Proinflammatory Th17 cells are expanded and induced by dendritic cells in spondylarthritis-prone HLA–B27–transgenic rats




HLA–B27/human β2-microglobulin–transgenic (B27-transgenic) rats, a model of spondylarthritis (SpA), develop spontaneous colitis and arthritis under conventional conditions. CD4+ T cells are pivotal in the development of inflammation in B27-transgenic rats. This study was undertaken to characterize the phenotype of CD4+ T cells in this model and to determine whether dendritic cells (DCs) induce proinflammatory T cells.


The phenotype of CD4+ T cells from rat lymph nodes (LNs) draining the sites of inflammation was analyzed by flow cytometry. Immunostaining was used to detect interleukin-17 (IL-17)–producing cells in the rat joints. DCs from B27-transgenic or control rats (transgenic for HLA–B7 or nontransgenic) were cocultured with control CD4+ T cells and stimulated with anti–T cell receptor α/β.


IL-17A– and tumor necrosis factor α (TNFα)–producing CD4+ T cells were expanded in mesenteric and popliteal LNs from B27-transgenic rats. The accumulation of Th17 cells correlated with disease development, in contrast to Th1 or Treg cells. IL-17–positive mononuclear cells were detected in the arthritic joints of B27-transgenic rats but not in the joints of control rats. Finally, in vitro cocultures demonstrated that Th17 cells were preferentially induced and expanded by DCs from B27-transgenic rats, by a process that may involve defective engagement of costimulatory molecules.


Our findings indicate that expanded CD4+ T cells in B27-transgenic rats exhibit a proinflammatory Th17 phenotype characterized by IL-17A and TNFα production. Furthermore, this population is preferentially induced by DCs from B27-transgenic rats. These data point toward an induction of Th17 cells as a possible pathogenic mechanism in this model of SpA. However, their pathogenic role still needs to be shown.

Spondylarthritis (SpA) is an inflammatory rheumatic disorder characterized by axial and peripheral enthesitis and/or arthritis and frequent extraarticular manifestations, such as uveitis, psoriasis, and inflammatory bowel diseases (Crohn's disease or ulcerative colitis). All skeletal and extraarticular features of SpA are thought to be determined by shared genetic factors, dominated by the class I major histocompatibility complex (MHC) allele HLA–B27 (1). Although the strong association between HLA–B27 and SpA was first described more than 35 years ago, the mechanism of this association has remained unexplained until now (2).

Evidence that HLA–B27 plays a direct role in determining SpA pathogenesis came from the description of an HLA–B27–transgenic rat model. A multisystem inflammatory disorder mimicking the most striking aspects of SpA (rat SpA), including ankylosing spondylitis, peripheral arthritis, psoriatic lesions, and ulcerative colitis, arises spontaneously in several lines of rats transgenic for HLA–B*2705 and human β2-microglobulin (hβ2m), but not in the control HLA–B*0702/hβ2m–transgenic line (3, 4). The expression of rat SpA is determined by several factors, including high levels of both HLA–B27 and hβ2m transgenes, a specific rat genetic background, and the presence of normal microbial flora (5, 6). Importantly, the role of the immune system in this model was highlighted in several ways. First, rat SpA was induced in nontransgenic recipients by transferring immature hematopoietic cells, but not mature lymph node (LN) cells, from disease-prone rats, indicating that a hematopoietic-derived cell is critical for disease induction (7). Second, nude HLA–B27–transgenic rats, which lack thymically derived T cells, were protected against disease but promptly developed rat SpA upon reconstitution with CD4+ T cells or thymus engraftment, whereas reconstitution with CD8+ T cells was relatively inefficient (6, 8). Moreover, experiments involving CD8+ T cell depletion confirmed that CD8+ T cells were not required for the development of rat SpA (9, 10).

Based on the results described above, we postulated that rat SpA could arise as a consequence of an interaction between antigen-presenting cells (APCs) expressing high levels of HLA–B27 and CD4+ T cells, and carried out experiments to examine the consequences of such interaction. Interestingly, these experiments revealed several aberrant functions in APCs from HLA–B27/hβ2m–transgenic rat lines. Most notably, mature splenic and LN dendritic cells (DCs) exhibited a decreased capacity to stimulate an allogeneic or a syngeneic T cell response, which paralleled disease susceptibility in a broad variety of lines, and was not a consequence of the inflammatory disease (11–13). This altered function could be linked to a defective engagement of costimulatory molecules, such as CD86, and also to a decreased capacity to form an antigen-independent immunologic synapse with CD4+ T cells (12, 14). Other aberrant functions in HLA–B27–transgenic rat DCs included altered cytoskeletal dynamics, decreased class II MHC expression, and enhanced apoptotic susceptibility, all potentially contributing to their decreased stimulatory function (15).

DCs are professional APCs that are capable of activating naive CD4+ T cells to differentiate and to proliferate. However, besides promoting T cell responses to antigen, evidence indicates that DCs play a role in establishing tolerance toward self antigens and in the maintenance of peripheral tolerance (16). Accordingly, spontaneous inflammation in this model could result from a breakdown of peripheral tolerance.

