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Clin Exp Immunol. Apr 2000; 120(1): 174–182.
PMCID: PMC1905626

Down-regulation of the T cell receptor CD3ζ chain in rheumatoid arthritis (RA) and its influence on T cell responsiveness

Abstract

T cells implicated in chronic inflammatory diseases such as RA respond weakly when stimulated in vitro with mitogen or antigen. The mechanism behind this hyporesponsiveness is unclear, but a depressed expression of the T cell receptor (TCR)-associated CD3ζ chain has been suggested. In the present work we describe a low expression of CD3ζ in synovial fluid (SF) T cells from RA patients compared with peripheral blood (PB) T cells, but no difference in CD3ζ expression between RA and healthy control PB T cells. In vitro studies demonstrated that granulocytes but not SF macrophages are able to down-regulate the expression of CD3ζ. Through stimulation with anti-CD3 antibodies we demonstrated that the TCR-dependent proliferative response was decreased in SF T cells compared with PB T cells. Stimulation with phorbol ester and ionomycin also resulted in a low proliferative response of SF T cells, indicating that both signal transduction through the TCR (stimulation with anti-CD3) and events further downstream in the signalling pathways (stimulation with phorbol ester and ionomycin) are affected. A similar depression of T cell activity was observed when induction of IL-2 and IL-4 was measured. However, SF T cells were not defective in the induction of interferon-gamma (IFN-γ) when stimulated with phorbol myristate acetate (PMA)/ionomycin, in contrast to the diminished IFN-γ response observed after stimulation with anti-CD3. This indicates that the hyporesponsiveness of SF T cells can not be generalized to all T cell functions. The differential response to external stimuli is likely to be of importance for the capacity of SF T cells to influence inflammatory reactions.

Keywords: CD3ζ, rheumatoid arthritis, synovial fluid, cytokine, proliferation

INTRODUCTION

It has long been known that T cells from patients with chronic inflammation or long-standing infections respond poorly when stimulated in vitro with antigen or mitogen (for review see [1]), but the mechanism(s) behind this hyporesponsiveness is unknown. For some of these conditions, i.e. RA, systemic lupus erythematosus (SLE), cancer and HIV infection, a depressed expression of the T cell receptor (TCR)-associated CD3ζ chain has been reported [212].

In studies of the expression of CD3ζ in different malignancies some attention has been given to the functional consequences of a low expression of CD3ζ. In tumour-infiltrating lymphocytes (TIL) from renal cell carcinoma patients CD3ζ expression varied from patient to patient, but all patients had a low proliferative response when stimulated with phytohaemagglutinin (PHA) or anti-CD3. Moreover, a correlation was evident between levels of tumour necrosis factor-alpha (TNF-α) produced after anti-CD3 or PHA stimulations and the expression of CD3ζ[13]. The CD3ζ expression on lymphocytes from colorectal carcinoma patients is low in TIL and increases with distance from the tumour, as analysed on T cells derived from nearby lymph nodes [14]. Subsequent studies have demonstrated that activated macrophages are able to down-regulate the expression of CD3ζ in vitro through the production of H2O2[1517].

In RA, peripheral blood (PB) and synovial fluid (SF) T cell responses to mitogen and antigen stimulations have previously been described as defective [8,1827], but it is still unclear whether a low expression of CD3ζ can fully explain the TCR-dependent hyporesponsiveness or is simply a consequence of some other down-regulatory mechanism. In the present study we analysed the expression of CD3ζ in different T cell subpopulations in PB and SF from RA patients and compared it with the expression in PB from healthy controls (HC). We recorded a depressed expression of CD3ζ in SF T cells, but observed a similar level of expression of CD3ζ between PB T cells from RA patients and HC. Hence, the role of inflammatory cells in the down-regulation of CD3ζ was studied in vitro by incubating PB T cells from RA patients together with autologous SF macrophages or granulocytes. In order to investigate whether a depressed expression of CD3ζ could explain the hyporesponsiveness observed in RA T cells, the ability of these cells to respond to anti-CD3 stimulation and stimulation with calcium ionophore in combination with phorbol ester was characterized. The T cell response was determined by analysing proliferation of different T cell subsets as well as the production of cytokines (interferon-gamma (IFN-γ), IL-2, IL-4, IL-10).

