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Abstract

  1. Top of page
  2. Abstract
  3. PATIENTS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. REFERENCES

Objective

To evaluate the possible role of activation-induced, T cell–derived, and chemokine-related cytokine (ATAC)/lymphotactin (Lptn) in the pathogenesis of rheumatoid arthritis (RA).

Methods

ATAC/Lptn levels in serum and synovial fluid samples were measured by sandwich enzyme-linked immunosorbent assay. Expression of messenger RNA for ATAC/Lptn in synovial tissues was analyzed by reverse transcription–polymerase chain reaction (PCR) and by in situ hybridization, and was quantitated by real-time PCR. The phenotype of peripheral blood mononuclear cells (PBMCs) expressing ATAC/Lptn was analyzed by intracellular cytokine staining and flow cytometry.

Results

Levels of ATAC/Lptn were similar in sera and synovial fluids from RA patients (n = 20) and osteoarthritis controls (n = 15). In phorbol myristate acetate/ionomycin–stimulated PBMCs, ATAC/Lptn expression was detected in CD8+ T cells and in a significantly increased proportion of CD4+,CD28− T cells from RA patients as compared with healthy controls. In synovial tissues, ATAC/Lptn was predominantly localized in CD3+ T cells in the sublining layer. Lymphocytes, synovial macrophages, and, unexpectedly, fibroblast-like synoviocytes (FLS) were identified as major target cells for ATAC/Lptn in RA synovium, as determined by analysis of the ATAC/Lptn receptor XCR1. In vitro, ATAC/Lptn stimulation of FLS resulted in a marked down-regulation of matrix metalloproteinase 2 production.

Conclusion

These data indicate that in RA synovium, ATAC/Lptn is mainly produced by T cells. Considering its function as a lymphocyte-specific chemoattractant, ATAC/Lptn might be a key modulator for T cell trafficking in the pathogenesis of RA. In addition, functional studies suggest that ATAC/Lptn may exert additional immunomodulatory effects in RA.

Rheumatoid arthritis (RA) is a chronic inflammatory disease of still-unknown etiology that leads to progressive destruction of affected joints by extracellular matrix degradation. The rheumatoid synovium is characterized by marked hyperplasia of synovial lining cells and prominent T lymphocyte infiltration (1). Fibroblast-like synoviocytes (FLS) and synovial macrophages have previously been shown to promote cartilage destruction and bone resorption by the production of matrix metalloproteinases (MMPs) (2), which are a family of zinc-dependent endopeptidases, and proinflammatory cytokines (3), such as interleukin-1β (IL-1β) and tumor necrosis factor α (TNFα). The contribution of T cells to the chronic inflammatory process in RA, however, remains to be defined. Association of the disease with HLA–DR4 antigens (4) and prominent accumulation of T cells in RA synovium (5) support a central role for these cells in the pathogenesis of RA. Until now, the mechanisms promoting T cell recruitment to sites of chronic inflammation have not been fully elucidated. T cell trafficking has previously been characterized as a multistep process involving selectins, adhesion molecules, and chemokines (6, 7).

Chemokines are secreted, low molecular weight proteins known to play a crucial role in transendothelial leukocyte migration and activation along chemoattractant gradients during inflammation (8). These molecules are structurally related and have previously been classified according to the organization of the N-terminal conserved cysteine motif into 4 groups, designated as chemokines CC, CXC, CX3C, and C (9). Whereas the CXC and CC groups both include several members, the C and CX3C chemokine subfamilies are represented so far by only 1 member each: lymphotactin (Lptn; XCL1) and fractalkine (or, neurotactin; CX3CL1), respectively. The 4 groups can also be distinguished according to their chromosomal localization and their biologic activities: CXC chemokines mainly target neutrophils and T cells, while CC chemokines generally attract monocytes and T cells. In contrast, both C and CX3C chemokines have a more restricted specificity for T cells.

The Lptn molecule was independently detected by Kelner et al (10), who called it lymphotactin, Yoshida et al (11), who called it single C motif 1 (SCM-1), and Müller et al (12), who called it activation-induced, T cell–derived, and chemokine-related molecule (ATAC). This novel chemokine is structurally related to the CC chemokine subfamily that lacks the first and third cysteine residues, and it is thus considered to represent the C chemokine subfamily. Two highly homologous genes encoding for XCL1 and XCL2, respectively, have thus far been detected in humans. ATAC/Lptn was found to be selectively expressed in activated CD8+ T cells and in a small proportion of activated CD4+ T cells (12), α/β-type thymocytes (10), intraepithelial γ/δ-type T cells (13), mast cells (14), and natural killer (NK) cells (15). ATAC/Lptn acts via a unique G protein–coupled receptor (XCR1) (16, 17).

The full spectrum of biologic functions of ATAC/Lptn is still unknown, but the functional properties that have been detected include chemotactic activity for both CD4+ and CD8+ T cells (18, 19) and induction of migratory responses in NK cells after activation with IL-2 (20). The ability of ATAC/Lptn to specifically recruit T cells and NK cells has been used successfully in gene therapy. Genetic modification of tumor RNA–pulsed dendritic cells to secrete ATAC/Lptn was shown to be associated with an enhanced therapeutic efficacy of dendritic cell–based tumor vaccines (21). Moreover, results of recent studies in a murine model of listeriosis suggest that ATAC/Lptn, macrophage inflammatory protein 1α (MIP-1α; CCL3), MIP-1β (CCL4), and RANTES (CCL5) are cosecreted with interferon-γ (IFNγ) by activated NK cells, CD8+ T cells, and CD4+ Th1 cells, and function as type 1 cytokines by up-regulating CD40, IL-12, and TNFα in macrophages (22).

Expression of ATAC/Lptn has been studied in several clinical and experimental models of inflammatory disease, such as acute allograft rejection (23, 24), autoimmune diabetes (25), encephalomyelitis (26), experimental crescentic glomerulonephritis (27), and chronic inflammatory bowel disease (28). Results of these studies support the concept of a potential role of ATAC/Lptn in Th1-type inflammatory processes. To analyze whether ATAC/Lptn may also participate in the pathogenesis of RA, we investigated the expression of ATAC/Lptn and its receptor in synovial tissues from RA patients and compared it with the expression in synovial tissues from osteoarthritis (OA) control patients. Our findings indicate that there is enhanced expression of ATAC/Lptn in rheumatoid synovium, on both CD4+ and CD8+ T cells. In addition, this study is the first to demonstrate the expression of the ATAC/Lptn receptor (XCR1) on FLS. Functional analysis revealed a marked down-regulation of MMP-2 production in cultured FLS during in vitro stimulation with ATAC/Lptn. These findings suggest a novel immunoregulatory function for ATAC/Lptn in RA.

PATIENTS AND METHODS

  1. Top of page
  2. Abstract
  3. PATIENTS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. REFERENCES

Serum, synovial fluid, and synovial tissue specimens.

