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Abstract

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

Objective

To investigate the messenger RNA (mRNA) and protein expression of the recently discovered platelet-derived growth factor C (PDGF-C) and PDGF-D in the synovial membrane (SM) of patients with rheumatoid arthritis (RA) and osteoarthritis (OA) and to assess the localization and cellular source of these proteins in the SM and their functional influence on synovial fibroblasts.

Methods

Expression of mRNA for PDGFs A, B, C, and D as well as for PDGF receptor (PDGFR) α and β chains in RA and OA SM samples was assessed by real-time reverse transcription–polymerase chain reaction. Protein levels of PDGF-C and PDGF-D were quantified by immunoblotting. Regional and cellular localization of PDGF-C and PDGF-D in the SM was investigated by double-staining immunohistochemistry. In addition, the influence of PDGF-D on the proliferation of synovial fibroblasts and their matrix metalloproteinase (MMP-1) mRNA expression were determined.

Results

The expression of mRNA for PDGFs A, B, and C and for PDGFR α and β chains was comparable in RA and OA SM samples; in contrast, the expression of mRNA for PDGF-D was significantly higher in OA SM. PDGF-C protein was not differentially expressed in OA and RA. The expression of PDGF-D protein was significantly higher in RA SM (full-length and activated form). PDGF-C and PDGF-D were expressed throughout the SM (lining layer, diffuse infiltrates, and stroma) by both synovial fibroblasts and macrophages. In addition, PDGF-D increased the proliferation of synovial fibroblasts and the expression of mRNA for MMP-1.

Conclusion

PDGF-C and PDGF-D are expressed by synovial fibroblasts and macrophages in RA and OA SMs. The levels of PDGF-D protein were significantly higher in RA SM. In addition, PDGF-D stimulated synovial fibroblast proliferation and expression of MMP-1. These findings may have pathogenetic implications for cellular transformation and matrix remodeling in the RA SM.

Platelet-derived growth factor (PDGF) is a major mitogen for fibroblasts and other cells of mesenchymal origin. Its biologic function is mediated by PDGF receptor tyrosine kinases, which are formed by homo- or heterodimers of α and β chains. The classic PDGF family consists of 2 members, PDGF-A and PDGF-B, which form homo- or heterodimers with distinct receptor binding abilities (1). Recently, 2 new family members have been discovered through database mining tools (2–4). Besides the core PDGF/vascular endothelial growth factor (VEGF) domain, PDGF-C and PDGF-D contain an additional CUB (complement subcomponents C1r/C1s, urchin epidermal growth factor–like protein, and bone morphogenetic protein 1) N-terminal domain, which is cleaved upon activation (2–4).

PDGF-C messenger RNA (mRNA) is expressed in many tissues, particularly the heart, liver, pancreas, and kidney. Although its detailed functional role remains to be investigated, PDGF-C seems to be a potent transforming factor, because it is expressed in many tumor cell lines (5), induces tumor formation, and efficiently transforms a murine fibroblast cell line (6). The ability to stimulate the growth of smooth muscle cells (5) and of microvessels in the murine cornea (7) also suggests an important role in angiogenesis. In addition, PDGF-C may contribute to fibrotic processes, since the hearts of PDGF-C–transgenic mice exhibit a progressive hypertrophy with strong proliferation of cardiac fibroblasts (2).

Transfection with PDGF-D results in transformation of murine NIH3T3 cells, which are then characterized by anchorage-independent growth, actin reorganization, an increased proliferation rate, and the capacity to induce tumor formation in mice (8, 9). In addition, PDGF-D is known to be involved in angiogenic processes (10).

Rheumatoid arthritis (RA) is characterized by progressive joint inflammation accompanied by synovial hyperplasia with some similarities to malignancy. Synovial cells located in pannus show an invasive behavior, are of variable cell size (11), and express high levels of protooncogenes (12) and tyrosine phosphorylated proteins (13). PDGF receptor (PDGFR) α and β chains are detected in fibroblast-like cells in the inflamed synovial membrane (SM) (14). PDGFRs, as well as PDGF-A and PDGF-B, are expressed to a significantly higher degree in the SM of RA patients compared with that of osteoarthritis (OA) patients (15). Furthermore, fibroblast-like cells from RA synovium respond to PDGF-BB stimulation with proliferation (16) and anchorage-independent growth in vitro (17), showing the importance of PDGFs in RA.

We therefore investigated the expression of the newly discovered PDGF-C and PDGF-D at the mRNA and protein levels in SM samples from patients with RA and OA, the cellular distribution of these factors within the SM, and the functional effects of PDGF-D on synovial fibroblasts.