Upon stimulation by DCs, naive CD4+ T cells are known to differentiate into distinct lineages of effector cells. While interleukin-12 (IL-12) drives the development of Th1 cells, producing predominantly interferon-γ (IFNγ) and IL-2 and eliciting cell-mediated immunity against intracellular pathogens (17), Th2 cells differentiate in response to IL-4; produce IL-4, IL-5, and IL-13; and are involved in the humoral response against parasites and allergy (17, 18). More recently, Th17 cells, a subset of T cells producing IL-17A, IL-17F, and IL-22, have been identified (19–21) and shown to be critical for the induction of several autoimmune disease models, such as collagen-induced arthritis and experimental autoimmune encephalomyelitis (22, 23). Th17 cells differentiate in mice in response to transforming growth factor β (TGFβ) and IL-6 or IL-21 (24) and differentiate in humans in response to a mixture of cytokines, including IL-1β, IL-6, tumor necrosis factor α (TNFα), IL-23, and TGFβ (25, 26). Finally, a CD4+ Treg cell population expressing the FoxP3 transcription factor has also been described (27). This Treg cell population appears to inhibit the proliferation of all of the former effector Th cells (28).

In the present study, we examined the phenotype of CD4+ T cells in a disease-prone HLA–B27–transgenic rat line and the putative influence of DCs from these rats on the differentiation of CD4+ T cells. First, we observed that CD4+ T cells with a proinflammatory Th17 phenotype accumulate in B27-transgenic rats, in parallel with the development of rat SpA. Furthermore, we showed that Th17 cells were preferentially induced and expanded by DCs from B27-transgenic rats, by a contact-dependent mechanism that may involve a previously described defective engagement of costimulatory molecules (12, 14).



The HLA–B27–transgenic rat line 33-3 and the HLA–B7–transgenic rat line 120-4 were originally produced at the University of Texas Southwestern Medical Center (Dallas, TX). Disease-prone rats of the 33-3 line, bearing 55 copies of HLA–B*2705 and 29 copies of hβ2m, and disease-free homozygous rats of the 120-4 line, bearing 52 copies of HLA–B*0702 and 31 copies of hβ2m, both on a Fischer background, were bred and maintained under conventional conditions (12, 14). Nontransgenic littermates from the 33-3 rat line were used as controls. Age- and sex-matched rats (1–12 months of age) were used in each experiment. Study procedures were approved by the institutional animal care committee.

Cell culture medium, monoclonal antibodies (mAb), and other reagents.

Cell cultures were performed in RPMI 1640 medium with Glutamax I (Life Technologies) supplemented with 10% fetal calf serum, streptomycin (100 μg/ml), 2% sodium pyruvate, 0.05 mM 2-mercaptoethanol, and 5 mM HEPES (complete medium), unless otherwise stated. The following mouse IgG1 or IgG2a anti-rat antigen mAb were obtained from European Collection of Cell Cultures or purchased from BD PharMingen: R73 (T cell receptor α/β [TCRα/β]), OX35 (CD4), OX8 (CD8α chain), 3.2.3 (natural killer cell receptor protein 1A), OX62 (αE2 integrin present on CD103+ DCs), OX33 (CD45 epitope specific to B cells and a subset of DCs), OX12 (Igκ chain), OX42 (C3bR [macrophages]), OX39 (CD25), 24F (CD86), JJ316 (CD28), and FJK-16s (FoxP3). Fluorescein isothiocyanate (FITC)–conjugated goat anti-mouse (GAM) IgG2a was from Caltag, and Cy5-conjugated GAM IgG1 was from Jackson ImmunoResearch. Allophycocyanin-conjugated anti-rat CD4, Cy7-conjugated anti-rat TNFα, phycoerythrin-conjugated anti-mouse IL-17A, and FITC-conjugated anti-rat IFNγ were from BD PharMingen. Rabbit anti-human IL-17A (sc-7927), goat anti-rat IL-23 p19 (sc-21083), and goat antihuman TGFβ1 (sc-31608) were from Santa Cruz Biotechnology. Peroxidase-conjugated goat anti-rabbit IgG was from Jackson ImmunoResearch. Recombinant rat IFNγ was from ImmunoTools. Phorbol 12-myristate 13-acetate (PMA), 5,6-carboxyfluorescein diacetate N-succinimidyl ester (CFSE), ionomycin, and brefeldin A were from Sigma.

Cell preparation.

Splenic DCs were obtained by two different methods, as previously described (12, 14). The first method was adapted from that described by Knight et al (29). Briefly, a single-cell suspension prepared from spleen was cultured overnight in RPMI Medium 1640 (Dutch Modification) (Life Technologies). Recovered nonadherent cells were subjected to a 14.5% (weight/volume) metrizamide gradient. Low-density cells were collected at the interface and consisted mostly of DCs.

