTech).
Paraffin-embedded sections of joint tissue were deparaffinized and rehydrated. Endogenous peroxidase was inhibited by addition of 0.5% hydrogen peroxide for 20 minutes in the dark. Tissue sections were blocked with rabbit serum and incubated with biotinylated rabbit anti-mouse leptin (PeproTech). After incubation for 2 hours, StreptABComplex/HRP (Dako) and 3,3′-diaminobenzidine tetrahydrochloride solution (Sigma-Aldrich) were added to develop brown precipitates. Mayer's hematoxylin was used for nucleus counterstaining. Control staining was performed by omitting the antileptin antibody.
Th17 cells in the knee joint were analyzed by immunofluorescence staining of frozen joint sections from leptin- and PBS-treated mice with CIA. Briefly, knee joint samples were fixed with 4% paraformaldehyde, decalcified with 14% EDTA, and embedded in OCT compound (Sakura) with 30% sucrose. Th17 cells were stained with fluorescein isothiocyanate–conjugated anti–IL-17 (clone TC11-18H10.1) and phycoerythrin-conjugated anti-CD4 antibodies (clone GK1.5) in a dark chamber for 30 minutes at room temperature. Nuclei were counterstained with DAPI (1 μg/ml) for 5 minutes. Leptin receptor was stained with biotinylated goat anti-mouse leptin receptor (R&D Systems). Slides were examined using a Leica Q550CW System.
Paraffin-embedded joint tissue sections (5 μm thick) were stained with hematoxylin and eosin. Histopathologic scoring of joint damage was performed under blinded conditions, according to a widely used scoring system for assessing synovitis, cartilage degradation, and bone erosion (29).
To determine whether leptin exerts any direct effect on Th17 cells, we first detected leptin receptor (Ob-Rb) transcripts in splenic naive CD4+ T cells and in vitro–differentiated Th17 cells from normal mice, by semiquantitative PCR analysis (Figure 1A). In addition, the expression of leptin receptor in CD4+ T cells and Th17 cells was further confirmed by flow cytometry and immunofluorescence microscopy, respectively (Figures 1B and C). When naive CD4+ T cells purified from the spleens of normal mice were cultured for 3 days under polarization conditions for Th17 differentiation, leptin treatment significantly enhanced Th17 cell generation, with a 1.5-fold increase in total cell numbers, an effect that was completely abolished when Ob-R chimera (Ob-R:Fc), a soluble blocker of leptin, was added to the culture of naive T cells (Figures 1D and E). Consistent with the above-described findings, leptin treatment exhibited no apparent effect on Th17 cell differentiation from naive T cells of db/db mice (Figures 1D and E), indicating a direct effect of leptin in enhancing Th17 generation via leptin receptor–mediated signaling transduction. As expected, significantly elevated IL-17 levels were detected in culture supernatants of leptin-treated T cells from normal mice (Figure 1F).
Moreover, increased levels of RORγt transcripts were readily detected when naive T cells were cultured for 15 minutes in the presence of leptin under serum-free Th17 polarization conditions, with a peak level reached by 30 minutes (Figure 2A). Upon treatment with leptin for 30 minutes at concentrations of 50 ng/ml and 250 ng/ml, levels of RORγt transcripts in naive T cells were significantly increased, by 3-fold and 5-fold, respectively, compared with those in T cells cultured in the absence of leptin (Figure 2B).
To verify whether leptin promotes Th17 cell expansion by enhancing the proliferation of Th17 cells, we cultured CFSE-labeled naive CD4+ T cells for 3 days in the presence or absence of recombinant leptin under polarization conditions for Th17 differentiation. Although the frequency of Th17 cells in culture was substantially increased, the proliferation rate of Th17 cells showed no obvious changes upon leptin treatment (Figure 2C). Taken together, these results indicate that leptin may play a role in driving Th17 cell–mediated immune response in vivo.
To examine the levels of leptin expression in the joint tissue during CIA development, we performed immunohistochemical staining on paraffin-embedded joint tissue samples from adjuvant-immunized control DBA/1J mice and mice with CIA at both the acute stage (within 2 weeks after the second CII immunization) and the chronic stage (within 8–10 weeks after the second CII immunization). As shown in Figure 3A, the number of leptin-expressing cells was substantially increased in inflamed synovium of mice with CIA at both the acute and chronic stages, consistent with markedly elevated levels of leptin transcripts in joint tissue as detected by qPCR analysis (Figure 3B). Moreover, levels of leptin in synovial fluid were significantly elevated during CIA development, peaking during the acute stage with a 10-fold increase (Figure 3C). Notably, a similar pattern of elevated IL-17 levels during CIA development was observed in the synovial fluid of mice with CIA (Figure 3D). Further examination of frozen joint sections by immunofluorescence microscopy revealed substantially increased numbers of Th17 cells in the synovial tissue, especially during the acute stage of CIA (Figure 3E). Taken together, these results indicate a positive correlation between local increases in leptin concentrations and Th17 cell expansion during the progression of CIA.