Serum samples (10 ml) were obtained from 40 RA patients and 20 healthy blood donors. Synovial fluid specimens were collected from 15 RA patients and 15 OA patients who underwent diagnostic or therapeutic joint puncture. For flow cytometry of peripheral blood mononuclear cells (PBMCs), blood samples (20 ml) from 10 RA patients and 3 healthy controls were drawn into tubes containing heparin. Synovial tissue specimens were obtained at the time of surgery from 20 RA patients who were undergoing synovectomy or total joint replacement.

Synovial tissue specimens were also obtained at the time of arthroplasty from 5 patients with psoriatic arthritis, 5 patients with reactive arthritis, and 15 age- and sex-matched control patients with OA. The diagnosis of OA was based on clinical and radiologic findings, rheumatoid factor negativity, and absence of acute-phase reactants. Baseline clinical characteristics of the RA patients from whom synovial tissues were obtained and the OA controls are summarized in Table 1.

Table 1. Baseline characteristics of RA patients from whom synovial tissues were obtained and OA control patients*
PatientsRA (n = 20)OA (n = 15)
  • *

    RA = rheumatoid arthritis; OA = osteoarthritis; RF = rheumatoid factor; DAS = Disease Activity Score; ESR = erythrocyte sedimentation rate; CRP = C-reactive protein; NSAIDs = nonsteroidal antiinflammatory drugs; DMARDs = disease-modifying antirheumatic drugs.

Sex, no. of males/females8/125/10
Age, mean (range) years61.6 (31–80)68.1 (42–87)
IgM-RF, % positive76
DAS, mean (range)3.64 (2.94–5.19)
No. of swollen joints, mean (range)5.32 (2–12)
ESR, mean (range) mm/hour35 (6–89)
CRP, mean (range) mg/liter28.1 (2.2–103)
Disease duration, median (range) years13.4 (1–56)
Therapy
 NSAIDs18
 Corticosteroids19
 DMARDs18
 Arthroplasty1115
 Synovectomy9

All patients with RA met the criteria established by the American College of Rheumatology (formerly, the American Rheumatism Association) (29). Their mean disease duration was 13.4 years. Most of the patients (18 of 20 [90%]) were rheumatoid factor positive and were receiving standard disease-modifying antirheumatic drug therapy (methotrexate or sulfasalazine) and low-dose prednisone (<10 mg/day). All patients had active disease, defined as the presence of at least 3 swollen and tender peripheral joints and morning stiffness of >1 hour's duration, with or without an elevated erythrocyte sedimentation rate (>28 mm/hour) or C-reactive protein level (>8 mg/liter). Clinical disease activity was assessed according to clinical and laboratory findings, as defined by van der Heijde et al (30).

Synovial tissue specimens were immediately split in half for cryopreservation and paraffin embedding. Samples were snap-frozen in liquid nitrogen for subsequent isolation of RNA and reverse transcription–polymerase chain reaction (RT-PCR) analysis. Paraffin-embedded tissue was subjected to histologic analysis, immunohistochemistry, and in situ hybridization studies.

The study was approved by the local Institutional Review Board.

Enzyme-linked immunosorbent assay (ELISA) for ATAC/Lptn.

For measurement of ATAC/Lptn levels in serum and synovial fluid specimens, a sandwich ELISA was established. Microtiter plates (96-well) were precoated overnight at 4°C with 100 μl of a polyclonal goat anti-human Lptn antibody (R&D Systems, Wiesbaden, Germany) diluted 1:50 in 50 mM carbonate buffer (pH 9.6). Plates were washed 4 times with 200 μl of washing buffer (phosphate buffered saline [PBS]–0.05% Tween 20) per well and were subsequently blocked with blocking buffer (PBS–1% bovine serum albumin [BSA]–5% sucrose) for 1 hour at room temperature. Microtiter plates were washed, and 100 μl of the samples (diluted 1:10 in dilution buffer containing PBS–0.05% Tween 20–5% nonfat dry milk) and standards was applied to each well. Wells were incubated for 2 hours at room temperature. A 7-point standard curve was generated by 2-fold serial dilutions of recombinant human Lptn (R&D Systems). Plates were washed 4 times and incubated for 1 hour with a rabbit anti-human Lptn antibody (Serotec, Düsseldorf, Germany) diluted 1:200 in dilution buffer.

Specifically bound antibody was detected by application of horseradish peroxidase–labeled polymer conjugated to secondary goat anti-rabbit antibody (EnVision rabbit; Dako, Hamburg, Germany) diluted 1:50 in dilution buffer for 1 hour at room temperature. Peroxidase activity was visualized by the addition of 100 μl of o-phenylenediamine substrate solution (Dako). After 30 minutes of incubation, 50 μl of stop solution (1N H2SO4) was added to each well, and the plates were read at 490 nm with a Tecan ELISA reader (Tecan, Männedorf, Switzerland). The detection limit of the assay was 0.4 ng/ml.

Flow cytometry of PBMCs.

PBMCs were isolated from heparinized blood samples by Ficoll-Hypaque density-gradient centrifugation (Biochrom, Berlin, Germany). Cells were resuspended at a density of 2 × 106/ml in Iscove's modified Dulbecco's medium (Gibco BRL, Eggenstein, Germany) supplemented with 100 IU/ml of penicillin, 100 μg/ml of streptomycin, and 10% fetal calf serum (FCS). Cells were then stimulated with phorbol myristate acetate (PMA; 20 ng/ml) (Sigma, Deisenhofen, Germany) and ionomycin (1 μg/ml; Sigma). After 2 hours of incubation, brefeldin A (5 μg/ml; Sigma) was added to inhibit chemokine release. Unstimulated PBMCs were cultured only in the presence of brefeldin A (5 μg/ml). After 5 hours of incubation, both stimulated and unstimulated cells were collected, centrifuged at 550g for 5 minutes at 4°C, and washed in PBS–0.1% BSA.

PBMCs (2 × 105) were then resuspended in 50 μl of PBS–2.5% FCS–0.1% NaN3 (FACS–PBS) containing 2 mg/ml of Ennobling (Immuno, Heidelberg, Germany) and stained for cell surface antigens with fluorophore-labeled antibodies (30 minutes at 4°C). Monoclonal antibodies (mAb) against the surface antigens CD3 (OKT3) and CD8 (OKT8; both from American Type Culture Collection [ATCC], Rockville, MD), CD4 (clone 91d6) (31), and CD28 (clone hCD28-103; kindly provided by H. W. Mages, Department of Molecular Immunology, Robert-Koch Institute, Berlin, Germany) were coupled to the following fluorophores according to standard procedures: fluorescein-N-hydroxysuccinimide ester (Molecular Probes, Leiden, The Netherlands), phycoerythrin (Cyanotech, Kailua-Kona, Hawaii), Cy5-N-hydroxysuccinimide ester (Pharmacia, Erlangen, Germany), and digoxigenin-N-succinimide ester (Roche, Mannheim, Germany). After washing in FACS–PBS, cells were fixed in 2% paraformaldehyde in PBS for 20 minutes at room temperature and washed again in FACS–PBS.

