Introduction
T cells are likely to play an important role in the pathogenesis of rheumatoid arthritis (RA) (reviewed in [
1]). In the synovial joint, infiltrating T cells are predominantly of the CD4
+ phenotype and are often found in the proximity of B cells and macrophages. These T cells could either represent cells potentiating the function of infiltrating leukocytes or represent suppressive regulatory T cells. Neither specific autoantigens nor autoreactive T cells have so far been conclusively demonstrated in RA. However, a distinct population of oligoclonally expanded proinflammatory CD4
+ T cells is found with increased frequencies in peripheral blood in RA patients compared with healthy control individuals [
2‐
4]. These cells display a proinflammatory phenotype, are terminally differentiated, express a variety of NK cell-related receptors and lack the co-stimulatory molecule CD28; the cells are therefore often referred to as CD4
+CD28
null T cells [
5,
6].
The presence of these CD4
+CD28
null T cells in peripheral blood has been associated with human cytomegalovirus (HCMV) seropositivity, extra-articular manifestations and cardiovascular disease in RA patients [
7‐
9]. Despite increased frequencies of CD4
+CD28
null T cells in the circulation of RA patients, however, their contribution to erosive disease is still unclear: while studies from Pawlik and colleagues and Goronzy and colleagues found associations between circulating CD4
+CD28
null T cells and erosive disease [
4,
10], Martens and colleagues and Gerli and colleagues did not observe such associations [
3,
9].
We had a unique opportunity to investigate the presence of these CD4+CD28null T cells in the synovial membrane, the synovial fluid and peripheral blood from the same patients in a large cohort of RA patients. The association with erosive disease and the levels of antibodies to citrullinated peptides/antigens was examined. Furthermore, CD4+CD28null T cells isolated from the synovial fluid were investigated with regard to antigen specificity and selective recruitment to the joint.
Materials and methods
Patients
One hundred and twenty-eight patients with RA were enrolled in the study. All fulfilled the American College of Rheumatology criteria for RA and attended the Rheumatology Clinic at Karolinska University Hospital, Stockholm, Sweden for corticosteroid injections of inflamed joints [
11]. Before the corticosteroid injections, synovial fluids were acquired from the knee joints (
n = 128), the elbow (
n = 1) or the shoulder joints (
n = 2). Eighty per cent of the patients were women, median age of 56 years (range, 25–82 years) and a median disease duration of 9 years (range, 0–45 years).
Assessment of erosive disease was performed by radiographic evaluations of the ankle joints or wrist joints by the same two rheumatologists. Radiographic changes in one or more joints were found in 51 out of 70 (73%) patients included in these analyses. The majority of the patients were treated either with nonsteroidal anti-inflammatory drugs, with systemic or local corticosteroid treatment, with methotrexate alone or in combination with corticosteroids (prednisolone), or with TNF blockers alone or in combination with methotrexate. Some patients were untreated.
This study was approved in compliance with the Helsinki Declaration by the Ethics Committee of the Karolinska University Hospital, and all patients and healthy subjects gave informed consent.
Arthroscopy and synovial biopsies
Knee joint synovial biopsies were acquired according to a previously described procedure [
12]. Biopsies were taken at the site of inflammation, either close to cartilage or not close to cartilage, defined as either less than 1.5 cm or more than 1.5 cm from cartilage, respectively.
Three-colour immunofluorescence microscopy
Frozen unfixed synovial biopsy sections were fixed with acetone. Sections were incubated overnight with the cocktail of primary antibodies – CD244 (R&D Systems, Minneapolis, MN, USA), CD4 (Becton Dickinson, San Jose, CA, USA), CD3 (DakoCytomation, Glostrup, Denmark) – or the isotype control antibodies – goat IgG (Caltag Laboratories, Burlingame, CA, USA), mouse IgG1 (DakoCytomation) and rabbit immunoglobulin (DakoCytomation). Excess of antibodies were washed away before incubation with the secondary antibodies – anti-sheep/goat immunoglobulin-biotin (The Bidning Site, Birmingham, UK), avidin-Oregon Green 488 (Molecular Probes, Eugene, OR, USA), anti-mouse IgG-Rhodamine RedTM-X (Jackson ImmunoResearch Laboratories, West Grove, PA, USA) and anti-rabbit IgG-AMCA (Jackson ImmunoResearch).