The second method was derived from the technique described by Josien et al (30), as follows. A single-cell suspension was prepared from spleen, minced, and digested in 2 mg/ml collagenase D (Roche Diagnostics). Low-density cells were then collected after centrifugation on a 14.5% Nycodenz gradient (Nycomed). Cells were then incubated with OX62 mAb–coated microbeads (Miltenyi Biotec) and positively selected on MS-type selection columns (Miltenyi Biotec). This population of OX62+ DCs was used after overnight incubation in complete medium containing 4% culture supernatant from murine hybridoma transfected with rat granulocyte–macrophage colony-stimulating factor.

T cells from HLA–B7–transgenic or nontransgenic control rats were isolated from LN single-cell suspensions by magnetic-activated cell sorting. Negative selection was performed, using combinations of OX33, OX42, 3.2.3, OX8, and OX39 to purify naive CD4+CD25− T cells. The purity of the selected populations was routinely in the range of 95–98% as determined by fluorescence-activated cell sorting (FACS) analysis.

Cell cultures.

Splenic DC populations were cocultured with CD4+CD25− T cells labeled with 5 μM CFSE with anti-TCRα/β (R73) mAb for 4 days in 24-well flat-bottomed plates in 500 μl complete medium. The DC:T cell ratio was 1:1. Proliferation of T cells was assessed by FACS, by evaluating the dilution of CFSE in TCRα/β+ cells.

In experiments using a Transwell apparatus (Corning Costar), CD4+CD25− T cells (5 × 105/well) were stimulated with DCs (5 × 105/well) in lower compartments of 24-well culture dishes, in a volume of 250 μl, in the presence of a Transwell insert (6.5-mm diameter, 0.4-μm pore size) containing either DCs and T cells (2 × 105 of each cell type per insert) or DCs alone, in 250 μl of medium. After 4 days of culture, cells from the lower compartments were separately transferred to 96-well plates before intracellular staining. In blocking experiments, anti-CD86 and anti-CD28 mAb were used at 5-μg/ml saturating concentrations.

Determination of cytokine levels.

The cytokines IL-4, IL-10, IL-17A, TNFα, and IFNγ were measured in rat sera and culture supernatants by specific enzyme-linked immunosorbent assays, according to the recommendations of the manufacturer (eBioscience). Results are expressed as the mean ± SEM.

Immunohistochemical analysis.

Paraffin-embedded sections of paws were dewaxed with Histo-Clear (twice for 5 minutes each time) and methanol (twice for 5 minutes each time) and subsequently added to distilled water. Sections were quenched with 3% H2O2 in Milli-Q water (twice for 5 minutes each time) to block endogenous peroxidase activity. After washing 3 times in phosphate buffered saline (PBS)–0.1% Tween, sections were incubated with goat serum diluted 1:5 (in PBS–0.1% Tween) for 30 minutes at room temperature to block nonspecific binding. Sections were subsequently incubated overnight at 4°C with 5 μg/ml rabbit anti-human IL-17A antibody or 5 μg/ml normal rabbit IgG as a negative control. In additional experiments, the primary antibody was first incubated for 2 hours with 250 ng murine recombinant IL-17A (eBioscience). After washing 3 times, a second blocking step with goat serum diluted 1:5 was applied. Sections were subsequently incubated with peroxidase-conjugated goat anti-rabbit IgG diluted 1:300 in PBS–0.1% Tween for 30 minutes at room temperature. After washing, enzyme activity was detected with diaminobenzidine chromogen, and sections were counterstained with hematoxylin.

Real-time polymerase chain reaction (PCR).

Total RNA was isolated from LN CD4+ T cell pellets using an RNeasy Mini kit (Qiagen). Genomic DNA was removed using the RNase-Free DNase set for DNA digestion during RNA purification (Qiagen). First-strand complementary DNA synthesis was performed using a Sensiscript RT kit with random hexamer primers (Qiagen). Levels of messenger RNA (mRNA) for the tested genes were quantified by real-time PCR (ABI 7900HT Sequence Detection System; Applied Biosystems) using SYBR Green Master Mix (Applied Biosystems). Thermocycling included incubation at 95°C for 10 minutes, followed by a 2-step PCR program of 95°C for 15 seconds and 55°C for 60 seconds and extension at 72°C for 10 seconds, for a total of 45 cycles. The total amount of mRNA was normalized across samples according to endogenous Hprt1 mRNA. The primer sequences (forward and reverse, respectively) were as follows: for Hprt1, 5′-TTTGTGTCATCAGCGAAAGTG-3′ and 5′-ATGGCCACAGGACTAGAAC-3′; for IL-21, 5′-AAGGCCAGATCACCTTCTGA-3′ and 5′-GCCCCTTTACATCTTGTGGA-3′; for IL-22, 5′-ACCAAGCTCAGCAGTCACCT-3′ and 5′-AGTTCCCCGATCGCTTTAAT-3′; for Tbx21, 5′-AGCCCACTGGATGCGACAGGA-3′ and 5′-GCGGCTGGTACTTATGGA-3′; for IFNγ, 5′-GGCCATCAGCAACAACATAAG-3′ and 5′-TGGGTTGTTCACCTCGAACTT-3′; and for Rorc, 5′-GGATGAGATTGCCCTCTACAC-3′ and 5′-GGAGGCCTTGTCGATGAGTC-3′.