PATIENTS and METHODS

Patients

Nineteen patients meeting the American College of Rheumatology classification criteria for RA [28] (16 females and three males, mean age 53 years, range 24–81 years) and 16 sex-matched HC (mean age 43 years, range 24–58 years) were included in this study.

Cell preparation

PB or SF were aseptically collected into heparinized tubes and diluted 1:2 for blood and 1:3 for SF in PBS within 1 h of sample collection. PB mononuclear cells (PBMC) or SFMC were density gradient separated and cells were washed three times in PBS followed by suspension either in PBS for CD3ζ staining, or for cell stimulation in RPMI 1640 (Flow Labs, Irvine, UK) supplemented with 10% fetal calf serum (FCS), penicillin, streptomycin, HEPES and glutamine (complete medium).

Purification of T cells, macrophages and granulocytes

T cells and macrophages were purified using negative selection with magnetic beads. For T cell purification, PBMC or SFMC were incubated with MoAb to CD32 (clone AT10 Serotec, Oxford, UK) for 30 min at room temperature. The cells were washed and anti-mouse IgG-coupled Dynabeads (Dynal, Oslo, Norway) were added and separation was performed according to the manufacturer’s recommendations. The purity of the T cells was typically >85% as determined by FACS analysis of CD3 expression. For macrophage purification, T cells were removed from SFMC by incubation with CD3-conjugated Dynabeads (Dynal) typically yielding a purity of 90% as measured by staining for CD14. For studies of the mechanism of CD3ζ down-regulation, granulocytes were taken from the bottom of the SF Ficoll separation followed by extensive washing. Contaminating cells in the granulocyte fraction were generally <10%.

Analysis of CD3ζ expression

Six million mononuclear cells were fixed with 5 ml 4% paraformaldehyde diluted in PBS for 10 min at 4°C immediately after separation. The cells were centrifuged and the paraformaldehyde discarded before washing the cells twice with 1% bovine serum albumin (BSA) and 0·02% NaN3 in PBS. The cell suspension was aliquoted for staining and permeabilized with 0·5 ml 0·1% saponin in PBS–BSA–NaN3 for 10 min at room temperature. One microgram/sample of an α-CD3ζ antibody (TIA-2; Coulter, Hialeah, FL), isotype-matched irrelevant antibody (Dakopatts, Glostrup, Denmark) or antibody specific for vimentin (Dakopatts) was added and incubated for 45 min at room temperature (the vimentin antibody was used as a permeabilization control). After two washes in PBS–BSA–saponin, 2 μl of a FITC-conjugated anti-mouse IgG antibody (Dakopatts) were added. After 30 min, the cells were washed twice in PBS–BSA–saponin and once in PBS. The anti-mouse antibody was blocked by adding 0·2 μg normal mouse IgG (Dakopatts) for 10 min before surface staining was performed using PE-conjugated antibodies to CD4, CD8, irrelevant antibody (Becton Dickinson, San José, CA) or Cy-5-conjugated antibody to CD3ε (PharMingen, San Diego, CA). The cells were washed twice in PBS and immediately analysed in a FACSort (Becton Dickinson).

To ensure that the low CD3ζ expression in RA SFMC was not due to protease degradation of CD3ζ in vitro, a mixture of protease inhibitors was added to two SF samples immediately after sampling and was present throughout the cell separation procedure. The protease inhibitor mixture has been described as adequate for inhibition of proteases produced by activated granulocytes [29] and consisted of 1 mm sodium orthovanadate (Sigma, St Louis, MO), 5 mm EDTA, 10 μg/ml leupeptin (Sigma), 10 μg/ml aprotinin (Sigma), 0·825 μg/ml p-nitrophenylguanidinobenzoate (Sigma), 2 mm PMSF (Sigma), 100 μg/ml chymostatin (Boehringer Mannheim, Germany) and 100 μg/ml trypsin-chymotrypsin inhibitor (Sigma).