PATIENTS AND METHODS

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

Patients.

Synovial tissue was obtained from patients with RA or OA who were undergoing synovectomy at the Orthopedic Clinic, Waldkrankenhaus Rudolf Elle (Eisenberg, Germany). All RA patients (n = 12) fulfilled the American College of Rheumatology (ACR; formerly, the American Rheumatism Association) 1987 criteria for RA (18), and all OA patients (n = 14) fulfilled the ACR criteria for OA of the knee (19). The study was approved by the Ethics Committee of the Friedrich Schiller University.

Synovial tissue samples were snap-frozen with or without embedding in tissue-freezing medium (Leica, Nussloch, Germany) and then stored at −70°C or were placed in RNAlater (Ambion, Austin, TX) overnight, then frozen and stored at −70°C.

Analysis of mRNA expression by real-time reverse transcription–polymerase chain reaction (RT-PCR).

Total RNA was isolated from RNAlater-treated samples after initial homogenization with an UltraTurrax homogenizer (Janke & Kunkel, Staufen, Germany) using an RNeasy kit (Qiagen, Hilden, Germany) according to the supplier's instructions. Complementary DNA (cDNA) was prepared using oligo(dT) primers and SuperScript reverse transcriptase (Invitrogen, Karlsruhe, Germany).

For the quantitation of PDGFs A, B, C, and D, PDGFR α and β chains, and the housekeeping gene GAPDH, the open-reading frame gene sequences (or parts of these) were cloned using the TOPO TA cloning kit (Invitrogen) and used as external standards. Real-time RT-PCR was performed on a LightCycler instrument (Roche Diagnostics, Mannheim, Germany) as previously described (20), using the primer pairs shown in Table 1.

Table 1. Primer and product sizes for real-time reverse transcription–polymerase chain reaction*
GeneForward primer (5′[RIGHTWARDS ARROW]3′)Reverse primer (5′[RIGHTWARDS ARROW]3′)Size, bpAnnealing temperature; amplification timeAdditional heating step
  • *

    An additional heating step to melt primer dimers was performed for 5 seconds at the temperatures indicated. PDGF = platelet-derived growth factor; MMP-1 = matrix metalloproteinase 1.

PDGF-AACACGAGCAGTGTCAAGTGCCCTGCAGTATTCCACCTTGG7760°C; 10 seconds87°C
PDGF-BAGATCGAGATTGTGCGGAAGCAGCTGCCACTGTCTCACAC9453°C; 10 seconds84°C
PDGF-CGCCAGGTTGTCTCCTGGTTATGCTTGGGACACATTGACAT8560°C; 10 seconds81°C
PDGF-DCCCAGGAATTACTCGGTCAAACAGCCACAATTTCCTCCAC13151°C; 10 seconds81°C
PDGFRαTGGGAGTTTCCAAGAGATGGTGTTCCTTCAACCACCTTCC8553°C; 10 seconds83°C
PDGFRβGTGCTCACCATCATCTCCCTACTCAATCACCTTCCATCGG7751°C; 10 seconds80°C
GAPDHCGGAGTCAACGGATTTGGAGCCTTCTCCATGGTGGTG30753°C; 20 seconds77°C
MMP-1CGACTCTAGAAACACAAGAGCAAGAAAGGTTAGCTTACTGTCACACGCTT32258°C; 25 seconds

The general amplification protocol (50 cycles) was set as follows: initial denaturation for 3 minutes at 95°C, then denaturation for 5 seconds at 95°C, specific primer annealing temperature for 10 seconds, and amplification at 72°C for the indicated time period (Table 1). The general settings for the melting curve protocol (1 cycle) were as follows: denaturation at 95°C, cooling to 5°C above the primer annealing temperature, heating to 95°C (temperature change 0.1°C/second), and final cooling at 40°C for 5 minutes. The fluorescence emitted by double-stranded DNA–bound SYBR Green was measured once at the end of each additional heating step and continuously during the melting curve program. The concentrations of cDNA present in each sample were calculated by the LightCycler software, using the external standard curves. In order to normalize the amount of cDNA in each sample and to guarantee the comparability of the calculated mRNA expression in all samples analyzed, the housekeeping gene GAPDH was also analyzed. Product specificity was confirmed by melting curve analysis and initial cycle sequencing of the PCR products. The amplification efficiency (defined as E = 10–1/slope, where the slope is derived from a plot depicting the cycle number versus the log concentration) was generally between 1.7 and 2.1, with a maximum difference of 8.5% between cloned standards and cDNA samples. These values fully comply with the published recommendations (21).

Western blotting analysis.