Stained tissue sections were examined with a Leica DM RXA2 microscope (Leica Microsystems, Wetzlar, Germany) equipped with a Leica DC 300F (Leica Microsystems DI, Cambridge, UK) digital colour video camera connected to a PC computer. Photographs were analysed with Leica IM500 software (Leica Microsystems, Heerbrugg, Switzerland).
CD4
+CD28
null T cells were identified by morphologically cell-like structures with co-localized immunostainings of CD3, CD4 and CD244, and were manually quantified in independent analyses performed by two persons. CD4
dim macrophages/monocytes and NK cells [
13], which also might express CD244, could be excluded using the combination of CD3 and CD4 in the three-colour stainings to identify CD4
+ T cells. The density of CD4
+CD28
null T cells were calculated by dividing the number of CD4
+CD28
null T cells by the total area of infiltrating T cells, measured with Image J software version 1.34s (National Institutes of Health, Bethesda, MD, USA).
Flow cytometry
The frequency of CD4+CD28null T cells in peripheral blood and synovial fluid was analysed by four-colour flow cytometry (FACSCalibur instrument; Becton Dickinson Immunocytometry Systems, San Jose, CA, USA) in peripheral blood mononuclear cells and synovial fluid mononuclear cells after Ficoll separation (Ficoll-Paque Plus; GE Healthcare Biosciences AB, Uppsala, Sweden). The antibodies used were CD3-FITC, CD28-APC (Pharmingen; Becton Dickinson, San Diego, CA, USA), CD4-PerCP (Becton Dickinson, San Jose) and CD244-PE (Immunotech, Marseille, France).
The TCR-Vβ usage was determined by the IOTest1 Beta Mark kit (Beckman Coulter, Marseille, France). The TCR-Vβ stainings were combined with antibodies to CD4 and CD28 (see above) to identify CD4+CD28null T cells and CD4+CD28+ T cells.
Peripheral blood mononuclear cells and synovial fluid mononuclear cells stimulated with HCMV antigens (see below) were analysed by flow cytometry after immunostaining with IFNγ-FITC, CD28-PE, CD3-APC and CD14-APC-Cy7 (all from Becton Dickinson, San Diego, CA, USA) and CD4-PerCp (Becton Dickinson, San Jose, CA, USA).
Flow cytometric data were analysed with CellQuest software (Becton Dickinson, Franklin Lakes, NJ, USA) or FlowJo software (Tree Star Inc., Ashland, OR, USA). The frequency of CD4+CD28null T cells was calculated as the percentage of CD28-negative cells in the gated CD3+CD4+ population.
Functional assays
The functional capacity of CD4+CD28null T cells from the synovial fluid and peripheral blood were assessed by IFN-γ production. Two million peripheral blood mononuclear cells and synovial fluid mononuclear cells from eight patients were either stimulated with plate-bound anti-CD3 antibodies (OKT-3) at 0 or 0.1 μg/ml for 4 hours, or by 2 μg/ml pp65 and immediate early HCMV antigens (JPT Peptide Technologies GmbH, Berlin, Germany) for 8 hours.
Activated cells were either detected by secretion (MACS Secretion Assay; Miltenyi Biotec, Bergisch Gladbach, Germany) or by upregulation of intracellularly stored IFN-γ (Becton Dickinson, San Diego, CA, USA) according to the manufacturers' protocols. The frequency of IFN-γ secreting CD3+CD4+CD28null T cells was analysed by flow cytometry (see above).
Enzyme-linked immunosorbent assay
The anti-CCP2 test (Immunoscan RA, Mark 2; Euro-Diagnostica, Arnhem, The Netherlands) was used to determine the levels of anti-citrullinated peptide/protein antibodies (ACPA) in the serum and synovial fluid. A cutoff value of 25 U/ml was used according to the manufacturer's instructions. Serum and synovial fluid samples were diluted equally (1:50) and were analysed on the same plate.
The presence of anti-HCMV IgG and IgM antibodies in the serum and the synovial fluid, from the same time point as the screening of CD4
+CD28
null T cells, was tested in an enzygnost anti-HCMV/IgG ELISA and an enzygnost anti-HCMV/IgM ELISA (Dade Behring, Marburg, Germany). Sera from HCMV-seronegative patients were further examined for detection of IgG against HCMV using antigens prepared from a HCMV clinical isolate (C6) using an ELISA as previously described by Rahbar and colleagues [
14]. Control antigen was prepared from uninfected fibroblasts.