In vitro assay of CD3ζ down-regulation

Purified PB T cells were incubated with or without SF macrophages or granulocytes (ratio 5:1) in complete medium overnight. Cells were then collected and the expression of CD3ζ was analysed as above.

Cell stimulation

Flat-bottomed 96-well cell culture plates (Costar, Cambridge, MA) were coated with 50 μl/well, 10 μg/ml of anti-CD3 (OKT3; ATCC, Rockville, MD) diluted in carbonate buffer pH 9·6 in PBS at 4°C overnight. Cells were diluted to 1 × 106/ml in complete medium and 300 μl were added to each well either coated with anti-CD3 or with the addition of 0·2 ng/ml phorbol 12-myristate 13-acetate (PMA; Sigma) and 0·5 μm ionomycin (Sigma). The concentration of PMA was chosen in preliminary tests to minimize cellular death and to give a comparable proliferative response to that of anti-CD3 stimulation. Cells added to wells not coated with anti-CD3 were used as unstimulated controls. For in vitro studies on functional consequences of granulocyte-induced down-regulation of CD3ζ, PBMC were stimulated with anti-CD3 in the presence of granulocytes at a ratio of 5:1. Granulocytes were added on day 0 and day 1. Supernatants were collected after 24 h for the analysis of IL-4 and at day 3 for the analysis of IFN-γ, IL-2 and IL-10.

Analysis of cell proliferation

The proliferation assay was essentially performed according to Esin et al. [30]. At day 3, 5-bromo-2′-deoxyuridine (BrdU; Sigma) was added to the above described cell cultures at a final concentration of 10 μm followed by incubation at 37°C for a further 6 h. Thereafter, cells were collected for proliferation analysis. The BrdU-incorporated cells (approximately 1 × 106 cells) were washed once in PBS and divided into four tubes for surface staining with PE-conjugated antibodies to CD3, CD4, CD8 (Becton Dickinson) or isotype-matched irrelevant control antibody, respectively. After one wash in PBS the cells were fixed with 0·5 ml 1% formaldehyde/0·01% Tween in PBS and stored at 4°C up to 1 week. The cell suspension was then washed once in Earle’s balanced salt solution (EBSS) (Gibco, Paisley, UK) and incubated with 0·5 ml DNase I (Sigma) diluted to 50 U/ml in EBSS for 30 min. The cells were washed once in 0·5% Tween/PBS and 0·125 μg FITC-conjugated anti-BrdU antibody (Becton Dickinson) or irrelevant FITC-conjugated isotype-matched control antibody (Dakopatts) was added and incubated for 45 min at room temperature. The cells were then washed twice in 0·5% Tween/PBS and analysed in a FACSort.

Cytokine immunoassay

Briefly, ELISA plates (Maxisorp; Nunc, Roskilde, Denmark) were coated with 50 μl of 2 μg/ml catcher antibody and blocked with 100 μl PBS−1% BSA. Washing was performed with PBS/0·05% Tween and standard recombinant cytokine (IFN-γ (R&D Systems, Minneapolis, MN), IL-2 (a generous gift from Hoffman-LaRoche, Basel, Switzerland), IL-10 (a generous gift from Shering-Plough, Madison, WI) diluted in culture medium or samples were added. After washing, the secondary biotinylated antibody was added at a concentration of 1 μg/ml. Plates were washed and avidin-alkaline phosphatase (Dakopatts) was added. Substrate (p-nitrophenyl-phosphate tablets (1 mg/ml; Sigma) in diethanolamine buffer pH 9·8) was added after washing and the reaction was monitored at 405 nm in a spectrophotometer. The antibodies used were: IFN-γ, 1-D1K (MabTech, Stockholm, Sweden) as catcher and 7-B6-1 (MabTech) as biotinylated detector; IL-10, 9D7 (PharMingen) as catcher and 12G8 (PharMingen) as biotinylated detector; IL-2, Genzyme’s duokit (Genzyme, Cambridge, MA). For IL‐4 determinations a commercial ELISA kit was used according to the manufacturer’s instructions (IL-4-HS; R&D).