For intracellular cytokine staining, PBMCs were resuspended in FACS–PBS containing 0.5% saponin (Sigma) and incubated for 30 minutes with a digoxigenin (DIG)–labeled mouse anti-human ATAC/Lptn primary mAb (clone ASA-1; 8 μg/ml) (19). After washing in FACS–PBS, PBMCs were incubated with the secondary reagent, anti–DIG–Cy5, as previously described (32). PBMCs were resuspended in 250 μl of FACS–PBS and subjected to flow cytometry analysis on a 2-laser FACSCalibur cytometer (Becton Dickinson, Heidelberg, Germany). Dead cells and monocytes were excluded by forward and side scatter gating. Typically, 10,000 events were acquired and analyzed using the CellQuest software (Becton Dickinson). Bivariate dot plots were generated upon data reanalysis to display the frequencies of individual cells coexpressing certain levels of cell surface antigens and intracellular ATAC/Lptn. Statistical analysis of intracellular cytokine coexpression was performed as previously described (22).

To reveal the specificity of staining, target cells were incubated with an isotype-matched immunoglobulin of irrelevant specificity at the same concentration for each fluorescence channel. In addition, blocking experiments were performed by incubation of cells with at least a 100-fold excess (usually 100–200 μg/ml) of unlabeled mAb prior to the addition of fluorophore-labeled antibody.

RT-PCR.

ATAC/Lptn messenger RNA (mRNA) expression in synovial tissue samples from 20 RA patients and 15 OA control patients was analyzed by RT-PCR. After total cellular RNA isolation (RNeasy kit, Qiagen, Hilden, Germany) and DNase I digestion (RNase-free DNase I; Pharmacia, Freiburg, Germany), first-strand complementary DNA (cDNA) was synthesized from 1 μg of isolated RNA with 1.5 μM oligo(dT)12-18 primer (Gibco BRL), 5× reverse transcriptase buffer, 0.4 μM of each dNTP (MBI Fermentas, St. Leon-Rot, Germany), and 200 units of reverse transcriptase (Superscript II; Gibco BRL) in a total volume of 50 μl at 42°C for 1 hour. The quality of cDNA was examined by PCR for β-actin as previously described (33). To detect ATAC/Lptn and ATAC/Lptn receptor expression, respectively, 1 μl of cDNA was amplified by PCR in a mixture containing 1 μl of each dNTP (10 mM), 1.5 mM MgCl2, 5 μl of PCR buffer, 0.5 μl of each primer (10 nmoles/μl), and 2.5 units of Taq polymerase (Gibco BRL) in a final volume of 50 μl.

Oligonucleotide primer sequences were designed according to the GenBank sequences for β-actin (accession no. X00351), ATAC/Lptn (accession no. NM002995), and ATAC/Lptn receptor XCR1 (accession no. NM011798). Cycle conditions were as follows: for β-actin forward (5′-CCCAGCCATGTACGTTGCTAT-3′) and β-actin reverse (5′-GGGTGGCTTTTAGGATGGCAA-3′), product size 1,047 bp, 3 minutes at 95°C, 35 cycles of 45 seconds at 95°C, 45 seconds at 60°C, 90 seconds at 72°C, followed by a final elongation step for 10 minutes at 72°C; for ATAC/Lptn forward (5′-GTGGAAGGTGTAGGGAGTGAAGTC-3′) and ATAC/Lptn reverse (5′-GTCCATGAGGGTGTAAAGTGAAAT-3′), product size 354 bp, 3 minutes at 95°C, followed by 35 cycles of 45 seconds at 95°C, 45 seconds at 57°C, 1 minute at 72°C, and a final elongation step for 10 minutes at 72°C; and for ATAC/Lptn receptor XCR1 forward 1 (5′-GCTTTCTTCGGGCTGTGATTATTC-3′) and ATAC/Lptn receptor XCR1 reverse 1 (5′-GCGGCAGATGAGCAGGGCGTATT-3′), product size 310 bp, 3 minutes at 95°C followed by 40 cycles of 45 seconds at 95°C, 45 seconds at 58°C, 1.5 minutes at 72°C, and a final elongation step for 10 minutes at 72°C.

PCR products were visualized on 1.5% agarose gels by staining with ethidium bromide. PCR products were cloned into the pSTBlue1 vector (Novagen, Madison, WI) to generate internal controls. Negative (distilled water) and positive (internal standard) controls were included in each PCR analysis. To control the specificity of the PCR assay, PCR products were sequenced in both directions by cycle sequencing with dye-dideoxy terminator dNTPs on an automated DNA sequencer (Applied Biosystems, Weiterstadt, Germany).

Real-time quantitative PCR.

To quantitate ATAC/Lptn mRNA expression, a real-time quantitative RT-PCR was established using LightCycler (Roche) technology as previously described (34). PCR was performed in a total volume of 20 μl containing 1 μl of cDNA, 2 μl of SYBR Green master mix (Roche), 4 mM MgCl2, and 11.25 pmoles of each oligonucleotide primer. Primer sequences were as follows: for ATAC/Lptn forward primer 5′-GCCGGTTAGCAGAATCAAGA-3′ and for ATAC/Lptn reverse primer, 5′-CTGGCTGGCTGGAGACG-3′, with a product size of 262 bp. Cycle conditions included denaturation of the template DNA for 1 cycle at 95°C for 30 seconds, amplification of the target DNA for 45 cycles at 95°C for 0 seconds, 62°C for 5 seconds, and extension at 72°C for 10 seconds, each with a temperature transition rate of 20°C per second.

Melting curve analysis was performed to determine the melting temperatures of the primer–dimer product and the specific PCR product. Fluorescence was measured in a separate detection step (2 seconds) at a temperature between the melting points of the primer–dimers and the specific PCR product (87°C for β-actin; 83°C for ATAC/Lptn) to exclude fluorescence attributable to the primer–dimers. Each run included external standards as positive controls and water as a negative control. Standards were generated by densitometric quantification of specific PCR products after agarose gel electrophoresis and preparation of a 10-fold dilution series ranging from 100 amoles/μl to 0.0001 amoles/μl. External standards were used to construct a standard curve. The cDNA concentration in each sample was then calculated automatically by reference to the standard curve. To standardize ATAC/Lptn mRNA concentrations, transcript levels of the housekeeping gene β-actin were determined in parallel for each sample. Results were then expressed as the ratio of the molar amounts of ATAC/Lptn mRNA and β-actin mRNA.

Double-label immunohistochemistry.

Double-label immunohistochemistry for ATAC/Lptn and cell surface markers CD3 (clone N/A; Dako), CD4 (clone NCL-CD4-368; Novocastra, Dossenheim, Germany), CD8 (clone NCL-CD8-4B11; Novocastra), tryptase (clone AA1; Dianova, Hamburg, Germany), Fascin (clone 55K-2; Dako), and the NK cell–like antigen (clone NK1; Dako) was performed on paraffin sections of synovial tissue samples as previously described (28). Briefly, the primary antibody, a rabbit anti-human ATAC/Lptn antibody (1:600 dilution; TEBU, Frankfurt, Germany), was applied overnight in a humidified chamber at 4°C and then detected by use of the biotin–tyramide signal amplification system (DuPont NEN, Boston, MA). Paraffin sections were subsequently incubated overnight with antibodies against the cell surface antigens. Specifically bound antibodies were detected by indirect immunofluorescence using rhodamine red-X–conjugated F(ab′)2 fragments of rabbit anti-mouse IgG (1:50; Dianova). Rhodamine red-X–conjugated goat anti-rabbit IgG (1:50; Dianova) was used for detection of the polyclonal anti-CD3 antibody.