For FACS analysis of surface antigen, cells were incubated with saturating concentrations of the appropriate primary mAb (mouse IgG1 or IgG2a isotype) for 30 minutes, washed, and then incubated with secondary Cy5-conjugated GAM IgG1 or FITC-conjugated IgG2a, as needed, for 30 minutes. Cytokine-producing cells were determined by surface staining using allophycocyanin-conjugated anti-rat CD4 and intracellular staining using fluorescent anticytokine mAb. Briefly, cells were stimulated with PMA and ionomycin (both at 0.5 μg/ml) in the presence of brefeldin A (10 μg/ml) for 4 hours. Cells were fixed in 2% paraformaldehyde, permeabilized with 0.1% saponin, and stained with fluorescent mAb. Then, the cells were analyzed using an FC500 cytometer (Beckman Coulter) and CXP analysis software (Beckman Coulter).

Statistical analysis.

Statistical comparisons between groups were performed using Student's t-tests. For analysis of real-time reverse transcriptase–PCR experiments, the ratios of the gene transcript levels in the B27-transgenic rats to those in the nontransgenic control rats were compared to the theoretical value of 1.0 (i.e., no difference) by 1-sample t-test. P values less than 0.05 were considered significant.


Increased CD4+ T cell numbers in HLA–B27–transgenic rat LNs.

To investigate the role of CD4+ T cells in rat SpA, we enumerated CD4+ T cells in mesenteric LNs (MLNs) and popliteal LNs from B27-transgenic rats and control rats (either nontransgenic or transgenic for HLA–B7). There was a notable increase in the number of CD4+ T cells obtained from the MLNs of HLA–B27–transgenic rats compared to their nontransgenic littermates; this difference was statistically significant at any time point (Figure 1A). Similar statistically significant differences were also observed in popliteal LNs, but to a lesser extent (data are available from the author upon request). There was no difference in the number of CD4+ T cells between HLA–B7–transgenic rats and nontransgenic rats, confirming the specificity of their expansion to disease-prone B27-transgenic rats and not to any HLA–B–transgenic rat line (data not shown).

Figure 1.

Increased numbers of CD4+ and CD4+CD25+ T cells in the lymph nodes (LNs) of HLA–B27–transgenic (B27) rats. Cell suspensions were prepared from the mesenteric LNs (MLNs) and peripheral LNs (PLNs) of age-matched control rats (nontransgenic [NTG] rats or B7-transgenic rats) and B27-transgenic rats. Cells were labeled with anti-CD4, anti-CD25, and anti-FoxP3 monoclonal antibodies to discriminate between activated T cells and Treg cells by fluorescence-activated cell sorting. A, Absolute numbers of CD4+ T cells in the MLNs of nontransgenic rats (shaded bars) and B27-transgenic rats (solid bars) at the indicated ages. Bars show the mean ± SEM (n = 5–7 rats per group). There was a significant difference between the nontransgenic group and the B27-transgenic group at each time point. B, Absolute numbers of CD4+CD25+ T cells in the peripheral LNs and MLNs of 6-month-old nontransgenic and B7-transgenic control rats (shaded bars) and B27-transgenic rats (solid bars). Bars show the mean ± SEM (n = 6 rats per group). C, Dot-blots showing the expression of CD25 and FoxP3 in CD4+-gated T cells from the MLNs of 6-month-old nontransgenic and B27-transgenic rats. D, Ratios of FoxP3− cells (activated cells) to FoxP3+ cells (Treg cells) among CD4+CD25+-gated cells in the peripheral LNs and MLNs of 6-month-old nontransgenic and B7-transgenic control rats (shaded bars) and B27-transgenic rats (solid bars). Bars show the mean ± SEM (n = 6 rats per group). ∗ = P < 0.05; ∗∗ = P < 0.005.

Higher numbers of activated CD4+ T cells than Treg cells in HLA–B27–transgenic rats.

Similar to the whole CD4+ T cell population, the subset of CD4+CD25+ T cells was significantly expanded in the MLNs and peripheral LNs of HLA–B27–transgenic rats (Figure 1B). Both activated CD4+ T cells and Treg cells may express CD25, but they can be distinguished on the basis of FoxP3 expression, which is restricted to the latter population. Both CD25+FoxP3+ (Treg) and CD25+FoxP3− (activated) CD4+ T cells were increased in the LNs of B27-transgenic rats (Figure 1C). However, this increase was more pronounced among activated cells than Treg cells (Figures 1C and D).

Preferential increase in Th17 cell number in parallel with disease development in the B27-transgenic rat.

