We previously demonstrated that CD4+,CD25+ regulatory T (Treg) cells are present in increased numbers in the synovial fluid (SF) of rheumatoid arthritis (RA) patients and display enhanced suppressive activity as compared with their peripheral blood (PB) counterparts. Despite the presence of these immunoregulatory cells in RA, chronic inflammation persists. The purpose of the present study was to investigate whether particular proinflammatory mediators that are associated with RA could abrogate CD4+,CD25+ Treg–mediated suppression.
Monocyte phenotype was determined by flow cytometry and cytokine levels by enzyme-linked immunosorbent assay. Magnetically sorted CD4+,CD25– and CD4+,CD25+ T cells derived from the PB and SF obtained from RA patients were stimulated alone or in coculture with anti-CD3 monoclonal antibody (mAb) and autologous antigen-presenting cells, in the absence or presence of anti-CD28 mAb or the proinflammatory cytokines interleukin-6 (IL-6), tumor necrosis factor α (TNFα), or IL-7.
Monocytes from the SF of RA patients displayed increased expression of HLA class II molecules, CD80, CD86, and CD40 as compared with PB-derived monocytes, indicating their activated status. Mimicking this increased costimulatory potential, addition of anti-CD28 mAb to cocultures of CD4+,CD25– and CD4+,CD25+ T cells resulted in reduced CD4+,CD25+ Treg–mediated suppression in both PB and SF. Furthermore, IL-7 and, to a limited extent, TNFα, both of which are produced by activated monocytes and were detected in SF, abrogated the CD4+,CD25+ Treg–mediated suppression. In contrast, IL-6 did not influence Treg-mediated suppression.
Our findings suggest that the interaction of CD4+,CD25+ Treg cells with activated monocytes in the joint might lead to diminished suppressive activity of CD4+,CD25+ Treg cells in vivo, thus contributing to the chronic inflammation in RA.
CD4+,CD25+ regulatory T (Treg) cells are major contributors to the homeostasis of the immune system. CD4+,CD25+ Treg cells have been shown to inhibit T cell proliferation, cytokine production, autoantibody production, and dendritic cell and monocyte/macrophage function (1–6). Recent studies suggest that their function is defective in certain human autoimmune diseases (7), including multiple sclerosis (8), type 1 diabetes mellitus (9), and rheumatoid arthritis (RA) (10).
RA is a potentially crippling disease that is characterized by chronic joint inflammation. The inflammation is located in the synovial tissue, where the presence of proinflammatory cells and cytokines leads to damage to the cartilage and bone. We and other investigators have recently shown that in RA and juvenile idiopathic arthritis (JIA) patients, the percentage of CD4+,CD25+ T cells in synovial fluid (SF) was increased as compared with the percentage in peripheral blood (PB) (11–13). Interestingly, the ability of CD4+,CD25+ Treg cells from the SF of RA patients to suppress T cell proliferation was increased as compared with CD4+,CD25+ Treg cells from the PB of RA patients or healthy controls, suggesting that highly functional CD4+,CD25+ Treg cells are present at sites of inflammation. However, it was suggested in one study that CD4+,CD25+ T cells from the PB of patients with active RA may be defective to a certain degree, since these cells were unable to inhibit the production of cytokine (tumor necrosis factor α [TNFα] and interferon-γ [IFNγ]) production by T cells and monocytes (10).
The interaction between T cells and monocytes is very important in the pathogenesis of RA (14–16). Monocytes at the cartilage–pannus junction produce high levels of TNFα and interleukin-1 (IL-1) upon interaction with T cells, and T cells become activated by monocytes via antigen presentation and/or their cytokines. In this interaction, costimulatory molecules such as CD86/CD80 on the antigen-presenting cells (APCs) and CD28/CTLA-4 on the T cell are critical players. CTLA-4 on CD4+,CD25+ Treg cells has been reported to be essential for suppression (17–19). CD28 signaling is important as well, since CD28−/− mice lack functional CD4+,CD25+ Treg cells (20). CD28 might play a dual role though, since strong stimulation through CD28 via high levels of CD86 on dendritic cells or via anti-CD28 monoclonal antibody (mAb) can lead to abrogation of CD4+,CD25+ Treg–mediated suppression in mice and healthy humans (21–24).