Sections were thoroughly washed between each incubation step with Tris buffered saline (TBS; 0.1M Tris HCl, pH 7.5, 0.15M NaCl, pH 7.4) containing 0.05% Tween 20. Negative controls included irrelevant antibodies of the same species as well as omission of the first or second antibody. Sections were finally mounted in fluorescent mounting medium (Dako) and analyzed on a Leica TCS NT laser scanning microscope (Leica, Heidelberg, Germany).

In situ hybridization.

In situ hybridization for the detection of ATAC/Lptn mRNA expression was performed on paraffin sections of synovial tissues according to a modified procedure described by Breitschopf et al (35). Briefly, the PCR product was cloned into the Sma I restriction site of the transcription vector pBluescript (Stratagene, Heidelberg, Germany). To generate DIG-11-UTP–labeled (Roche) antisense (complementary) and sense (anticomplementary) probes, PCR was performed using oligonucleotide primers specific for the T3 and T7 promoter regions of the vector. After in vitro transcription with T3 or T7 RNA polymerase (Roche), the labeling efficiency of probes was controlled by dot-blot analysis of serial probe dilutions.

For in situ hybridization, tissue sections were deparaffinized and permeabilized with proteinase K (10 μg/ml at 37°C for 20 minutes; Sigma). After denaturation of sections (5 minutes at 85°C) on a heating block, the DIG-labeled probe was applied, and hybridization was carried out overnight at 58°C in a sealed, humidified chamber. Posthybridization washes included 1× sodium citrate–sodium chloride (SSC)/0.1% sodium dodecyl sulfate (SDS) (twice for 5 minutes at room temperature) and 0.2× SSC/0.1% SDS at hybridization temperature (twice for 10 minutes). Finally, sections were washed in TBS containing 0.1% Tween 20. Specifically bound probe was detected with a polyclonal sheep anti-DIG antibody conjugated with alkaline phosphatase F(ab′)2 fragments (1:500 dilution; Roche). Enzyme activity was visualized after addition of the BCIP/nitroblue tetrazolium substrate solution (Roche). The sense probe was included in each experiment as a negative control.

To characterize the phenotype of ATAC/Lptn mRNA–positive cells, immunohistochemical staining for cell surface antigens was performed after in situ hybridization, using a modification of the procedure described above. In situ–hybridized sections were incubated with mAb directed against the Ki-M1P antigen (kindly provided by Prof. Dr. R. Parwaresch, Kiel, Germany) (36) or against CD3 (Beckman Coulter, Heidelberg, Germany). Antibodies were detected by indirect immunofluorescence with a fluorescein isothiocyanate–conjugated rabbit anti-mouse IgG (1:50; Dako). Staining for CD4 and CD8 antigens (Novocastra) was performed on serial paraffin sections. Primary antibodies were detected using a biotin–streptavidin amplification system with alkaline phosphatase–conjugated streptavidin and fast red/naphthol phosphate substrate according to the manufacturer's instructions (BioGenex, Hamburg, Germany).

In situ hybridization for the detection of the ATAC/Lptn receptor (XCR1) was performed with a riboprobe generated from a 653-bp PCR product. The primer sequences for the XCR1 PCR were as follows: for XCR1 forward 2, 5′-TGGGGCTGGGTGCTGGGAGACT-3′ and for XCR1 reverse 2, 5′-GCGGCAGATGAGCAGGGCGTATT-3′. PCR was carried out as described above. Cycle conditions were denaturation at 95°C for 3 minutes, followed by 35 cycles of 95°C for 45 seconds, 66°C for 45 seconds, 72°C for 1.5 minutes, and a final elongation step at 72°C for 10 minutes. The PCR product was cloned into the Sma I restriction site of the cloning vector pBluescript SK+ (Stratagene). After in vitro transcription, in situ hybridization was carried out as described above.

Cell culture.

ATAC/Lptn receptor (XCR1) expression was studied in 4 different cell lines and cultured FLS from 3 different donors. The 4 cell lines were 2 renal fibroblast cell lines (TK 173 and TK 188), a T cell acute lymphoblastic leukemia cell line (Jurkat), and a cell line of human immature monocytes (U937). The Jurkat (TIB-152) and U937 (CRL-1593.2) cell lines were purchased from ATCC.

For generation of FLS cultures, synovial tissue specimens were digested with Dispase (Roche) as previously described (37). Briefly, fat and fibrous tissues were removed, and synovium was cut into small pieces of ∼5 mm3. To isolate FLS, the synovial pieces were finely minced and digested for 3–4 hours at 37°C with Dispase in 50% PBS and 50% Dulbecco's modified Eagle's medium (DMEM; Gibco BRL) containing 5% FCS. After centrifugation, cells were resuspended in complete medium (DMEM with 2 mML-glutamine, 100 IU/ml penicillin, 100 μg/ml streptomycin, and 10% heat-inactivated FCS; Gibco BRL) and plated in 24-well culture dishes (Costar, Cambridge, MA) at 37°C in a humidified atmosphere of 5% CO2, 95% air. After 48 hours, nonadherent cells were removed and adherent cells were cultured in complete medium. At confluence, cells were trypsinized and passaged in 150-cm2 culture flasks, with 2 medium changes each week. To ensure exclusion of macrophages from the culture, FLS between passages 3 and 8 were used for the RT-PCR analysis.

Stimulation of FLS cultures with recombinant human ATAC/Lptn.

For stimulation experiments, FLS were spread at a density of 2 × 104/ml in 24-well tissue culture plates and cultured in DMEM containing 10% FCS, 100 IU/ml of penicillin, and 100 μg/ml of streptomycin. At confluence, medium was removed, cells were washed once in PBS, and the medium was changed to DMEM supplemented with 1% insulin–transferrin–selenium (Gibco BRL), 100 IU/ml of penicillin, and 100 μg/ml of streptomycin. Cells were subsequently stimulated for 48 hours with recombinant human ATAC/Lptn (R&D Systems) at a concentration of 100 ng/ml. To control the specificity of ATAC/Lptn action, 5 μg/ml of anti-human ATAC/Lptn antibody (R&D Systems) was added to the control wells. After 24 and 48 hours, supernatants were collected, and the cells were trypsinized and resuspended in TRIzol reagent (Gibco BRL) for subsequent RNA isolation and quantitative RT-PCR.

MMP-2 mRNA was measured by real-time quantitative RT-PCR as described above. The MMP-2 primer sequences used for LightCycler analysis were as follows: for MMP-2 forward 1, 5′-CGCCGTCGCCCATCATCAAGTT-3′ and for MMP-2 reverse 1, 5′-GGAAGGCACGAGCAAAGGCATCAT-3′.