Statistical analyses
Comparisons of nonparametrically distributed data in two independent groups or compartments were performed by the Mann–Whitney test. The Spearman test for correlation was used for analyses of covariation of two nonparametrically distributed data.
Discussion
Herein we demonstrate that only minor populations of CD4+CD28null T cells were present in the inflamed joints of RA patients, despite significant percentages in peripheral blood. The presence of CD4+CD28null T cells in peripheral blood and the synovial fluid was strongly associated with HCMV IgG seropositivity, but not with ACPA or erosive disease.
Our results on the presence of CD4
+CD28
null T cells in the synovial fluid were based on screening for CD3
+CD4
+CD28
- cells. It was therefore important to consider the stability of the CD28-negative phenotype. Data from
in vitro experiments indicate that cytokines such as TNF and IL-12 in synovial fluid can modify CD28 expression, complicating the analyses of CD4
+CD28
null T cells in the joint [
19,
20]. We believe our data, however, not to be biased by the cytokines present in the synovial fluid since there was no over-representation of synovial CD4
+CD28
+ T cells expressing the TCR-Vβ chains preferentially expressed by peripheral blood CD4
+CD28
null T cells (Figure
4). Interestingly, previous studies have shown a reduction in the frequency of CD4
+CD28
null T cells in peripheral blood after TNF blockade [
9,
21‐
23]. In our cohort, neither TNF blockade nor any of the other most frequently used medical treatments (untreated, nonsteroidal anti-inflammatory drug, methotrexate) was associated with the distribution of CD4
+CD28
null T cells in the synovial fluid, peripheral blood and synovial tissue. It is therefore likely that the effect of TNF blockade on CD4
+CD28
null T cells is primarily seen when comparing the same patients before and after treatment, rather than comparing heterogeneous groups of patients with and without this treatment.
CD4
+CD28
null T cells from both peripheral blood and synovial fluid demonstrated reactivity to HCMV-derived antigens. That the frequency of responding CD4
+CD28
null T cells was higher in the synovial fluid compared with the peripheral blood does not necessarily mirror an accumulation of HCMV-reactive CD4
+CD28
null T cells in this compartment, since the same effect was seen for CD4
+CD28
+ T cells (data not shown). These differences might instead be due to a different status of accessory cells from the two compartments. We also analysed the HCMV specificity of the dominant TCR-Vβ subsets of CD4
+CD28
null T cells from two patients comprising 20% and 29% of the total CD4
+CD28
null population in both peripheral blood and synovial fluid. Interestingly, the TCR-Vβ dominant CD4
+CD28
null T-cell subsets did not respond either to pp65 or to immediate early antigens, indicating that the TCR-Vβ dominant subsets might be reactive to antigens other than those considered immunodominant for HCMV (data not shown). This might be explained by a hypothesis suggested by Davenport and colleagues, who demonstrate that during chronic Esptein–Barr virus infections T-cell clones reactive to the most dominant epitopes rapidly decrease after primary infection and that clonotypes reactive to less dominant epitopes control the recurrent infections [
24]. At present, however, we can not exclude that some of the CD4
+CD28
null T cells with access to the joint have specificity for cartilage-derived and/or citrullinated candidate antigens.
It is intriguing that only certain subsets of CD4
+CD28
null T cells reach the synovial fluid. Since we were not able to detect all possible TCR-Vβ chains, we cannot exclude that there is an even more pronounced discrimination of CD4
+CD28
null T cells expressing nondetectable TCR-Vβ chains. The reason for this skewed distribution of CD4
+CD28
null T cells in peripheral blood and synovial fluid, both with regard to subsets of CD4
+CD28
null T cells and to the size of whole populations, can with present knowledge only be speculated upon. Owing to the strong association of CD4
+CD28
null T cells to HCMV seropositivity, it is tempting to assume that the location or status of the HCMV infection plays an important role. Because of the increased frequencies in peripheral blood and exclusion from the synovial fluid, it is probable that tissues other than the rheumatic joint are the primary homing sites for CD4
+CD28
null T cells in these patients. The few CD4
+CD28
null T cells found in the joint could instead be a consequence of general patrolling initiated by infection in other tissues. A widespread distribution of virus-specific T cells in a site other than the actual site of virus infection has previously been demonstrated in mouse models [
25]. Further investigations considering the expression of chemokine receptors/integrins, antigen specificity, location of the HCMV infection and the presentation of HCMV antigens are needed to clarify this issue. The frequency of CD4
+CD28
null T cells in the synovial fluid does not necessarily reflect the situation in the inflamed synovia, although in our cohort the low CD4
+CD28
null T-cell frequencies in the synovial fluid were in agreement with the modest numbers in the inflamed synovial membrane.