Frozen tissue was homogenized in complete lysis buffer (Nuclear Extraction kit; Active Motif, Rixensart, Belgium) containing proteinase inhibitors (Complete; Roche Diagnostics). Lysates were prepared with the Nuclear Extraction kit according to the supplier's instructions (whole cell lysate protocol). Protein content was determined using the bicinchoninic acid assay (Pierce, Rockford, IL) following acetone precipitation of a 25-μl sample aliquot. PDGF-C and PDGF-D were detected by immunoblotting of 10 μg of lysate, resolved by a reducing sodium dodecyl sulfate–polyacrylamide gel electrophoresis, using goat anti-human PDGF-C (sc-18228; Santa Cruz Biotechnology, Santa Cruz, CA) and goat anti-human PDGF-D (AF1159; R&D Systems, Wiesbaden, Germany) as primary antibodies and horseradish peroxidase (HRP)–conjugated donkey anti-goat IgG (sc-2020; Santa Cruz Biotechnology) as secondary antibody. The blots were stripped and reprobed with mouse anti-human β-actin (clone AC-15; Sigma, Deisenhofen, Germany) and HRP-conjugated goat anti-mouse IgG (Sigma) to ensure equal loading.

Immunohistochemistry analysis.

Analyses were performed on serial cryostat sections (5 μm) of RA or OA SM samples (fixed with acetone for 10 minutes at 4°C and air-dried), using the above antibodies for PDGF-C and PDGF-D, the fibroblast-specific antibody AS02 (CD90; Dianova, Hamburg, Germany), and the marker antibody for macrophages CLB-CD14B (CD14; Janssen Biochimica, Beerse, Belgium). Monoclonal antibodies against PDGF were diluted in phosphate buffered saline (PBS)/5% normal human serum, added to sections that had been preincubated with 5% normal human serum in PBS for 20 minutes, and incubated for 30 minutes at room temperature in a humidified chamber. HRP-conjugated donkey anti-goat IgG (in PBS/5% normal human serum) was added for 30 minutes. Peroxidase staining was revealed with diaminobenzidine (0.5 mg of diaminobenzidine in 1 ml of PBS containing 30 μl H2O2) treatment for 5 minutes. The slides were then washed, blocked with 20% normal human serum, and incubated with monoclonal antibody against CD90 or CD14 in PBS/5% horse serum for 30 minutes. As secondary antibody, alkaline phosphatase–conjugated goat anti-mouse IgG was used in conjunction with Naphthol AS-MX phosphate, Fast Blue BB salt, and levamisole. The slides were washed and covered with Aquatex (Merck, Darmstadt, Germany). Isotype-matched mouse monoclonal antibody or normal goat serum served as controls for all immunohistochemical analyses, which always yielded negative results.

Functional assays.

Synovial fibroblasts from 3 RA patients were purified as previously described (22). For proliferation assays, the cells (first passage) were seeded at 1.7 × 103 cells/well in a 96-well flat-bottomed plate and cultured in DMEM containing 100 μg/ml of gentamicin, 100 μg/ml of penicillin/streptomycin, 20 mM HEPES, and 10% fetal calf serum (FCS) for 24 hours at 37°C in an atmosphere containing 5% CO2. The cells were then serum-starved in the same medium containing 1% FCS for 72 hours and then stimulated for 72 hours by the addition of 10, 20, 50, or 100 ng/ml of PDGF-D core protein (kindly provided by U. Eriksson, Stockholm, Sweden). Bromodeoxyuridine (BrdU) was added for the last 24 hours of culture, after which the incorporation of BrdU was measured using a cell proliferation enzyme-linked immunosorbent assay (Cell Proliferation ELISA, BrdU; Roche Diagnostics) according to the manufacturer's instructions.

For analysis of matrix metalloproteinase 1 (MMP-1) mRNA expression, synovial fibroblasts were cultured and serum-starved as described above at a density of 2 × 105 cells/well (6-well plate). Subsequently, the cells were stimulated with 100 ng/ml of PDGF-D for 24 hours in medium with 1% FCS and then lysed. Total RNA was isolated, cDNA was synthesized, and real-time RT-PCR was performed using the primers and conditions mentioned above or as shown in Table 1.

Statistical analysis.

The nonparametric Mann-Whitney U test was applied for the comparison of differences among mRNA or protein expression in the SM samples from patients with RA or OA and in the synovial fibroblasts at different time points of stimulation. P values less than or equal to 0.05 were considered significant.

RESULTS

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

Transcriptional expression of PDGF isoforms and receptors.