MMP-2 concentrations were measured in culture supernatants by ELISA (Amersham Pharmacia Biotech, Braunschweig, Germany) according to the manufacturer's instructions. In addition, MMP-2 and MMP-9 were analyzed by gelatin zymography of culture supernatants, as previously described (38). After loading cell supernatants and equal amounts of total protein on SDS gels containing 1.0 mg/ml gelatin, electrophoresis on nonreducing SDS–10% polyacrylamide gels (Bio-Rad, Hercules, CA) was performed. Gels were subsequently incubated in renaturation buffer (2.5% Triton X-100) for 30 minutes, followed by overnight incubation at 37°C in 5 mmoles/liter CaCl2, 50 mmoles/liter Tris HCl, pH 7.5, and subsequent staining for 1 hour at room temperature with Coomassie brilliant blue (0.5% Coomassie blue in 40% methanol/10% acetic acid).

Gels were destained in 40% methanol/10% acetic acid for 15 minutes, scanned with a Fluor-S MultiImager (Bio-Rad), and analyzed by densitometry using Multi-Analyst software (Bio-Rad). The molecular sizes of the bands displaying enzymatic activity were determined by comparison with standard proteins and with purified MMPs (Calbiochem-Novabiochem, San Diego, CA).

Statistical analysis.

Values are expressed as the mean ± SEM. Statistical differences between groups were evaluated by Student's t-test. P values were determined by a 2-sided calculation. P values less than 0.05 were considered significant.

RESULTS

  1. Top of page
  2. Abstract
  3. PATIENTS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. REFERENCES

ATAC/Lptn levels in sera and synovial fluids.

To determine ATAC/Lptn concentrations in sera and synovial fluids, a sandwich ELISA technique was established. Serum levels of ATAC/Lptn in RA patients ranged between 0.85 ng/ml and 200 ng/ml (mean ± SEM 15.62 ± 5.61 ng/ml) (Figure 1). Serum levels of ATAC/Lptn in RA patients were similar to those in healthy controls.

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Figure 1. Levels of activation-induced, T cell–derived, and chemokine-related cytokine (ATAC)/lymphotactin (Lptn) in serum and synovial fluid (SF) specimens from rheumatoid arthritis (RA) patients, osteoarthritis (OA) control patients, and healthy control subjects. ATAC/Lptn concentrations were measured by enzyme-linked immunosorbent assay (ELISA) in A, serum samples from 40 RA patients and 8 healthy controls and B, synovial fluid samples from 15 RA patients and 15 OA patient controls. No significant difference in ATAC/Lptn levels between RA patients and controls was detected.

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In synovial fluid samples from RA patients, ATAC/Lptn concentrations varied between 0.19 and 100 ng/ml (mean ± SEM 18.05 ± 8.26 ng/ml). There was no significant difference in synovial fluid ATAC/Lptn levels between RA patients and OA control patients. In addition, there was no correlation between the Disease Activity Score and serum or synovial fluid concentrations of ATAC/Lptn in RA patients (data not shown).

Phenotype of PBMCs expressing ATAC/Lptn.

PBMCs were isolated from RA patients and healthy controls and analyzed for the expression of ATAC/Lptn by intracellular cytokine staining and flow cytometry. No constitutive production of ATAC/Lptn could be detected in unstimulated PBMCs isolated from RA patients or healthy controls. However, ATAC/Lptn could be induced by activation of PBMCs with PMA and ionomycin. After 5 hours of PMA/ionomycin stimulation, a mean ± SEM of 23.76 ± 4.11% CD8+ T cells and 2.56 ± 0.55 CD4+ T cells from RA patients stained positive for ATAC/Lptn (Figure 2A). These ATAC/Lptn levels were not significantly different from those induced in PBMCs from healthy controls.

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Figure 2. Flow cytometric analysis of ATAC/Lptn expression in peripheral blood mononuclear cells (PBMCs) after intracellular staining. PBMCs were isolated from 10 RA patients and 3 healthy controls. A, Analysis of phorbol myristate acetate/ionomycin–stimulated PBMCs revealed that ATAC/Lptn was predominantly expressed in CD8+ T cells in both RA patients and healthy controls. B, In RA patients, however, a significantly greater proportion of the CD4+,CD28− T cells stained positive for ATAC/Lptn. ∗∗∗ = P < 0.001. C, Most of the ATAC/Lptn-producing CD8+ T cells were deficient for the costimulatory molecule CD28 (CD8+,CD28−) in both groups. Values are the mean and SEM. See Figure 1 for other definitions.

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To further characterize the subtype of ATAC/Lptn-expressing CD4+ and CD8+ T cells, PBMCs were stained for the costimulatory CD28 molecule. The results revealed a significantly greater proportion of ATAC/Lptn-positive CD4+,CD28− T cells in RA patients compared with healthy controls (P < 0.0001) (Figure 2B). In addition, the majority of the ATAC/Lptn-expressing CD8+ T cells lacked the CD28 surface molecule (CD8+,CD28−) in both RA patients and healthy controls (Figure 2C).

Expression of ATAC/Lptn mRNA in RA synovium.

ATAC/Lptn mRNA expression was analyzed in synovial tissues from 20 RA patients by RT-PCR with oligonucleotide primers specific for human ATAC/Lptn. Synovial biopsies obtained from 15 patients with OA were used as controls. As demonstrated in Figure 3A, ATAC/Lptn mRNA was detected in 17 of the 20 RA tissue samples (85%), but in only 3 of the 15 OA specimens (20%). We used β-actin as an internal control for gene expression.

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Figure 3. Activation-induced, T cell–derived, and chemokine-related cytokine (ATAC)/lymphotactin (Lptn) mRNA expression in synovial tissues from 20 rheumatoid arthritis (RA) and 15 osteoarthritis (OA) patients. A, Total cellular RNA was isolated from synovial tissue specimens. After standardization of cDNA samples against β-actin, polymerase chain reaction (PCR) for ATAC/Lptn revealed the expression of ATAC/Lptn mRNA in 17 RA samples, but only 3 OA samples. B, ATAC/Lptn mRNA expression was investigated by real-time quantitative reverse transcription–PCR. Values were determined in duplicate. ATAC/Lptn mRNA levels were standardized against β-actin to compare ATAC/Lptn mRNA between samples. ATAC/Lptn gene expression related to β-actin expression was significantly elevated in RA patients compared with OA patients. Bars show the mean.

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Quantification of ATAC/Lptn mRNA levels was performed by using a real-time quantitative RT-PCR in combination with LightCycler technology. ATAC/Lptn mRNA levels calculated by reference to a standard curve were determined in relation to β-actin mRNA levels for each sample. Levels of ATAC/Lptn mRNA were up to 3.5-fold higher in RA samples than in OA control samples (Figure 3B). The mean levels of ATAC/Lptn transcripts were also significantly increased in RA synovium compared with OA synovium (P < 0.001).

Localization of ATAC/Lptn expression in RA synovium.