The balance between inflammation and regulation in RA might be further influenced by the presence of proinflammatory cytokines in the synovium and/or SF (25). TNFα produced by synovial macrophages is a key player in RA, and increased levels of TNFα in RA patients correlate with radiologic evidence of bone damage (14, 26). Blockade of TNFα by therapy with anti-TNFα has proven to be beneficial by inhibiting inflammation and preventing joint damage in RA patients (26). This clinical effect might be mediated in part by a restoration of the defective function of CD4+,CD25+ Treg cells in patients with active RA (10, 27). Levels of IL-6 are also elevated in RA SF (28, 29), and blockade of IL-6 is beneficial in RA (30). In mice, IL-6 production by dendritic cells was shown to block CD4+,CD25+ Treg–mediated suppression, which was independent of costimulatory molecules (31). Recent work by us and by other investigators showed that IL-7 is produced by CD68+ monocytic cells in the rheumatoid synovium (32) as well as fibroblast-like synoviocytes (33). Furthermore, IL-7 induces TNFα production by RA monocytes, and IL-7 levels were shown to correlate with disease activity (32, 34). IL-7 was recently suggested to reduce Treg-mediated suppression in JIA patients (35).
Based on these data, we postulated that enhanced costimulation and proinflammatory cytokines, such as those present in vivo during synovial inflammation in RA, could abrogate CD4+,CD25+ Treg function in RA patients. Our findings are presented herein.
PATIENTS AND METHODS
RA patients were randomly selected from among those attending our outpatient clinic. PB and SF samples were obtained from the patients, and mononuclear cells were isolated using gradient centrifugation on Ficoll-Isopaque (Amersham Pharmacia Biotech, Uppsala, Sweden). Characteristics of the patients evaluated in the SF enzyme-linked immunosorbent assay (ELISA) studies and the medications they were taking are shown in Table 1.
Table 1. Presence of proinflammatory cytokines in the synovial fluid of 14 RA patients*
Approval for the study was obtained from the Institutional Medical Ethics Review Board of the University Medical Center Utrecht.
Magnetic cell separation.
PB or SF mononuclear cells were incubated with antibodies against CD8, CD14, CD16, CD19, CD36, CD56, CD123, T cell receptor γ/δ, and glycophorin A, followed by incubation with goat anti-mouse IgG magnetic beads, and column purification (Miltenyi Biotec, Bergisch Gladbach, Germany). The isolated CD4+ T cells (purity >90%) were separated into CD4+,CD25– (purity 98%) and CD4+,CD25+ (purity >80%) T cell fractions by incubation with mouse anti-human CD25 magnetic beads (Miltenyi Biotec) followed by column purification (11).
Anergy of CD4+,CD25+ T cells was defined as a proliferative response that was lower than 20% of the response of CD4+,CD25– T cells. For this analysis, CD4+,CD25– and CD4+,CD25+ T cells (105/well in 200 μl) were cultured for 3 days at 37°C in a 96-well round-bottomed plate (Nunc, Roskilde, Denmark) with anti-CD3 mAb (PeliCluster, 0.4 μg/ml; CLB, Amsterdam, The Netherlands) and irradiated (5,000 rads) autologous PB or SF mononuclear cells (105/well) as APCs. Cells were cultured in the absence or presence of anti-CD28 mAb (0.4 μg/ml; CLB), IL-6 (50 ng/ml; PeproTech, Rocky Hill, NJ), IL-7 (10 ng/ml; PeproTech), TNFα (50 ng/ml; PeproTech), or a combination of these cytokines. All cells were cultured for 3 days at 37°C in RPMI 1640 medium (Gibco-BRL Life Technologies, Merelbeke, Belgium) supplemented with 1% penicillin/streptomycin, 1% L-glutamine, and 10% heat-inactivated pooled human AB serum (ICN Biochemicals, Zoetermeer, The Netherlands). During the last 18 hours of culture, 3H-thymidine was added at 18.5 kBq/well (specific activity 925 GBq/mmole; New England Nuclear, Groningen, The Netherlands). Proliferative responses are expressed as the mean ± SEM counts per minute in triplicate wells.