To determine whether locally elevated leptin concentrations can enhance CIA progression, we injected recombinant leptin (5 μg in 10 μl PBS) intraarticularly into the knee joint of DBA/1J mice once every 3 days for a total of 3 injections, starting on day 17 after the first CII immunization (Figure 4A). We found that the onset of arthritis occurred 1 week earlier in leptin-treated mice compared with PBS-treated control mice (Figure 4B). In addition, arthritis symptoms were markedly more severe in the leptin-treated mice with CIA (Figures 4C and D). Interestingly, draining popliteal lymph nodes in leptin-treated mice with CIA were enlarged and exhibited significantly increased cellularity (Figure 4E). Moreover, histopathologic examination of joint samples revealed more pronounced synovial hyperplasia, cartilage damage, and bone erosion in leptin-treated mice with CIA compared with PBS-treated CIA controls (Figure 4F). Taken together, these results indicate that locally increased leptin concentrations contribute directly to the exacerbation of synovial inflammation and pathologic changes in the joints of mice with CIA.
To evaluate the effect of local leptin injection on T cell responses in vivo, we examined the frequencies of Th1 and Th17 cells in peripheral lymphoid organs in PBS- and leptin-treated mice with CIA. Both the percentages and the total numbers of IFNγ-producing CD4+ Th1 cells were moderately increased in the spleen, draining lymph nodes, and joint tissue of leptin-treated mice with CIA, but the increases were not significant compared with findings in PBS-treated controls with CIA (Figures 5A and B).
In contrast, IL-17–producing CD4+ Th17 cells were significantly expanded (∼3-fold) in both draining lymph nodes and joint tissue of mice with CIA (Figures 5C and D), although increases in the spleen were mild and not significant. In addition, levels of IL-17 transcripts were increased by ∼4-fold in joint tissue of leptin-treated mice with CIA, consistent with elevated IL-17 levels in synovial fluid as detected by ELISA (Figures 6A and B). Notably, the number of IL-17–secreting CD4+ Th17 cells showed a marked 6-fold increase in cell suspensions of joint tissue samples from leptin-treated mice with CIA, as determined by ELISpot assay (Figure 6C). Further examination of frozen joint sections by immunofluorescence microscopy confirmed the substantially increased numbers of CD4+ Th17 cells in the synovial tissue of leptin-treated mice with CIA when compared with PBS-treated controls with CIA (Figure 6D).
In this study we have shown for the first time that leptin can promote Th17 cell differentiation from naive CD4+ T cells in culture. Moreover, local intraarticular injection of leptin markedly enhances synovial hyperplasia with increased numbers of Th17 cells present in the joint tissue and results in exacerbated cartilage damage and bone erosion in mice with CIA. Thus, our current findings indicate a previously unrecognized role of leptin in enhancing Th17 response and driving autoimmune inflammation during CIA development.
There is compelling evidence that leptin is involved in modulating immune responses in addition to its well-recognized functions in controlling food intake and energy homeostasis. Extensive investigations, including previous studies by our group, have characterized an important role of leptin in regulating the maturation and functions of macrophages, dendritic cells, natural killer cells, and B cells (30). In particular, several studies have clearly demonstrated the crucial function of leptin in suppressing T cell apoptosis and promoting T cell activation via STAT-3–mediated signaling transduction. Recent findings have further revealed a critical role of leptin in the control of Treg cell proliferation, which highlights the diversified functions of leptin in modulating the complex relationships between immunity and inflammatory response (16). Interestingly, a link between leptin and IL-17 in obesity and cancer has been established (31, 32).