Analysis of intracellular IFN-γ by flow cytometry

After stimulation with PMA/ionomycin (20 ng/ml and 0·5 μm, respectively) overnight in the presence of 5 μg/ml brefeldin A (Sigma), the cells were collected and fixed for 10 min at 4°C with 4% paraformaldehyde in PBS. Cells were then washed once in PBS–BSA and permeabilized as described for the CD3ζ assay. FITC-conjugated anti-IFN-γ antibody (1 μg; PharMingen) was added and allowed to bind for 30 min at room temperature. Cells were then washed twice with PBS–BSA–saponin and once with PBS before surface staining with PE-conjugated antibodies specific for CD94 (PharMingen) together with Cy-5-conjugated antibody specific for CD3ε (PharMingen). Cells were incubated for 15 min at room temperature followed by washing twice in PBS and were thereafter analysed in a FACSort.

Statistical analysis

Non-parametric methods were used throughout the study. Differences between groups were analysed using Mann–Whitney’s U-test and paired analyses utilized Wilcoxon’s signed rank test. The limit for statistical significance was set to 0·05 (two-tailed). Box plots depict the median as a line and the 25th and 75th percentiles limiting the box with the 10th and 90th percentiles indicated with bars.

RESULTS

CD3ζ is down-regulated in SF T cells

A pronounced down-regulation of CD3ζ was observed in RA SFMC compared with RA PBMC in both CD4+ and CD8+ T cells. No difference in CD3ζ expression between RA PBMC and HC PBMC could be detected in any cell population (Fig. 1). The low expression of CD3ζ in RA SF T cells was not due to degradation in vitro by proteases released by activated granulocytes since addition of protease inhibitors immediately upon SF sampling resulted in comparable levels of CD3ζ expression to cells separated without inhibitors (data not shown). A comparable degree of down-regulation of CD3ζ on SF T cells was also observed using another CD3ζ-specific MoAb (6B10.2; Santa Cruz Biotechnology, Santa Cruz, CA, data not shown).

Fig. 1
Down-regulated CD3ζ expression in RA synovial fluid mononuclear cells (SFMC) but not in RA peripheral blood mononuclear cells (PBMC). Expression of CD3ζ was analysed by intracellular cytometric staining on density-separated PBMC and SFMC ...

Granulocytes but not macrophages are capable of down-regulation of CD3ζ in vitro

In order to investigate the role of accessory cells in the down-regulation of CD3ζ, purified PB T cells were coincubated with autologous macrophages or granulocytes isolated from SF. The expression of CD3ζ was measured after overnight incubation. As evident in Fig. 2, granulocytes, but not macrophages, were able to down-regulate CD3ζ in vitro. The extent of CD3ζ down-regulation was dependent on the T cell:granulocyte ratio, a ratio of 5:1 (Fig. 2) but not 40:1 (data not shown) resulting in a marked down-regulation. Similar results were obtained if peripheral granulocytes were used and also if cells were incubated in serum-free AIM-V medium (Gibco), indicating that anti-oxidants present in FCS do not affect the in vitro down-regulation of CD3ζ (data not shown). The down-regulation of CD3ζ in vitro was accompanied by a diminished T cell responsiveness upon stimulation with anti-CD3 as measured by proliferation and IFN-γ secretion (Table 1).