For suppression assays, CD4+,CD25– T cells (105/well in 200 μl) were stimulated with anti-CD3 mAb and irradiated APCs in the absence or presence of CD4+,CD25+ T cells (responder:suppressor ratios of 1:0 and 1:1, respectively). To control for consumption of nutrients or high cell numbers, experiments were set up in parallel in which CD4+,CD25– T cells were cocultured with CD4+,CD25– T cells at a ratio of 1:1, which did not lead to suppression, but rather an increase in proliferation (data not shown). Cultures were costimulated with anti-CD28 mAb and proinflammatory cytokines (IL-6, IL-7, and TNFα) as described above. The percentage inhibition was calculated as the relative difference in proliferative response between CD4+,CD25– T cells cultured alone and CD4+,CD25– T cells cultured in the presence of CD4+,CD25+ T cells.
Additional suppression assays were performed using responder T cells from RA SF that had been labeled with carboxyfluorescein diacetate succinimidyl ester (CFSE-DA) (4 μM; Molecular Probes Invitrogen, Paisley, UK). Cells (105/well in 200 μl) were stimulated with CD3/CD28 Dynabeads (1 bead per cell; Invitrogen, Paisley, UK) in the absence or presence of CD4+,CD25+ Treg cells at a 1:1 ratio. Additional costimulation was provided by adding anti-CD28 mAb (0.4 μg/ml; CLB) to the wells. After 3 days, the cells were analyzed by flow cytometry to determine the percentage of cells that had divided.
The presence of IL-6 (BioSource, Etten Leur, The Netherlands), IL-7 (Diaclone, Besançon, France), and TNFα (R&D Systems, Minneapolis, MN) in SF samples was detected using commercially available kits. Detection limits were 10, 1, and 10 pg/ml, respectively. SF samples obtained from RA patients were centrifuged, and the supernatants were stored at −80°C until they were used. Before use, SF samples were treated for 30 minutes at 37°C with 25 units/ml of hyaluronidase (Sigma, Zwijndrecht, The Netherlands).
For analysis of monocyte phenotype, mononuclear cells were isolated from paired PB and SF samples obtained from RA patients and were stained for fluorescence-activated cell sorter analysis using anti-CD4 phycoerythrin (PE)–Cy5–labeled (Dako, Glostrup, Denmark) and mAb against CD25 (PE/fluorescein isothiocyanate [FITC]–labeled), HLA class II (FITC-labeled), CD80 (FITC-labeled), CD86 (PE-labeled), CD40 (PE-labeled), CD127 (FITC-labeled), and CD210 (PE-labeled) (all from BD Biosciences, San Jose, CA). Viable T cells and monocytes were gated based on their forward-scatter/side-scatter profile and their CD4high (T cells) or CD4low (monocytes) expression. The expression of cell markers was analyzed using CellQuest software (BD Biosciences), and the results are expressed as geometric mean fluorescence intensity (MFI).
To test the differences in monocyte phenotype, T cell proliferative response, and percentage inhibition between cultures with and without costimulation or cytokines, we used Wilcoxon's matched pairs signed rank test.
Enhanced costimulation-induced reversal of CD4+,CD25+ Treg–mediated suppression of T cell proliferation in PB and SF from RA patients.