To investigate the phenotype of expanded CD4+ T cells, we performed intracellular staining and monitored the proportions of MLN CD4+ T cells expressing IFNγ, IL-17A, and TNFα, alone or in combination, in rats ages 1 month (prior to rat SpA development) up to 1 year (Figure 2A). Th1 cells producing IFNγ alone predominated in control rats, and the number of these cells increased with age in both B27-transgenic rats and controls, without any differences between the 2 groups (Figure 2A). The proportion of Th17 cells producing IL-17A remained low (<1%) across all ages in controls (Figure 2A). In contrast, in the MLNs of B27-transgenic rats, the proportion of IL-17A+CD4+ cells, which was as low as that in controls at 1 month, rose steadily thereafter, reaching 20% at 1 year. This pattern was observed for the major population of IL-17A+IFNγ− cells as well as for a minor subset of IL-17A+IFNγ+ cells (Figure 2A).

Figure 2.

CD4+ T cells are biased toward a proinflammatory Th17 phenotype in the B27-transgenic rat. In vitro–stimulated MLN (A) and popliteal LN (B) cells from age-matched nontransgenic rats, B7-transgenic rats, and B27-transgenic rats at the indicated ages (n = 5 rats per group) were first surface-labeled with anti-CD4 and then stained intracellularly with fluorescent anti–interferon-γ (anti-IFNγ), anti–tumor necrosis factor α (anti-TNFα), and anti–interleukin-17A (anti–IL-17A) and analyzed by fluorescence-activated cell sorting. Results are the percentages of CD4+-gated cells and represent the evolution of Th1 cells (IFNγ+IL-17−), Th17 cells (IFNγ−IL-17+), Th1/17 cells (IFNγ+IL-17+), and TNFα-producing CD4 T cells over time. Symbols represent individual rats; horizontal lines show the mean ± SEM. ∗∗ = P < 0.005; ∗∗∗ = P < 0.001, B27-transgenic rats versus pooled controls (B7-transgenic and nontransgenic rats). See Figure 1 for other definitions.

In addition, we quantified CD4+ T cells producing TNFα, a cytokine that can be produced by either Th1 or Th17 cells. The proportion of TNFα+CD4+ T cells was significantly increased in the MLNs of B27-transgenic rats (Figure 2A). Interestingly, this TNFα+CD4+ T cell population contained a majority of the IL-17A+ cells in B27-transgenic rat LNs, and a minor subset of the TNFα+IL-17A+ cells were also IFNγ+ (data are available from the author upon request). A similar pattern, consisting of an increased proportion of IL-17A+ and TNFα+ CD4+ T cells, but a normal proportion of IFNγ+IL-17A− CD4+ T cells, was observed in the popliteal LNs of B27-transgenic rats (Figure 2B). Notably, this increase appeared later in the popliteal LNs than in the MLNs (i.e., at 6 months versus 3 months), parallel to the development of arthritis, which follows that of colitis in this model.

The preferential increase in the number of Th17 cells was further evaluated by transcription analysis of MLN CD4+ T cells. Using real-time PCR, we quantified the expression of IL-21, IL-22, and IFNγ mRNA. IL-21 and IL-22 are typically produced by Th17 cells, and IFNγ is a Th1-specific cytokine. We also quantified the expression of retinoic acid receptor–related orphan nuclear receptor (coded by Rorc) and T-bet (coded by Tbx21) as typical Th17 and Th1 transcription factors, respectively. The 5 transcripts were detected in all CD4+ T cell samples purified from the MLNs of 5 nontransgenic and 8 diseased B27-transgenic rats ages 6–9 months. The transcription levels of IL-21, IL-22, and Rorc, the combination of which is characteristic of a Th17 phenotype, were significantly increased in the B27-transgenic rats relative to the nontransgenic controls, with mean ratios of 1.8 (95% confidence interval [95% CI] 1.2–2.4) (P = 0.02), 32 (95% CI 14.5–49.5) (P = 0.004), and 8.8 (95% CI 3.2–14.3) (P = 0.013), respectively. In contrast, the expression levels of IFNγ and Tbx21, which are representative of a Th1 phenotype, did not differ between B27-transgenic and nontransgenic rats (mean ratio 1.0 [95% CI 0.6–1.5] [P = 0.92] and 1.4 [95% CI 0.5–2.3] [P = 0.4], respectively) (data are available from the author upon request).

Consistent with the results described above, IL-17A was detected at significantly higher levels in the sera of 6-month-old B27-transgenic rats (mean ± SEM 32.2 ± 4.7 pg/ml; n = 6), than in B7-transgenic controls (2.85 ± 1.6 pg/ml; n = 6) (P < 0.005) and nontransgenic controls (0 ± 0 pg/ml; n = 6) (P < 0.001). These levels were higher than the levels of IFNγ and TNFα in the B27-transgenic rats, which were not significantly different from those observed in the B7-transgenic and nontransgenic controls (data are available from the author upon request). Neither IL-4 nor IL-10 was detected in any of the sera (data not shown).

Finally, IL-17A–positive mononuclear cells were detected in the hyperplastic lining layer and in synovial infiltrates of the arthritic paws of 7-month-old HLA–B27–transgenic rats (Figures 3A, B, and D). In contrast, the paucicellular synovium of the nontransgenic controls (Figures 3C and D) yielded negative signals, as did that of the nonarthritic HLA–B7–transgenic controls (Figures 3C and D).