In accordance with previous findings of leptin receptor expression on CD4+ T cells and Treg cells (14), we detected the expression of leptin receptor on Th17 cells. In addition, we provide new evidence that leptin can effectively promote Th17 cell expansion in culture. This action is mediated through leptin receptor, since leptin treatment exerted no obvious effect on Th17 generation from cultured naive T cells from db/db mice. Notably, the observed Th17 cell expansion is mainly due to enhanced Th17 differentiation from naive T cells, since leptin does not appear to have an effect on promotion of Th17 cell proliferation in culture. Together with the observation of up-regulated RORγt transcripts in leptin-treated naive T cells, these findings demonstrate a previously unrecognized function of leptin in enhancing Th17 cell differentiation. Whether leptin exerts any effect on the generation of and cytokine production by memory T cells is currently unknown, and future studies are needed to obtain further insight into the extensive functions of leptin in regulating T cell maturation and function.
As a key effector cell subset of CD4+ T cells, Th17 cells are known to play a potent proinflammatory role in the pathogenesis of autoimmune disease (33), and several recent investigations have suggested the critical involvement of Th17 cells in the progression of autoimmune arthritis (34–36). In this study we have demonstrated that locally increased leptin concentrations via intraarticular injection in the knee joint trigger earlier onset of arthritis and increased severity of clinical symptoms. Moreover, local leptin administration exacerbates synovial inflammation, with increased numbers of Th17 cells in both draining lymph nodes and joint tissue.
It is currently unclear whether the increased numbers of Th17 cells in joint and synovial tissue result from increased de novo Th17 cell differentiation in situ or from enhanced migration of Th17 cells into local inflamed joint tissue. However, we have found significantly increased levels of IL-6 in both joint tissue and synovial fluid after local leptin injection in mice with CIA (Deng J, et al: unpublished observations), indicating that the microenvironment in the inflamed joint may favor Th17 cell differentiation in situ. Interestingly, blockade of IL-6 using anti–IL-6 receptor monoclonal antibody has been shown to suppress autoimmune arthritis and experimental autoimmune encephalomyelitis in mice, by inhibiting inflammatory Th17 responses (37, 38). Thus, it is plausible to reason that overproduction of leptin and IL-6 may collectively drive Th17 response and promote progression of autoimmune disease (39).
Busso et al have reported a significantly reduced Th1 response with increased production of the Th2 cytokine IL-10 in both db/db and ob/ob mice, indicating a role of leptin in promoting the Th1 response (24). In the present study, local administration of leptin was shown to trigger a markedly enhanced Th17 response with modestly increased numbers of IFNγ-producing Th1 cells in situ, which is concordant with the finding that Th17 cells act as a crucial inducer of RANKL to stimulate osteoclastogenesis and promote bone erosion in mice with CIA (40). Thus, our observations of markedly increased leptin levels in inflamed joint tissue during CIA development suggest a pathogenic role of leptin in driving synovial inflammation and joint damage in situ. Since synoviocytes are also known to express leptin receptor, it remains to be investigated whether and how increased leptin levels in local joint tissue affect synoviocyte functions and contribute to synovial hyperplasia during arthritis progression. Moreover, further studies are needed to examine the function of leptin in influencing IL-17 response and joint inflammation during the chronic stage of CIA.
There have been conflicting reports on the relationships between leptin levels and disease activity in RA (41). Although serum leptin concentrations have been reported to be significantly increased in patients with erosive RA (21), in other studies higher levels of serum leptin were associated with reduced joint damage (22). Nevertheless, these contrasting results may suggest that local leptin levels in joint tissue could be directly correlated with synovial inflammation and joint damage in RA. In a recent study by Tan et al, high adiponectin expression was detected in synovial fluid and synovial tissue of RA patients (42), while adiponectin stimulates synovial fibroblasts to produce monocyte chemoattractant protein 1 and IL-6 (19), which further indicates that locally produced adipokines are closely implicated in synovial inflammation during the pathogenesis of RA.
In summary, our results have identified a previously undescribed function of leptin in modulating Th17 response and exacerbating autoimmune arthritis. Further studies are needed to determine whether local leptin gene silencing or targeting of leptin signaling may exert any therapeutic effect in CIA, which may have important implications for the development of new strategies for the treatment of patients with RA and other autoimmune diseases.
All authors were involved in drafting the article or revising it critically for important intellectual content, and all authors approved the final version to be published. Dr. Lu had full access to all of the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.
Study design. Deng, Liu, Yang, Lu.
Acquisition of data. Deng, Liu, Yang, Ko.
Analysis and interpretation of data. Deng, Liu, Yang, S. Wang, Zhang, X. Wang, Ko, Hua, Sun, Cao, Lu.
We thank the staff of the Medical Faculty Core Facility, University of Hong Kong for technical support.