To investigate the effects of costimulation on CD4+,CD25+ Treg–mediated suppression in RA, we determined the expression levels of costimulatory and HLA class II molecules on monocytes from RA patients in paired samples of PB and SF (Figure 1). CD80, CD86, and HLA class II molecule expression levels were significantly higher on monocytes from SF than on monocytes from PB (mean ± SEM 9 ± 1 versus 27 ± 7 MFI for CD80; 85 ± 17 versus 131 ± 22 MFI for CD86, and 140 ± 34 versus 687 ± 85 MFI for HLA class II; P < 0.05). CD40 expression was low on SF-derived monocytes but was still significantly higher than that on PB-derived cells (9 ± 2 versus 17 ± 4 MFI; P = 0.030) (data not shown).
The enhanced expression of CD80, CD40, and CD86 on SF-derived monocytes suggested that increased costimulatory signals were received by synovial T cells, which might alter the balance between regulation and inflammation. To investigate specifically the influence of B7/CD28-mediated costimulation on CD4+,CD25+ Treg–mediated anergy and suppression, CD4+,CD25– T cells and CD4+,CD25+ Treg cells were isolated from PB or SF samples from RA patients and cultured either alone or in coculture with each other, in the absence or presence of anti-CD28 mAb. Anti-CD3 mAb and autologous APCs were present in all cell cultures.
Addition of anti-CD28 mAb (0.4 μg/ml) led to a dramatic increase in proliferation of both CD4+,CD25– T cells and CD4+,CD25+ T cells from the PB of RA patients (from a mean ± SEM of 18,000 ± 4,000 cpm to 110,000 ± 14,000 cpm [P = 0.018] for CD25– T cells and from 1,600 ± 1,000 to 40,000 ± 7,000 cpm [P = 0.028] for CD25+ T cells) (Figures 2A and C). According to our definition of anergy (i.e., a proliferative response <20% of the response of CD4+,CD25– T cells), the anergic state of the CD4+,CD25+ T cell population was reversed, since their relative proliferative response compared with the CD25– T cell response increased from a mean ± SEM of 8 ± 3% to 41 ± 8% (P = 0.028).
CD4+,CD25+ T cells from SF also showed a significant increase in proliferation upon addition of anti-CD28 mAb (from 3,800 ± 3,000 cpm to 30,000 ± 14,000 cpm; P = 0.043), leading to a noticeable reversal of their unresponsive state (6 ± 4% versus 33 ± 22%; P = 0.06) (Figure 2D). Despite this increase in proliferation, CD4+,CD25+ Treg cells did not produce IFNγ (PB and SF; n = 2) (data not shown), indicating that these cells did not convert into fully competent effector T cells. CD4+,CD25– T cells from SF displayed a much stronger proliferative potential in response to anti-CD3 mAb and APCs than did PB-derived CD4+,CD25– T cells (77,000 ± 12,000 cpm versus 18,000 ± 4,000 cpm) (compare Figure 2B with Figure 2A), which was further enhanced by the addition of anti-CD28 mAb (105,000 ± 15,000 cpm; P = 0.043).
Next, we investigated the influence of anti-CD28 mAb on the actual suppressive capacity of CD4+,CD25+ Treg cells from PB or SF. In the absence of anti-CD28 mAb, CD4+,CD25+ Treg cells from PB were able to suppress CD4+,CD25– T cell proliferation by 52 ± 4%, which is consistent with our previous findings (11). Addition of anti-CD28 mAb led to a moderate decrease in this inhibition (to 33 ± 8%; P = 0.025) (Figure 2E). This coincided with a significant increase in proliferation of the cocultures (CD25– and CD25+ T cells cultured together) upon addition of anti-CD28 mAb (8,000 ± 1,900 cpm versus 70,000 ± 12,000 cpm; P = 0.018) (data not shown).