Figure 3.

Presence of interleukin-17 (IL-17)–positive mononuclear cells in the arthritic joints of HLA–B27–transgenic rats. Immunohistochemical analysis for IL-17A was performed in paws from 7-month-old rats. A and B, IL-17–positive cells in the synovium of an arthritic female HLA–B27–transgenic rat. Negative controls showing preincubation of the antibody with recombinant IL-17 or control IgG substitution of the primary antibody are also shown. C, Near absence of IL-17–positive mononuclear cells in the paucicellular normal synovium from an HLA–B7–transgenic (HLAB7tg) rat and a nontransgenic (wild-type) control rat. Original magnification × 400. D, IL-17–positive cells in nontransgenic, HLA–B7–transgenic, and HLA–B27–transgenic rats. Six randomly chosen zones of the synovium from 2 rats in each group were scored in a blinded manner. Bars show the mean ± SEM. P = 0.0006, HLA–B27–transgenic rats versus nontransgenic rats; P = 0.0005, HLA–B27–transgenic rats versus HLA–B7–transgenic rats.

DCs from HLA–B27–transgenic rats favor Th17 cell induction.

We next addressed whether DCs in B27-transgenic rats, which display aberrant function (14), contributed to the notable increase in the number of Th17 cells. To test this possibility, we cocultured splenic DCs with CFSE-labeled syngeneic CD4+CD25− T cells from control rats (either B7-transgenic or nontransgenic) and stimulated the coculture with anti-TCRα/β. In these assays, we used 2 different splenic DC populations: CD103+ DCs or DCs isolated by the method described by Knight et al (29) (Figure 4). Both types of DCs showed the expansion and/or differentiation of IL-17A–producing T cells, which were detected among the CD4+ T cells that divided the most (Figure 4A). This effect was 3- to 15-fold greater with B27-transgenic rat DCs than with control DCs (Figures 4A and B). In this coculture assay, both control and B27-transgenic rat DCs also induced the production of IFNγ and TNFα by control T cells (Figure 4C). Most interestingly, the proportion of T cells producing those cytokines was considerably greater after coculture with B27-transgenic rat DCs (Figure 4C). Furthermore, as shown in the MLNs of B27-transgenic rats, the majority of IL-17A–producing T cells induced by B27-transgenic rat DCs also produced TNFα (Figure 4D). Thus, B27-transgenic rat DCs directly participate in the induction of proinflammatory cytokine–producing T cells.

Figure 4.

Splenic dendritic cells (DCs) from HLA–B27–transgenic (B27) rats induce a biased expansion of Th17 cells. Mature nontransgenic (NTG) or B27-transgenic rat CD103+ DCs or DCs isolated by a method adapted from Knight et al (29) (Knight DCs) were cocultured with 5,6-carboxyfluorescein diacetate N-succinimidyl ester (CFSE)–labeled lymph node CD4+CD25− T cells from nontransgenic or B7-transgenic control rats with anti–T cell receptor α/β (anti-TCRα/β) (R73 monoclonal antibody; 1 μg/ml) stimulation. After 4 days, cells were surface-labeled with anti-TCRα/β, then stained intracellularly with anti–IL-17A and analyzed by fluorescence-activated cell sorting (FACS). Results are shown for TCRα/β+-gated cells. A, Dot-blots showing results with Knight DCs and with CD103+ DCs. Results are representative of 10 independent experiments using T cells from nontransgenic or B7-transgenic control rats. B, IL-17A+CSFEintermediate/low T cells resulting from coculture with Knight DCs and CD103+ DCs from nontransgenic and B27-transgenic rats. Bars show the mean ± SEM (n = 10 experiments with Knight DCs and 7 experiments with CD103+ DCs). ∗∗∗ = P < 0.001. C, Dot-blots showing results after DCs isolated by the method adapted from Knight et al were cocultured as described above with T cells from nontransgenic control rats. After 4 days, cells were surface-labeled with anti-TCRα/β and then stained intracellularly with anti-IFNγ, anti–IL-17A, and anti-TNFα and analyzed by FACS. Results are representative of 3 independent experiments. D, Dot-blots showing the expression of TNFα and IL-17A in TCRα/β+-gated cells (same experiment as described in C). See Figure 2 for other definitions.

Th17 induction is contact dependent.

B27-transgenic rat DCs may favor the expansion of Th17 cells by an increased secretion of stimulatory factors or a decreased capacity to produce inhibitory factors. To investigate whether soluble factors such as cytokines were involved in this process, we used a Transwell assay (Figure 5). The proportion of Th17 cells induced upon contact with control DCs in the lower well was not modified by the presence of B27-transgenic rat DCs in the upper well, whether alone or in coculture with control T cells (0.1% of proliferative Th17 cells, with or without B27-transgenic rat DCs in the upper compartment) (Figure 5). Nor did the presence of nontransgenic DCs in the upper well, alone or in coculture with control T cells, influence the induction of Th17 cells upon contact with B27-transgenic rat DCs in the lower well (0.4–0.5% of proliferative Th17 cells, with or without nontransgenic DCs in the upper compartment) (Figure 5). Moreover, neither the addition of anti–IL-23 or anti-TGFβ neutralizing antibodies to the coculture to block major soluble factors known to stabilize or to induce Th17 phenotype, nor the addition of IFNγ, which may oppose Th17 cell differentiation, influenced the observed pattern (results not shown).