To localize ATAC/Lptn mRNA expression in the synovial tissue, paraffin sections were subjected to in situ hybridization with DIG-labeled RNA probes. Hybridization sites were detected by enzyme detection methods using alkaline phosphatase–conjugated anti-DIG antibodies. Consistent with the results of the quantitative RT-PCR analysis, ATAC/Lptn mRNA was strongly expressed in sections of synovial tissue from RA patients. ATAC/Lptn mRNA–positive cells were predominantly localized in lymphocytic infiltrates in the sublining layer (Figure 4A). There was an apparent relationship between the expression of ATAC/Lptn and the density of the mononuclear cell infiltrate. The expression of ATAC/Lptn mRNA in inflammatory cells of the lining layer was weaker and largely restricted to perivascular areas. In synovial tissue sections from OA control patients, ATAC/Lptn mRNA–expressing cells were rarely seen in the lining and sublining areas (<2%). Sense probes served as negative controls and were always included (Figure 4B).

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Figure 4. Localization of ATAC/Lptn mRNA expression in RA synovium by in situ hybridization. Paraffin sections of synovial tissue samples from RA patients were stained by in situ hybridization with a digoxigenin-labeled riboprobe and enzyme detection. ATAC/Lptn mRNA–expressing cells were predominantly localized in the sublining layer of the RA synovium (A). No staining was detected with in situ hybridization with a sense probe (B). By immunofluorescence, ATAC/Lptn mRNA–producing cells detected by in situ hybridization in the same section were shown to be negative for the Ki-M1P antigen (C), but mostly coexpress the CD3 surface antigen (D). In addition, immunohistochemistry for CD4 and CD8 antigens in serial sections revealed that both CD4+ (E) and CD8+ (F) T cells represent a cellular source of ATAC/Lptn in RA synovium. (Original magnification × 200.) See Figure 1 for definitions.

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To further characterize the immunophenotype of ATAC/Lptn mRNA–expressing cells, tissue sections were sequentially stained for ATAC/Lptn mRNA by in situ hybridization and for cell surface antigens by immunofluorescence (CD3 and Ki-M1P) or immunohistochemistry (CD4 and CD8), respectively. Double-staining experiments revealed that the majority of ATAC/Lptn mRNA–positive cells were T cells (CD3+) (Figure 4D). In contrast, no ATAC/Lptn mRNA expression was detected in cells of the monocyte/macrophage lineage (Ki-M1P) (Figure 4C). Immunostaining for CD4 and CD8 antigens further demonstrated that ATAC/Lptn mRNA was present in both CD4+ (Figure 4E) and CD8+ (Figure 4F) T cells of the synovial infiltrate.

To analyze ATAC/Lptn expression on the protein level, paraffin sections of synovial tissue specimens were stained by immunohistochemistry using a polyclonal rabbit anti-human ATAC/Lptn antibody. Consistent with the results of the in situ hybridization studies, ATAC/Lptn was found to be strongly expressed in lymphocytic infiltrates of the sublining layer (Figure 5A). No positive staining was observed for the isotype control (Figure 5B). Double staining for cell surface antigens confirmed that most of the cells containing ATAC/Lptn protein were T cells (CD3+) (Figure 5C). Consistent with the results of the flow cytometric analysis of PBMCs, CD8+ T cells were the major source of ATAC/Lptn (Figure 5D). Other cellular sources of ATAC/Lptn in the rheumatoid synovium were identified, including some CD4+ T cells (Figure 5E), NK cells (NK1+) (Figure 5F), dendritic cells (Fascin positive) (Figure 5G), and mast cells (tryptase positive) (Figure 5H). In contrast, no positive staining was detected in synovial macrophages (Ki-M1P+) (data not shown).

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Figure 5. Localization of ATAC/Lptn expression in RA synovium by double-label immunohistochemistry. Paraffin sections of synovial tissues were stained for ATAC/Lptn with a polyclonal rabbit anti-human ATAC/Lptn antibody and detected by the biotin–tyramide signal amplification system (fluorescein isothiocyanate). Double staining for cell surface antigens (rhodamine red-X) was performed by indirect immunofluorescence. Sections were analyzed on a Leica TCS NT laser scanning microscope. A, ATAC/Lptn was predominantly detected in the sublining layer of the rheumatoid synovium. B, No staining was observed in the isotype control. C, Double staining for cell surface antigens revealed that ATAC/Lptn-positive cells mostly coexpressed the CD3 antigen. D, CD8+ T cells and E, CD4+ T cells were both identified as a source of ATAC/Lptn. In addition, some F, NK cells, G, dendritic cells, and H, mast cells stained positive for ATAC/Lptn. (Original magnification × 400.) See Figure 1 for definitions.

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ATAC/Lptn expression was also analyzed in different stages of RA as well as in other inflammatory arthritides. In OA (Figure 6A) and reactive arthritis (Figure 6B), only weak staining signals were detected in mononuclear cells scattered throughout the synovial tissue. In psoriatic arthritis (Figure 6C), ATAC/Lptn-positive cells were found predominantly in perivascular lymphocytic infiltrates. Analysis of early RA (Figure 6D) compared with late RA (Figure 6E) revealed a more prominent ATAC/Lptn staining of inflammatory cells that were predominantly localized in the sublining layer in early RA.

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Figure 6. Expression of ATAC/Lptn in early and late RA and in other inflammatory arthritides. In A, OA and B, reactive arthritis, only weak staining signals were detected. In C, psoriatic arthritis, ATAC/Lptn-positive cells were found predominantly in perivascular lymphocytic infiltrates. In D, early RA, as compared with E, late RA, a more prominent staining of inflammatory cells was seen mainly in the sublining area. (Original magnification × 200.) See Figure 1 for definitions.

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Expression of the ATAC/Lptn receptor (XCR1) in different cell lines, cultured FLS, and RA synovium.

To characterize the biologic activity of ATAC/Lptn in RA synovium by definition of target cells for ATAC/Lptn action, we analyzed the expression of the ATAC/Lptn receptor (XCR1) by RT-PCR in 4 different cell lines (TK 173, TK 188, Jurkat, and U937), cultured FLS, and in RA synovial biopsy tissues obtained from 3 different donors. After cDNA synthesis and standardization against β-actin, PCR was performed with oligonucleotide primers specific for the human ATAC/Lptn receptor XCR1. XCR1 expression was present in the T cell line (Jurkat), the renal fibroblast cell lines (TK 173 and TK 188), and in FLS generated from 3 different donors (FLS 1–3) as well as in the corresponding synovial tissues (ST 1–3) (Figure 7A). The level of XCR1 expression in FLS was dependent on the number of passages in culture. The receptor was more strongly expressed in early-passage cultures (FLS 1, passage 4) than in late-passage cultures (FLS 2 and 3, passage 8). No signal could be detected in the monocyte cell line (U937). As negative control, water (instead of cDNA) was always included. To exclude genomic DNA contamination in XCR1 RT-PCR analysis, RT-negative controls were analyzed with each sample.