Fig. 2
Down‐regulation of CD3ζ in T cells by granulocytes but not by macrophages. Purified RA peripheral blood (PB) T cells were coincubated with autologous synovial fluid (SF) macrophages or autologous SF granulocytes overnight and the T cell ...
Table 1
Functional effects of granulocyte‐mediated down‐regulation of CD3ζ

The proliferative response of SF T cells is suppressed

The proliferative response to anti-CD3 stimulation was markedly suppressed in SF T cells compared with RA PB T cells in both the CD4 and CD8 subsets (Fig. 3a). An unexpected tendency for an increased proliferative response to anti-CD3 stimulation was apparent in RA PB T cells compared with HC PB T cells, but this only reached the limit of significance when analysing all CD3+ T cells.

Fig. 3
Defective proliferative response of RA synovial fluid (SF) T cells but not of RA peripheral blood (PB) T cells. Proliferation in response to (a) anti-CD3 stimulation and (b) phorbol 12-myristate 13-acetate (PMA)/ionomycin stimulation was measured by flow ...

When stimulating with PMA/ionomycin, a poor proliferative response was observed in SF T cells compared with RA PB T cells (Fig. 3b). No statistically significant difference in the proliferative response to PMA/ionomycin stimulation could be detected between RA PB T cells and HC PB T cells, although a trend towards a poor response of RA PB was noted in the CD8+ T cell population.

Analysis of cytokine secretion

The kinetics of in vitro cytokine production of PMA/ionomycin or anti-CD3-stimulated HC PBMC was analysed in pilot experiments and determined to be similar to previously published kinetics of PHA stimulation of PBMC [31]. Results are presented for immunoassay analyses of supernatants after 24 h stimulation (IL-4) or at day 3 (IFN-γ, IL-2 and IL-10) (Fig. 4). Analysis of IL-2 secretion after 24 h of stimulation yielded similar results as when analysing day 3. IL-4 secretion could not be detected in supernatants from cells stimulated for 3 days, however.

Fig. 4
Cytokine concentration in supernatants after stimulation of peripheral blood mononuclear cells (PBMC) or synovial fluid mononuclear cells (SFMC) with (a) anti-CD3 or (b) phorbol 12-myristate 13-acetate (PMA)/ionomycin. Cells were incubated for 24 h (IL-4) ...

When stimulating with anti-CD3, RA SFMC produced significantly less IFN-γ and IL-4 compared with RA PBMC, but no statistically significant difference could be deduced in the IL-10 response (Fig. 4a). IL-2 secretion after CD3 stimulation could not be detected by the immunoassay used. No differences were recorded between RA PBMC and PBMC from HC as to the production of IFN-γ or IL-10 after stimulation with anti-CD3, whereas an increased production of IL-4 was observed in RA PBMC compared with PBMC from HC.

The RA PBMC IFN-γ response to PMA/ionomycin stimulation was markedly depressed compared both with RA SFMC and with HC PBMC (Fig. 4b). The same tendency also held true for IL-10 (although not statistically different), whereas the IL-2 response, and to some degree also the IL-4 response, were lower in SFMC compared with PBMC.

Identification of IFN-γ-producing cells

In order to define the phenotype of the responding cells after stimulation with PMA/ionomycin, intracellular staining of IFN-γ was performed in combination with analyses of cell surface markers (Fig. 5). Since IFN-γ is mainly produced by T cells and natural killer (NK) cells, IFN-γ+ cells were counterstained with a T cell-specific antibody (CD3) and a marker for NK cells (CD94). The proportion of cells producing IFN-γ after stimulation was higher in RA SF T cells compared both with RA PB T cells and with HC T cells (Fig. 6a). However, this was not observed among T cells expressing CD94 or in the NK cell population. Comparing the responsiveness of PB T cells between RA patients and HC demonstrated a lower percentage of IFN-γ+ cells in RA PB, supporting our data of IFN-γ secretion as measured by ELISA. It is evident from Fig. 6b that PMA/ionomycin-induced IFN-γ was mainly derived from T cells (both CD4+ and CD8+) in both SFMC and PBMC. Interestingly, some of the IFN-γ+ T cells expressed CD94 (29% in HC PBMC, 20% in RA PBMC and 6% in SFMC).