In SF, the addition of anti-CD28 mAb led to a strong increase in proliferation of the cocultures (26,000 ± 8,000 versus 86,000 ± 16,000; P = 0.043) (data not shown) and a profound decrease in CD4+,CD25+ Treg–mediated suppression (60 ± 13% inhibition versus 15 ± 17% in the absence and presence of anti-CD28 mAb, respectively; P = 0.043). Since 3H-thymidine incorporation assays do not distinguish between CD4+,CD25– and CD4+,CD25+ T cell proliferation, we performed additional assays using CFSE-DA–labeled responder T cells from SF. For this analysis, we had to alter our stimulation conditions in order to achieve sufficient cell division to be observed by flow cytometry. Responder T cells were therefore stimulated with anti-CD3/CD28 mAb–coated beads in the absence or presence of CD4+,CD25+ Treg cells, with or without additional anti-CD28 mAb (0.4 μg/ml) to provide enhanced costimulation.
Upon stimulation with beads, 78% of the responder T cells had divided after 3 days. This cell division was reduced by almost 30% in the presence of Treg cells (average of 2 experiments) (data not shown). When the same coculture was performed in the presence of enhanced CD28-mediated costimulation, cell division was inhibited by only 19% (average of 2 experiments). These data support our results from the 3H-thymidine incorporation experiments, showing that costimulation through CD28 can reduce CD4+,CD25+ Treg–mediated suppression in RA patients.
IL-7–induced reduction of the suppressive function of CD4+,CD25+ T cells from RA synovial fluid.
We next investigated whether the proinflammatory cytokines TNFα, IL-6, and IL-7, which are produced in large amounts by monocytes and other cells in the rheumatoid joint, could diminish the suppressive capacities of CD4+,CD25+ T cells from RA PB or SF. First, we confirmed the presence of these cytokines in 14 SF samples from RA patients (Table 1). TNFα was detected in 10 of the 14 samples (mean ± SEM 66 ± 26 pg/ml), IL-7 in 6 of the 14 samples (62 ± 29 pg/ml), and IL-6 in 11 of the 14 samples (16 ± 7 ng/ml).
To investigate the effects of these cytokines on the proliferation and immune regulation of T cells from RA SF, we cultured SF-derived CD4+,CD25– T cells or CD4+,CD25+ Treg cells either alone or in coculture with each other in the absence or presence of TNFα, IL-7, or IL-6, or a combination of these cytokines (Figure 3). In all cultures, autologous APCs and anti-CD3 mAb were present. In the absence of additional cytokines, CD4+,CD25+ T cells from SF proliferated poorly in response to stimulation with anti-CD3 mAb and APCs (mean ± SEM 4 ± 1% relative to CD4+,CD25– T cell proliferation). The addition of TNFα (50 ng/ml) led to some increase in proliferation in both CD4+,CD25– T cells and CD4+,CD25+ T cells in SF, but this difference was not statistically significant. In contrast, IL-7 (10 ng/ml) induced a significant increase in proliferation of CD4+,CD25+ T cells in SF (700 ± 300 versus 6,000 ± 1,700; P = 0.028), whereas IL-6 (50 ng/ml) had no effect on the proliferation of either T cell subset. Adding a mixture of the 3 cytokines led to a significant increase in proliferation of CD4+,CD25+ T cells (2,000 ± 1,700 to 15,000 ± 4,500; P = 0.03).
We then looked at the effects of these cytokines on T cell suppression. Initially, we cultured T cells from SF at a 1:1 ratio of CD4+,CD25– to CD4+,CD25+ T cells. At this ratio, a higher percentage of inhibition was observed in SF as compared with PB (53 ± 3% in PB [n = 6] versus 97 ± 1% in SF [n = 2]), which is consistent with our previous findings (11). When cytokines were added to these cultures, no effects of the cytokines on the percentage of inhibition were observed (data not shown).