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Figure 7. Expression of the ATAC/Lptn receptor XCR1 in different cell lines, cultured fibroblast-like synoviocytes (FLS), and RA synovium. Total cellular RNA was extracted from 4 cell lines (renal fibroblast lines TK 173 and TK 188, acute lymphoblastic leukemia T cell line Jurkat, and monocyte cell line U937) as well as from synovial tissues (ST 1–3) and cultured FLS (FLS 1–3) from 3 different RA patients. After cDNA synthesis and standardization against β-actin, PCR was performed with oligonucleotide primers specific for XCR1. The T cell line, renal fibroblast cell lines, and RA FLS and synovial tissue specimens were shown by reverse transcription–PCR to express XCR1 (A). To exclude genomic DNA contamination, reverse transcriptase (RT)–negative controls were analyzed with each sample. Results are representative of at least 3 independent experiments. In situ hybridization for XCR1 in cultured FLS (early-passage FLS 1) revealed clear hybridization signals in the cytoplasm (B). Hybridization with a sense probe revealed no staining (C). The ATAC/Lptn receptor was also detected by in situ hybridization of RA synovium (D), especially in synovial fibroblasts (FLS) (E). In addition, lymphocytes (Lc) (F) and some synovial macrophages (MP) (E) were identified by morphology to be positive for XCR1 mRNA. No specific staining was detected in hybridization experiments with the sense probe (G–I). See Figure 3 for other definitions.

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To verify RT-PCR results, in situ hybridization for XCR1 was performed on cultured FLS generated from RA synovial tissue. As demonstrated in Figure 7B, fibroblasts grown on chamber slides showed a clear hybridization signal after in situ hybridization for the ATAC/Lptn receptor. No staining was observed for hybridization with the sense probe (Figure 7C). The XCR1 receptor could also be detected in FLS from OA control patients. By quantitative RT-PCR analysis, XCR1 mRNA levels in OA-derived FLS were significantly lower than those in RA-derived FLS (data not shown).

Similarly, in situ hybridization of paraffin sections of RA synovial tissue samples (Figure 7D) revealed XCR1 transcripts in FLS (Figure 7E). In addition, hybridization signals were detected in lymphocytes (Figure 7F) and some synovial macrophages (Figure 7E) by microscopic analysis of hybridized sections. Negative controls included hybridization experiments with a sense probe and hybridization after nuclease digestion of the target RNA (Figures 7G–I).

Stimulation of cultured FLS with ATAC/Lptn.

Due to the novel finding of ATAC/Lptn receptor mRNA expression in FLS, we were interested in analyzing the effects of ATAC/Lptn on FLS function. Cultured FLS stimulated with human recombinant ATAC/Lptn and RNA samples isolated from stimulated and unstimulated cultures were screened by a cDNA microarray technique (data not shown). The results suggested that ATAC/Lptn may down-regulate MMP-2 production in cultured FLS. To verify this finding, RNA was isolated from 10 different FLS cultures and analyzed by quantitative real-time RT-PCR. As shown in Figure 8A, stimulation of FLS with 100 ng/ml of ATAC/Lptn decreased MMP-2 mRNA levels up to 10-fold after 24 hours of incubation. This effect could be blocked by addition of anti-ATAC/Lptn antibody at a dose (5 μg/ml) that, in preliminary experiments, completely abrogated ATAC/Lptn stimulation.

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Figure 8. Down-regulation of matrix metalloproteinase 2 (MMP-2) production in ATAC/Lptn-stimulated cultured fibroblast-like synoviocytes (FLS) from RA patients. FLS were stimulated with human recombinant ATAC/Lptn (100 ng/ml) for 48 hours. A, MMP-2 mRNA transcription was significantly reduced after 24 hours of incubation, as measured by quantitative reverse transcription–polymerase chain reaction (RT-PCR) (P = 0.01). This effect could be reversed by the addition of an ATAC/Lptn-specific antibody (5 μg/ml). B, A significant reduction in MMP-2 production was detected by ELISA of 48-hour culture supernatants. C, Gelatin zymography for MMP-2 activity confirmed the results shown in B. MMP-2 and MMP-9 were visualized by negative staining with Coomassie brilliant blue, and proteolytic bands were scanned densitometrically. No effect on MMP-9 production was detected. D, MMP-2 production in FLS stimulated with ATAC/Lptn was shown to be down-regulated in a dose-dependent manner. Values are the mean and SEM. Results were consistent in at least 3 independent experiments. ∗∗ = P < 0.01. See Figure 1 for other definitions.

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In parallel, supernatants of FLS cultures were analyzed for secreted MMP-2 by ELISA. After 48 hours of incubation, production of MMP-2 was significantly reduced in cultures stimulated with 100 ng/ml ATAC/Lptn (P = 0.01). This effect could also be reversed by addition of the ATAC/Lptn-specific antibody (Figure 8B). The difference between the levels of ATAC/Lptn-induced changes in MMP-2 mRNA and protein concentrations could be explained by an accumulation of MMP-2 protein in culture supernatants, since MMP-2 protein levels were measured by ELISA in culture supernatants (without medium change) after 48 hours of incubation.

To further analyze the effect of ATAC/Lptn on MMP-2 activity in FLS culture supernatants, zymography for MMP-2 and MMP-9 was performed. Consistent with the mRNA and protein data, ATAC/Lptn (at 48 hours after stimulation) markedly reduced the activity of MMP-2 (Figure 8C) but had no effect on the activity of MMP-9 (data not shown). The inhibition of MMP-2 production by ATAC/Lptn was shown to be dose-dependent by both quantitative RT-PCR (Figure 8D) and ELISA. The results shown in Figures 8A–D were consistent among at least 3 independent experiments.

DISCUSSION

  1. Top of page
  2. Abstract
  3. PATIENTS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. REFERENCES

To our knowledge, this is the first study to demonstrate a prominent expression of ATAC/Lptn and its receptor XCR1 in the synovial tissue of RA patients. We established a highly sensitive ELISA to examine the levels of ATAC/Lptn in the peripheral blood and in synovial fluid from affected joints. ATAC/Lptn levels in serum and synovial fluid from RA patients were not significantly elevated levels compared with those in healthy controls and in OA control patients, respectively. This finding is in contrast to the observed overexpression of ATAC/Lptn mRNA and protein in RA compared with OA synovium. A possible explanation for this discrepancy is a strictly local induction of ATAC/Lptn in RA synovium. ATAC/Lptn may be secreted locally and may exert its effects directly at the site of chronic inflammation in RA. A tight regulation of ATAC/Lptn production, with induction only in response to proinflammatory stimuli, is also suggested by our finding that intracellular ATAC/Lptn was detectable in PBMC cultures only after stimulation with PMA and ionomycin.

Consistent with previous results (12), ATAC/Lptn expression was predominantly detected in CD8+ T cells and in a small proportion of CD4+ T cells. Analysis of the immunophenotype of ATAC/Lptn-positive T cells revealed that the majority of CD4+ and the CD8+ T cells containing ATAC/Lptn lacked expression of the costimulatory molecule CD28. Interestingly, expansion of CD4+,CD28− and CD8+,CD28− T cell clones in RA has previously been identified (39–41).

CD4+,CD28− T cells are infrequent in normal subjects, but account for up to 50% of CD4+ T cells in RA patients (42). In line with a role of these cells in the pathogenesis of RA, a correlation between the expansion of CD4+,CD28− T cells and the manifestation of clinical symptoms has previously been observed: Patients with extraarticular disease manifestations were found to carry the highest frequency of this unusual T cell subset. CD4+,CD28− T cells were shown to infiltrate the synovial lesions, to produce high amounts of IFNγ (43), and to lyse target cells by producing perforin and granzyme B (44). These properties of CD4+,CD28− T cells further support a direct contribution of this T cell subset to the chronic inflammatory process in RA.