Fig. 5
Flow cytometric analysis of IFN-γ-producing cells in RA peripheral blood mononuclear cells (PBMC) and synovial fluid mononuclear cells (SFMC). Cells were stimulated with or without phorbol 12-myristate 13-acetate (PMA)/ionomycin overnight in the ...
Fig. 6
Identification of IFN-γ-producing cells after phorbol 12-myristate 13-acetate (PMA)/ionomycin stimulation. RA peripheral blood mononuclear cells (PBMC; n = 5), synovial fluid mononuclear cells (SFMC; n = 6) and healthy control (HC) PBMC (n = 5) ...

DISCUSSION

The results of our study clearly demonstrate a low expression of CD3ζ in all SF T cells, while there was no difference in CD3ζ expression in PB T cells between RA patients and HC. This is in agreement with Maurice et al. [8] but partly contradicts the study by Matsuda et al. [11], which reported a low expression of CD3ζ in RA PBMC compared with HC PBMC. In the study by Maurice et al. the results obtained by flow cytometry were confirmed by Western blotting using a different antibody. We also demonstrate that granulocytes, but not SF macrophages, have the capacity to down-regulate the expression of CD3ζ on T cells in vitro, and that this down-regulation affects the ability to respond by proliferation and IFN-γ secretion upon stimulation with anti-CD3. We have in preliminary studies observed that this effect of granulocytes on TCR CD3ζ chain expression is cell contact-dependent and that incubation of PBMC with SF does not cause a down-regulation of CD3ζ (data not shown). Granulocytes are abundant in SF and it is plausible that granulocytes contributing to the local inflammation down-regulate the expression of CD3ζ on T cells in vivo. Other reports have identified activated macrophages as potent down-regulators of the expression of CD3ζ via their production of free oxygen radicals [16] or via cell contact-dependent interaction [15], but although macrophages derived from SF have been described as activated [32,33], we could not detect any down-regulatory ability of SF macrophages. Neither have we been able to inhibit the in vitro granulocyte-mediated down-regulation of CD3ζ using the anti-oxidants catalase or N-acetylcysteine (data not shown), suggesting that mechanisms other than secretion of oxygen radicals play a major role in down-regulation of CD3ζ.

Analysis of the ability of SFMC to respond to in vitro stimulation demonstrates that although the proliferation and secretion of T cell-derived cytokines upon stimulation with anti-CD3 are poor in SFMC, in agreement with the low CD3ζ expression of these cells, the proliferative response and induction of IL-2 are also poor when stimulating with PMA/ionomycin. This implies that the hyporesponsiveness of SFMC might in part be due to a low CD3ζ expression, but that pathways downstream of TCR complex signalling must also be involved. Since stimulation with ionomycin induces influx of Ca2+ and PMA activates the calcium-dependent protein kinase C (PKC), we conclude that either these signal transduction mechanisms or events further downstream are affected in the SF T cell population.

Although our results for proliferative responses and cytokine induction after anti-CD3 stimulation clearly indicate a hyporesponsiveness of T cells in the rheumatic joint, it is also evident from our data that direct stimulation of PKC in combination with increase of intracellular Ca2+ in SFMC induces IFN-γ much more efficiently than in paired samples from RA PBMC, and is at least as efficient as in HC PBMC. Hypothetically, these apparently contradictory findings could be explained by a significant expression of IFN-γ by activated non-T cells in SF (i.e. NK cells). However, intracellular staining of IFN-γ combined with cell surface markers clearly demonstrated that IFN-γ is predominantly produced by T cells in both SF and PB, and that PMA/ionomycin induces a larger number of IFN-γ+ T cells in SF compared with both RA and HC PB T cells. The responding T cells were equally distributed among the CD4 and CD8 subsets in both compartments, and interestingly, a small but significant fraction of the IFN-γ-producing cells were identified as NK T cells (CD3+CD94+).