CD4
+CD28
null T cells isolated from the synovial fluid could function as effector T cells by rapid secretion of IFN-γ. Interestingly, IFN-γ is only scarcely found in the T-cell infiltrates of the rheumatic synovial membrane and has therefore not been considered a key cytokine in the pathogenesis of RA [
26]. Several reports instead indicate the importance of IL-17, and recent publications have further shown that IFN-γ counteracts the differentiation of IL-17-producing T cells [
27‐
29]. Since CD4
+CD28
null T cells produce IFN-γ but not IL-17 [
6,
30], it is possible that CD4
+CD28
null T cells, if activated in the joint by secretion of IFN-γ, might even inhibit the synovial inflammation in RA.
The limited presence of CD4
+CD28
null T cells in the synovial fluid, despite increased frequencies in peripheral blood and their equal distribution in patients with and patients without erosive disease, indicates no significant role for CD4
+CD28
null T cells in the local inflammation driving joint destruction. Instead, these data indirectly support the previously suggested role for these cells in extra-articular manifestations and cardiovascular disease. That is, CD4
+CD28
null T cells are exclusively present in HCMV IgG-seropositive RA patients and are reactive to HCMV antigens (present study and [
7]), CD4
+CD28
null T cells only have limited access to the inflamed joint despite increased frequencies in the circulation (present study), increased frequencies of CD4
+CD28
null T cells do not correlate with erosive disease (present study and [
3,
9]), RA patients with high frequencies of circulating CD4
+CD28
null T cells display increased risk for cardiovascular events compared with patients lacking CD4
+CD28
null T cells [
9,
31], CD4
+CD28
null T cells have been found in atherosclerotic plaques and can mediate lysis of endothelial cells
in vitro [
32,
33], HCMV is frequently found in atherosclerotic and nonatherosclerotic vascular walls [
34], and HCMV increases the thrombogenicity of endothelial cells [
35].
CD4+CD28null T cells and HCMV might not be the only mediators of cardiovascular events in RA, but these studies together strongly link CD4+CD28null T cells and HCMV infection to cardiovascular events, which is found with increased prevalence and is the major cause of death in patients with RA.
Acknowledgements
The authors would like to thank Dr Cecilia Söderberg-Naucler for helpful discussion in HCMV-related issues, Dr Florian Kern and Charlotte Tammik for providing the protocol and reagents for the HCMV reactivity assay, Eva Jemseby for organizing sampling, storage and administration of biomaterial, and Marianne Engström for technical support with the immunofluorescence microscopy stainings. This study was supported by Alex and Eva Wallstrom, Borje Dahlin, Tore Nilsson, Magn. Bergvall, Nanna Svartz, and Åke Wiberg Foundations, the Swedish Association againt Rheumatism, the Swedish Medical Association, the King Gustaf the V:s 80 year Foundation, the Swedish Research Council and the EU FP6 project, AutoCure LSHB CT-2006-018661, 2. This publication reflects only the author's views; the European Community is not liable for any use that may be made of the information herein.
Competing interests
The authors declare that they have no competing interests.
Authors' contributions
AERF was responsible for the study design, performed laboratory work (preparation of blood and synovial fluid samples, flow cytometry analyses, development, analyses of the immunofluorescence microscopy, and HCMV reactivity assays), statistical analyses, interpretation of the data, and drafted the manuscript. OS performed and interpreted data from in vitro stimulation assays as well as ACPA ELISAs of serum and synovial fluid. AATJ performed and analysed the immunofluorescence stainings. BN contributed to the clinical evaluation of the patient cohort together with EaK, who also performed the arthroscopies. AR carried out and evaluated the results from the HCMV ELISAs. NKB set up the flow cytometric assays for investigation of the CD244 expression. A-KU was responsible for the biobank of arthroscopic biopsies and participated in the development of immunofluorescence stainings. RvV contributed to the clinical evaluations of patients and manuscript preparation. VM and CT were the principle investigators and participated equally in the planning and coordination of the study, interpretation of data, and drafting the manuscript. All authors read and approved the final manuscript.