Quantitative RT-PCR for the detection of ATAC/Lptn mRNA expression revealed that ATAC/Lptn transcript levels were significantly elevated in RA synovium compared with OA synovium (P < 0.001). In situ hybridization and immunohistochemistry studies further demonstrated that on both the transcript (mRNA) level and the protein level, ATAC/Lptn-positive cells were mainly localized in lymphocytic infiltrates of the sublining layer. Double immunostaining for cell surface antigens revealed that most of these ATAC/Lptn-positive cells were CD3+ T cells, whereas cells of the monocyte/macrophage lineage did not contain ATAC/Lptn.

Further characterization of the phenotype of ATAC/Lptn-positive T lymphocytes revealed the expression of ATAC/Lptn in both the CD8+ and the CD4+ T cell subsets. These findings were consistent with those of previous studies demonstrating that activated CD8+ T cells and a small proportion of CD4+ T cells represent the most important source of this chemokine in the peripheral blood (19). In RA synovium, ATAC/Lptn expression could also be demonstrated in mast cells, NK cells, and dendritic cells. Expression of ATAC/Lptn in these 3 different cell types has previously been described. Rumsaeng et al (14) demonstrated that ATAC/Lptn synthesis and release could be induced in mast cells in response to Fcε receptor I aggregation. In addition, ATAC/Lptn expression has been demonstrated in NK cells (15) and was shown to be up-regulated in these cells by activating Ly-49 NK receptors (45). In Crohn's disease, ATAC/Lptn expression has been demonstrated in dendritic cells (27). Since there is evidence that expression of ATAC/Lptn is up-regulated in dendritic cells during maturation (46), our finding of ATAC/Lptn-positive dendritic cells in RA synovium may indicate the presence of mature dendritic cells in the inflamed synovium. Our present data and our previous data (12) indicate the absence of ATAC/Lptn in cells of the monocyte/macrophage lineage, including cytokine-activated monocytes (U937), fibroblasts, and HeLa cells.

Analysis of ATAC/Lptn expression in other inflammatory arthritides, including OA, reactive arthritis, and psoriatic arthritis, revealed a prominent staining of inflammatory cells only in psoriatic arthritis, which suggests a role for ATAC/Lptn in Th1 cytokine–associated disease states. Comparison of early-stage and late-stage RA demonstrated a more pronounced expression of ATAC/Lptn in lymphocytic infiltrates of the sublining layer in early RA, which also indicates that ATAC/Lptn might exert an important function in the early phase of RA pathogenesis.

To further elucidate a potential functional role in RA, we studied the expression of the ATAC/Lptn receptor XCR1 in different cell types known to be involved in the pathogenic process of RA. As detected by RT-PCR, T cells and, surprisingly, FLS isolated from the synovial tissue of RA patients were shown to express the ATAC/Lptn receptor mRNA. XCR1 has been identified as a specific functional, G protein–coupled receptor for ATAC/Lptn and has been detected in selected tissues, including placenta, spleen, and thymus, but was rarely found in PBMCs (16, 17). In addition, B cells and neutrophils have also been shown to express the ATAC/Lptn receptor (47) and to respond to ATAC/Lptn by chemotaxis. XCR1 expression was recently detected in tissue macrophages in a murine model of listeriosis (22). The RT-PCR results in our study indicate that XCR1 is also highly expressed in the rheumatoid synovium.

Since T cells represent the main source of ATAC/Lptn expression in RA synovium, expression of XCR1 on these cells may indicate autocrine and/or paracrine regulation by ATAC/Lptn. The finding that FLS were also positive for XCR1 expression is rather striking and suggests that these cells also represent targets of ATAC/Lptn activity in RA synovium. The presence of functional ATAC/Lptn receptors on FLS is supported by our finding that ATAC/Lptn dose-dependently regulated the production of MMP-2 in cultured FLS at the mRNA and protein levels.

Previous studies have largely focused on the chemoattractant properties of ATAC/Lptn. It has been shown to induce chemotactic responses in T cells (10, 18, 48), NK cells (15), and B cells and neutrophils (47), but this functional activity has not been confirmed by other investigators (16, 19). More recent data indicate that Lptn may particularly enhance the migration of antigen-activated CD62Llow T cells (49) and memory T cells (50). In addition, ATAC/Lptn gene–modified dendritic cells were capable of attracting T cells and NK cells in chemotaxis assays (21) and were successfully used to enhance antitumor immunity. In light of these in vitro functions of ATAC/Lptn, the expression of ATAC/Lptn in RA synovium may contribute to the selective recruitment of T cells to the synovium of affected joints, where accumulated T cells are known to be harbored.

Recent studies further suggest that apart from its chemotactic properties, ATAC/Lptn might exert multiple biologic functions. In a murine model of listeriosis, ATAC/Lptn as well as MIP-1α, MIP-1β, and RANTES were shown to be cosecreted with IFNγ and to act as type 1 cytokines by up-regulating CD40, IL-12, and TNFα in macrophages (22). As part of a proinflammatory cytokine milieu in RA, ATAC/Lptn may enhance the expression of these mediators in synovial macrophages as well, and thus may perpetuate the inflammatory destruction of the articular tissue. However, these hypothesized effects of ATAC/Lptn stimulation on synovial macrophage function have not yet been analyzed in detail.

Our data on the regulation of MMP-2 in FLS suggest a more complex role of ATAC/Lptn in RA that may include additional immunomodulatory effects or even antiinflammatory effects. Down-regulation of MMP-2 production was demonstrated on both the RNA and protein levels in cultured FLS after stimulation with ATAC/Lptn. This effect of ATAC/Lptn could be abrogated by the addition of neutralizing anti-ATAC/Lptn antibodies. Thus, apart from T cell recruitment, ATAC/Lptn might have an additional function in regulating the mechanisms of disease progression in RA. However, at this point, it cannot be excluded that the in vitro effects observed in cultured FLS may not correlate with the effects in vivo, especially in the environment of a proinflammatory cytokine milieu characteristic of RA. Further studies, including studies in animal models, are therefore needed to more precisely characterize the role of ATAC/Lptn in RA.

In summary, our study demonstrates an increased expression of ATAC/Lptn in CD4+,CD28− T cells in the peripheral blood as well as in CD4+ and CD8+ T cells in the synovial infiltrate in patients with RA. Our study also shows the expression of the ATAC/Lptn receptor XCR1 in lymphocytes, synovial macrophages, and, interestingly, FLS. Functional analysis of cultured FLS stimulated with ATAC/Lptn in vitro suggests that this chemokine may exert an immunomodulatory effect in RA in addition to its known chemotactic activity for T cells. Our findings suggest a crucial role for this chemokine in the pathogenesis of the chronic inflammatory process in RA.

REFERENCES

  1. Top of page
  2. Abstract
  3. PATIENTS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. REFERENCES