We have previously shown that the balance between CD25– responder T cells and CD25+ suppressor T cells determines the level of suppression. Therefore, we hypothesized that a change in the ratio of responder and suppressor T cells to 1:0.5 would alter the impact of proinflammatory cytokines on CD4+,CD25+ Treg–mediated suppression. A 1:0.5 ratio of CD4+,CD25– to CD4+,CD25+ T cells more closely approaches the physiologic ratio of these cells in SF (11). Indeed, when this ratio was used, profound effects were observed when IL-7 was added to the cultures (n = 5), with the percentage of inhibition decreasing from 67 ± 14% to 21 ± 10% in the presence of IL-7 (P = 0.04) (Figure 4C). Addition of TNFα reduced CD4+,CD25+ Treg–mediated suppression from 72 ± 10% to 45 ± 13%, although this effect did not reach statistical significance (P = 0.08 [n = 4]). Addition of IL-6 (n = 5) to these cocultures had no effect on CD4+,CD25+ Treg–mediated suppression. The mixture of cytokines also led to a significant decrease in CD4+,CD25+ Treg–mediated suppression (from 67 ± 14% to 16 ± 9%; P = 0.043), which was likely due to the presence of IL-7 in combination with TNFα.
Importantly, despite the fact that IL-7 or the mixture of cytokines significantly enhanced the proliferation of CD4+,CD25+ Treg cells from SF (Figure 4B), they did not reverse the anergic state as compared with CD4+,CD25– T cells. The relative proliferation of CD4+,CD25+ Treg cells increased from 1% in the absence of cytokines to 7% or 12% in the presence of IL-7 or the mixture, respectively. The CD4+,CD25– T cells from SF did not proliferate significantly more upon stimulation with anti-CD3 mAb and IL-7 or the cytokine mixture as compared with stimulation with anti-CD3 mAb alone. These observations suggest that the decreased percentage inhibition in SF cultures upon addition of IL-7 or the cytokine mixture was due to the effects of the cytokines on the suppressive capacity, rather than the proliferative response, of either CD4+,CD25– T cells or CD4+,CD25+ Treg cells.
We also tested the effects of the cytokines on the reversal of CD4+,CD25+ Treg–mediated suppression in PB. Similar to our findings in SF, no significant reversal of suppression was observed in PB when CD25– and CD25+ T cells were cocultured at a ratio of 1:1 (data not shown). IL-7 and the cytokine mixture significantly increased the proliferation of both CD4+,CD25– and CD4+,CD25+ T cells in PB, leading to a moderate reversal of T cell anergy (8% versus 23% or 37% for the relative response of CD25+ T cells cultured without additional cytokines, with IL-7, or with the mixture, respectively). In contrast to SF, when CD4+,CD25– and CD4+,CD25+ T cells from PB were cocultured at a ratio of 1:0.5, no inhibition was detected, and thus the influence of the cytokines could not be assessed (data not shown).
To elucidate how IL-7 might be altering Treg-mediated suppression, we investigated several possible mechanisms. Since IL-7 receptor α (IL-7Rα) expression on T cells is essential for the effects of IL-7, we studied the expression of IL-7Rα chain (CD127) on CD4+,CD25– and on CD4+,CD25bright T cells from PB (n = 8) and SF (n = 6) samples from RA patients (the CD4+,CD25bright T cell subset contains the Treg cells with the most suppressive function ). We observed a significantly lower expression of CD127 on CD4+,CD25bright T cells as compared with CD4+,CD25– T cells in both PB (P = 0.0078) and SF (P = 0.03) (Figure 4), suggesting that IL-7 may be acting on the responder T cells rather than the regulatory T cells.
We also investigated whether IL-7 diminished IL-10 or TGFβ production by synovial Treg cells. However, we found that Treg-mediated suppression of T cell proliferation in SF was independent of these 2 cytokines, since suppression was still observed when neutralizing mAb to IL-10 and TGFβ were added in suppression assays (75 ± 8 versus 73 ± 4% suppression in the absence or presence of neutralizing mAb; n = 2) (data not shown). Based on these findings, we did not expect to find an effect of IL-7 on IL-10R (CD210) expression on responder T cells, and this was indeed confirmed by flow cytometry (n = 2) (data not shown).