Figure 5.

Th17 cell induction by B27-transgenic rat DCs is contact dependent. Mature nontransgenic or B27-transgenic rat DCs isolated by a method adapted from Knight et al (29) were cocultured with CFSE-labeled lymph node CD4+CD25− T cells from a nontransgenic control rat with anti-TCRα/β (R73 monoclonal antibody; 1 μg/ml) stimulation, in the lower compartment of a Transwell apparatus. The upper compartment contained DCs alone or DCs and T cells, as indicated below each panel. After 4 days, cells in the lower compartment were harvested, labeled, and analyzed by FACS as described in the Figure 4 legend. Dot-blots are shown for TCRα/β+-gated cells. Data are representative of 3 independent experiments. IL-17A = interleukin-17A (see Figure 4 for other definitions).

Correlation of Th17 cell induction with impaired engagement of costimulatory molecules.

We have previously shown that DCs from B27-transgenic rats have an impaired capacity to form a mature antigen-independent immunologic synapse with CD4+ T cells, which involves a defective engagement of costimulatory molecules such as CD86 (12). Thus, in the present study we examined whether such abnormal function could be involved in the expansion of Th17 cells. We performed cocultures in the presence of either a neutralizing anti-CD86 mAb or an anti-CD28 “superagonist,” mAb JJ316, which stimulates T cell proliferation independently of DC–T cell contact (31) (Figure 6). As expected, T cell proliferation resulting from the coculture of nontransgenic DCs with T cells from control rats (either B7-transgenic or nontransgenic rats) was strongly inhibited by the addition of anti-CD86 but not by the addition of JJ316. Most interestingly, the proportion of Th17 cells was increased in the presence of both mAb, irrespective of the level of T cell proliferation (P = 0.0018). In contrast, the proliferation of nontransgenic T cells supported by B27-transgenic rat DCs was much weaker than that supported by nontransgenic DCs and further decreased in the presence of anti-CD86, which was consistent with the findings of our previous study (14). Nevertheless, strong T cell proliferation was induced in the presence of B27-transgenic rat DCs when JJ316 was added (Figure 6). Finally, the increased proportion of Th17 cells detected in the coculture with B27-transgenic rat DCs remained unchanged in the presence of either mAb (Figure 6). Thus, the expansion of Th17 cells during the coculture appeared to be independent of the level of proliferation induced but correlated notably with the impairment of CD86–CD28 engagement (Figure 6).

Figure 6.

Th17 cell induction by rat DCs correlates with CD86/CD28 blockade. Mature nontransgenic or B27-transgenic rat DCs isolated by the method adapted from Knight et al (29) were cocultured with CFSE-labeled lymph node CD4+CD25− T cells from nontransgenic control rats with anti-TCRα/β (R73 monoclonal antibody; 1 μg/ml) stimulation in the presence of anti-CD86 neutralizing monoclonal antibody, anti-CD28 superagonist JJ316, or isotype control. After 4 days of culture, cells were harvested, labeled, and analyzed by FACS as described in the Figure 4 legend. Dot-blots are shown for TCRα/β+-gated cells. Results are representative of 3 independent experiments. IL-17A = interleukin-17A (see Figure 4 for other definitions).


A critical role for T cells in the B27-transgenic rat model of SpA was previously established (7). Furthermore, several lines of evidence strongly support a direct role for CD4+ T cells, whereas CD8+ T cells appear to be dispensable (8–10). However, the phenotype of CD4+ T cells that might drive the disease process has not been previously thoroughly ascertained. In the present study, we observed an expansion of CD4+ T cells in the LNs draining the sites of inflammation in B27-transgenic rats. Interestingly, the frequency of CD4+ T cells has also been shown to be increased in the peripheral blood of SpA patients (32).

Among CD4+ T cells, the expansion was most striking in the CD4+CD25+ subset, which may correspond to activated or Treg cells. We showed a preferential expansion of FoxP3− activated cells compared to FoxP3+ Treg cells. We had previously shown that DCs from B27-transgenic rats have a decreased capacity to activate naive CD4+ T cells, by a mechanism implying defective costimulatory function (12, 14). We had proposed that such abnormal function could predominantly impair the capacity of B27-transgenic rat APCs to activate Treg cells. Our present results corroborate this hypothesis by showing an unbalanced control of CD4+ T cell activation.