Altogether, our T cell responsiveness data indicate that T cells in SF are only partially hyporesponsive. Since activation bypassing the TCR of SF T cells induces high levels of IFN-γ but not IL-4, such stimulation primarily promotes a cell-mediated immune response (type 1) rather than a humoral immune response (type 2). These data concord with a recent report of PMA + ionophore stimulation of synovial tissue CD4+ cells and PB CD4+ cells [34]. The ability of SF T cells to produce IFN-γ is further supported by our previous studies demonstrating that SFMC of RA patients have an increased expression of IFN-γ mRNA [35] and a higher proportion of cells spontaneously secreting IFN-γ ex vivo[36] compared with PBMC. Since IL-4 was not detected in SFMC, the data imply a shift in the cytokine balance towards a type-1 profile. However, we and others have demonstrated high levels of IL-10 in SF by measurement of mRNA expression in SF cells [35] and secretion of the protein [37,38], indicating that the proinflammatory type-1 response is modulated by the immunosuppressive cytokine IL-10. In the present study, a tendency of increased production of IL-10 was observed in RA SFMC compared with patient PBMC when stimulating with PMA/ionomycin. We can therefore not exclude that mechanisms inducing IFN-γ in SF also activate production of IL-10, although the level of expression may not be sufficient for down-modulation of the immune activation. One plausible explanation for the differential responsiveness of SF T cells is that the ability to produce IFN-γ is augmented by activation via accessory cells. It is well documented that granulocytes and monocytes/macrophages in the synovial compartment are preactivated in vivo[32,3942]. Thus, it is possible that accessory cells in SF have a high capacity to activate T cells to produce IFN-γ by pathways bypassing the TCR, thereby being able to partly overcome the relative hyporesponsiveness of the SF T cell population.

Comparison of the T cell responses in PBMC between RA patients and HC revealed a suppressed induction of, and fewer cells producing, IFN-γ in RA patients when stimulating with PMA/ionomycin. This observation cannot be explained by a general hyporesponsiveness of RA PB T cells since: (i) the PMA/ionomycin-induced secretion of the other T cell-derived cytokines and the proliferative response were similar in the two groups, and (ii) direct activation of T cells by anti-CD3 did not result in depressed IFN-γ production. Similarly, the decreased ability to secrete IFN-γ cannot be explained by a general defective function of accessory cells in RA PB, since these cells spontaneously secrete TNF-α ex vivo as efficiently as HC PB (data not shown). Furthermore, it has previously been demonstrated that RA PB monocytes are phenotypically more activated compared with HC PB monocytes [43,44]. Hypothetically, our findings could be explained by a specific defect in the interplay between costimulatory cells and T cells in RA PB, resulting in a decreased ability of T cells to produce IFN-γ, although none of the cell populations can be defined as generally hyporesponsive.

In conclusion, we have described in detail a defective function of RA SF T cells when stimulating through the TCR, which in part might be explained by the low expression of CD3ζ on these cells. However, since stimulation with PMA/ionomycin also gave a poor proliferative response, events further downstream in the signalling pathways must also be affected. We also demonstrated that the hyporesponsiveness of SF T cells cannot be generalized to all T cell functions, since PMA/ionomycin-stimulated induction of IFN-γ was not suppressed. Thus, it is evident that T cells in the rheumatoid SF, although defective when stimulated directly via the TCR, are capable of responding to external stimuli by other stimulatory mechanisms bypassing the TCR. We have demonstrated that such activation primarily promotes a type-1 immune response rather than a type-2 response.

Acknowledgments

We thank Assoc. Prof. Robert A Harris for linguistic advice. This work was supported by the Swedish Medical Research Council, the Swedish Rheumatism Association, King Gustav V 80th jubilee fund, Prof. Nanna Svartz fund, Ulla and Gustaf af Ugglas fund and Harald, Greta Jeanson’s fund and Tore Nilson’s foundation. The work was performed at the Department of Medicine, Rheumatology Unit, Karolinska Institute, Stockholm, Sweden.

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