We have previously shown that CD4+,CD25+ Treg cells from RA patients have the capacity to inhibit both T cell proliferation and cytokine production ex vivo (11). However, in vivo, these T cells operate in a highly inflammatory milieu and interact with activated cells, such as T cells and monocyte/macrophages. We hypothesized that the interaction with these activated cells, through costimulatory molecules and proinflammatory cytokines, determines the level of suppression in vivo.
We demonstrated that monocytes from SF displayed an activated phenotype, with high levels of CD80, CD86, CD40, and HLA class II molecules compared with PB-derived monocytes. This is consistent with the literature, which describes highly activated macrophages in the joint, mainly at the cartilage–pannus junction (14). To mimic signaling via the B7 family of costimulatory molecules, we added anti-CD28 mAb to cocultures of responder and regulatory T cells. This led to a partial abrogation of CD4+,CD25+ Treg–mediated suppression in PB and a more profound reversal of suppression in SF cell cultures. This novel finding in RA patients adds to the findings of previous studies on the effects of anti-CD28 mAb on Treg cells in mice and in healthy human controls (21–24).
Besides the effects of costimulation, we investigated the effects of a set of proinflammatory cytokines that are produced in large amounts in the rheumatoid synovium and have been shown to be important in the pathogenesis of RA. Due to limitations in patient samples and numbers of Treg cells that can be isolated for functional studies, we had to restrict our studies to a selected set of cytokines. We showed previously that IL-7 levels are increased in the serum of RA patients as compared with that of healthy controls and that these levels correlate with increased disease activity (34). Moreover, we have shown that IL-7 is abundantly present in RA synovial tissue and in RA synovial fluid (Table 1 and ref.32). In addition, it was recently shown that IL-7 may inhibit Treg function in JIA (35). IL-7 could therefore be an important immunoregulatory cytokine in RA. Indeed, when we tested the effects of IL-7 on CD4+,CD25+ Treg–mediated suppression in SF, we found that IL-7 induced a profound loss of suppression. IL-7 did not reverse the anergic state of CD4+,CD25+ Treg cells, however, and we found that the IL-7Rα chain was expressed at much lower levels on CD4+,CD25+ Treg cells than on CD4+,CD25– T cells. Together, these data suggest that IL-7 might act on the CD4+,CD25– responder T cells rather than the CD4+,CD25+ Treg cells.
In our experiments, we added 10 ng/ml of recombinant human IL-7, whereas the concentration of IL-7 in SF ranged from 15 to 186 pg/ml, and up to 500 pg/ml in previous studies (32). However, we believe this is a physiologically relevant concentration, since we previously showed that IL-7 influences effector T cell function (e.g., IFNγ production) at very low dosages, starting at 10–100 pg/ml, with optimal responses at 10 ng/ml (34). Moreover, we expect that the local concentration of IL-7 in the synovium will exceed the levels measured in SF, given the abundant expression in RA synovial tissue. In addition, we found a synergistic effect of the addition of low doses of IL-7 (1 ng/ml) and CD28-mediated costimulation (0.04 μg/ml). The inhibitory capacity of Treg cells was reduced, with 7 ± 3%, 37 ± 31%, and 62 ± 9% in the presence of IL-7 (1 ng/ml), anti-CD28 (0.04 μg/ml), or a mixture of IL-7 and anti-CD28, respectively (n = 3) (data not shown). Taken together, we are confident that the data obtained with IL-7 are physiologically relevant.