We next characterized the cytokines produced by CD4+ T cells in the LNs draining the sites of inflammation. Notably, we observed a preferential expansion of TNFα- and IL-17A–producing cells, which paralleled disease development, likely corresponding to Th17 cells (20, 26, 33). Such an interpretation is strongly supported by the increased transcription of Rorc, IL-21, and IL-22 (which are involved in the Th17 lineage) in the LN CD4+ T cells from B27-transgenic rats (34, 35), and by the detection of IL-17A–positive mononuclear cells in arthritic joints from B27-transgenic rats (although we cannot exclude the possibility that at least some of these IL-17–producing cells were not T cells). Accordingly, up-regulation of IL-17A in CD4+ T cells from the colonic lamina propria was recently shown in the same B27-transgenic rats (36). We also detected increased levels of IL-17A in the serum of the B27-transgenic rats, similar to findings reported in human SpA (37). All of these findings support a direct role of Th17 cells in this model, although this remains to be formally demonstrated by experiments such as those involving cell transfer. (Of note, it has been impossible until now to purify live Th17 cells in the rat.)

Former studies of T cell cytokines produced in the B27-transgenic rat model of SpA demonstrated an early increase in IFNγ and IL-2 levels in the inflammatory colonic mucosa, suggesting a Th1-mediated disorder (5). Interestingly, in the present study the proportion of Th1 cells producing IFNγ alone and the expression levels of IFNγ and Tbx21 transcripts in LN CD4+ T cells from B27-transgenic rats were comparable to that in controls, suggesting that Th1 cells may not be the most critical T cells mediating the disease process. Similar observations have been reported in the context of human SpA (38). Nevertheless, we also detected a minor population of CD4+ T cells that produced IFNγ, TNFα, and IL-17A together, which was specific to the B27-transgenic rats and appeared late during the disease process. A similar phenotype of polyfunctional T cells producing both Th1 and Th17 cytokines has previously been described as selectively enriched in the peripheral blood of SpA patients (38, 39).

The role of Th17 cells has recently emerged as pivotal in several models of autoimmune and inflammatory diseases (40). Our results suggest that Th17 cells could contribute to both intestinal and joint inflammation in the B27-transgenic rat model of SpA. Such an interpretation is consistent with recent findings in human SpA. Genetics studies have identified several polymorphisms in the IL23R gene, which codes for a receptor involved in Th17 cell differentiation, as being associated with susceptibility to ankylosing spondylitis (41). Furthermore, the expansion of cells producing both TNFα and IL-17A is consistent with the striking efficacy of anti-TNFα agents, both in the B27-transgenic rat model and in human SpA (42, 43). In contrast, anti-rat IL-17 treatment failed to prevent disease in B27-transgenic rats (Glatigny S, et al: unpublished observations). Nevertheless, this result does not rule out a role for Th17 cells in rat SpA pathogenesis, since we observed an increased number of IL-17–producing T cells in the LNs of the anti–IL-17–treated rats. It is known that IL-17 negatively regulates its own production by way of negative feedback (44). Therefore, blocking IL-17 in the B27-transgenic rat led to an increase in the proportion of Th17 cells, which could still exert a pathogenic effect via the production of other mediators.

We next showed that interaction between DCs from B27-transgenic rats and CD4+ T cells from control rats contributes to the expansion of Th17 cells, despite their defective capacity to support a T cell– proliferative response (11–13). Furthermore, those IL-17A–producing cells were detected among the T cells that divided the most, suggesting that the B27-transgenic rat DCs stimulated potentially autoreactive Th17 cells, similar to what has been described with DCs isolated from human psoriatic lesions (45).

Regarding the mechanism responsible for the biased Th17 cell induction by B27-transgenic rat DCs, Transwell experiments indicated that it was not explained by a difference in soluble factor production, such as an excess of IL-23 or TGFβ, or a lack of IFNγ (results not shown). Alternatively, blocking the interaction between CD86 and CD28 during DC–T cell coculture resulted in an increased induction of Th17 cells in the nontransgenic DC condition. This effect was not correlated with the level of T cell proliferation, which was inhibited by the anti-CD86 mAb, but was conversely enhanced by the anti-CD28 superagonist mAb JJ316. Taken together, these results are consistent with a critical role played by the defective costimulatory capacity of B27-transgenic rat DCs in the biased induction of Th17 cells and could link aberrant characteristics of B27-transgenic rat DCs to their putative pathogenic role in this model.

In conclusion, these results provide a mechanism that could explain how defective function in HLA–B27–transgenic rat DCs would contribute to disease development by skewing CD4+ T cell differentiation toward a proinflammatory over a regulatory phenotype. Whether similar mechanisms also apply to SpA in humans remains to be demonstrated. However, it is worth noting that a defective stimulatory capacity of autologous and heterologous CD4+ T cells has been observed in human DCs from HLA–B27–positive ankylosing spondylitis patients (46).


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. Breban 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 conception and design. Glatigny, Fert, Lories, Chiocchia, Breban.

Acquisition of data. Glatigny, Fert, Blaton, Lories, Araujo.

Analysis and interpretation of data. Glatigny, Fert, Blaton, Lories, Araujo, Chiocchia, Breban.