Because of the good clinical results following anti-TNFα therapy (26), we were interested in the effect of TNFα on Treg activity. This interest was further nurtured by recent reports suggesting that anti-TNFα therapy increased CD4+,CD25+ T cell numbers and/or function in the PB of RA patients (10, 27). Effects of TNFα on CD4+,CD25+ Treg cells were further observed in type 1 diabetes mellitus–prone NOD mice, which have a relative deficiency of CD4+,CD25+ Treg cells: neonatal administration of TNFα decreased, while anti-TNFα mAb increased the number of these cells in thymus and spleen of these mice. In addition, transfer of CD4+,CD25+ T cells from mice treated with TNFα led to a rapid onset of diabetes mellitus, whereas cells from untreated mice successfully delayed disease onset (37). These studies suggested that anti-TNFα therapy may influence suppression by Treg cells in vivo, although it remains possible that some of these effects are achieved indirectly via changes in disease activity.
Our data showed that the in vitro addition of TNFα to cultures of SF-derived CD4+,CD25+ Treg cells led to some reversal of the Treg-mediated suppression, suggesting that TNFα may indeed have direct effects on Treg function. This is supported by a recent study showing that TNFα blocked the suppressive activity of CD4+,CD25+ Treg cells from the PB of healthy controls (27). We also observed that upon addition of TNFα, the proliferation of the cocultures (CD25– and CD25+) was significantly increased (P = 0.043) (data not shown). This shows that TNFα does lead to enhanced T cell proliferation, which might abrogate tolerance and contribute to joint damage in RA.
No effects of IL-6 on T cell proliferation or Treg function were observed in SF or PB. This was quite surprising, since recent data in mice suggested that IL-6 production by dendritic cells could abrogate the suppressive capacities of CD4+,CD25+ Treg cells, either directly (38) or via effects on CD25– responder T cells (31). An explanation for our findings might be that human Treg cells and/or responder T cells are less susceptible to these effects of IL-6 than are their murine counterparts.
Our findings demonstrate that proinflammatory mediators such as IL-7 and TNFα, and costimulation affect CD4+,CD25+ Treg function in RA. These findings complement recently reported data on the abrogation of CD4+,CD25+ Treg function in PB from patients with JIA (35), and together the results indicate that proinflammatory mediators might contribute to various chronic inflammatory or autoimmune diseases directly, as well as through their effects on CD4+,CD25+ Treg cells. Importantly, our data also show that raising the ratio of Treg cells to responder T cells negates the reversing effects of IL-7. These data might provide some answers as to why chronic inflammation in the rheumatoid joint persists despite the increased presence of CD4+,CD25+ regulatory T cells. In this context, 2 recent studies are of interest since they revealed that TGFβ, which is thought to play an important role in Treg induction and/or function in vivo (39–42), skews naive T cells toward an IL-17–producing (Th17) phenotype in the presence of an inflammatory cytokine milieu (IL-6/TNFα/IL-1β) (43, 44). Th17 cells are noted for their role in mediating inflammation, including that in RA. Importantly, both RA SF and synovium contain large amounts of TGFβ (45, 46), IL-17 (47), as well as IL-6 and TNFα (Table 1) (26, 28, 29, 48). These findings thus indicate that the proinflammatory environment may counteract immunoregulation at several levels.
Our findings provide further evidence that elimination of proinflammatory mediators such as IL-7 or TNFα from the joint by antibody therapy might be beneficial not only by suppressing proinflammatory processes, but also by enlarging the effects of CD4+,CD25+ Treg–mediated suppression. This argument, together with the notion that CD4+,CD25+ T cells can inhibit T cell (11) and monocyte (6) function, strengthens the concept that an increase in the function and/or numbers of regulatory T cells in the joints of RA patients could be an attractive therapeutic goal.
Dr. Taams 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. Van Amelsfort, van Roon, Bijlsma, Lafeber, Taams.
Acquisition of data. Van Amelsfort, van Roon, Noordegraaf, Jacobs, Taams.
Analysis and interpretation of data. Van Amelsfort, van Roon, Noordegraaf, Jacobs, Bijlsma, Lafeber, Taams.
Manuscript preparation. Van Amelsfort, van Roon, Bijlsma, Lafeber, Taams.
Statistical analysis. Van Amelsfort, van Roon, Lafeber, Taams.