The mRNA expression of the PDGFRs and their ligands in SM samples was examined by real-time RT-PCR. Both receptors were expressed at relatively low, but similar, levels, with no significant differences between OA (mean ± SD fold expression relative to GAPDH for PDGFRα, 8.5 × 10−3 ± 2.1 × 10−3, and for PDGFRβ, 4.9 × 10−3 ± 1.9 × 10−3; n = 10) and RA (for PDGFRα, 4.3 × 10−3 ± 1.0 × 10−3, and for PDGFRβ 1.3 × 10−3 ± 3.0 × 10−4; n = 9).

PDGF-A was detectable at only very low levels in 2 OA patients and 1 RA patient (0.9 × 10–4 to 5 × 10–4–fold expression relative to GAPDH). The most strongly expressed isoform was PDGF-B; its expression was not significantly different in OA (518.9 ± 289.2; n = 10) versus RA (226.7 ± 38.3; n = 9) samples. The 2 recently discovered PDGFR ligands, PDGF-C and PDGF-D, were found to be expressed at moderate levels. Whereas the amounts of PDGF-C transcripts were comparable in OA and RA (Figure 1A), the expression of PDGF-D in RA was significantly lower that than in OA (P = 0.018; Figure 1B).

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Figure 1. Real-time reverse transcription–polymerase chain reaction quantification of A, platelet-derived growth factor C (PDGF-C) and B, PDGF-D mRNA expression in synovial membranes. Individual (circles) and mean (bars) expression values (in relation to GAPDH) are shown for samples from patients with osteoarthritis (OA; n = 10) and rheumatoid arthritis (RA; n = 9). ∗ = P = 0.018 versus OA, by Mann-Whitney U test.

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Protein expression of PDGF-C and PDGF-D.

Because of the novelty and/or significantly different mRNA expression in OA and RA SM, the expression of these proteins was also investigated in SM lysates by immunoblot analysis. Full-length PDGF-C was detected in both OA and RA SM as a band of 46 kd (Figure 2A, top), which was at similar levels in the 2 diseases compared with β-actin (Figure 2B, left). Although there was a lower expression of mRNA for PDGF-D in the SM of RA patients, a significantly stronger signal for the full-length PDGF-D protein was observed compared with that in OA (Figure 2A, bottom), which is also shown in relation to β-actin (Figure 2B, right) (P = 0.018). Furthermore, the 35-kd bands (open arrowhead in Figure 2A, bottom), representing the protease-cleaved, and therefore activated, PDGF-D, was detected at significantly higher levels in RA patients than in OA patients (Figure 2B, right) (P = 0.018).

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Figure 2. Protein expression of PDGF-C and PDGF-D in lysates of synovial membranes from RA (n = 5–6) and OA (n = 6) patients. A, Western blot analysis of PDGF and the respective bands for β-actin. B, Quantification of bands (in relation to β-actin), where bars show the mean expression, and circles show individual expression. Solid arrowheads indicate molecular mass standards; open arrowhead indicates the activated form of PDGF-D. ∗ = P = 0.018 versus OA, by Mann-Whitney U test. See Figure 1 for definitions.

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Immunolocalization of PDGF-C and PDGF-D in the SM.

PDGF-C and PDGF-D were expressed throughout all compartments of the RA SM (i.e., in the synovial lining layer, in diffuse infiltrates, as well as in the stroma and around microvessels) (Figures 3A and B). In terms of tissue localization, comparable results for PDGF-C and PDGF-D were obtained with RA SM (Figures 3A and C) and OA SM (Figures 3B and D). Double-staining of RA SM showed that PDGF-C and PDGF-D were colocalized with CD90+ synovial fibroblasts within the stroma and diffuse infiltrates (Figures 3E and F), as well as with CD14+ macrophages within the synovial infiltrates (Figures 3G and H). In addition, a strong double-staining was detected for endothelial cells around the microvessels, known to be CD90+ in the RA SM (Figures 3E and F).

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Figure 3. Immunohistochemistry analysis of PDGF-C and PDGF-D in synovial tissue cryosections from a representative patient with RA (A, B, E, F, G, and H) and a representative patient with OA (C and D). Staining for PDGF-C (A and C) or PDGF-D (B and D) (brown staining) is observed in the synovial lining layer (l), diffuse cell infiltrates (d), and stroma (s) and around microvessels (v) (C and D were counterstained with hematoxylin). To characterize the cells expressing PDGF-C (E and G) and PDGF-D (F and H) (brown staining), sections of RA synovial membrane were counterstained with cell-specific antibodies to fibroblasts (E and F) and macrophages (G and H) (blue staining). Arrowheads indicate double-labeled cells. Bars in A and C = 50 μm (for A–D); bar in F = 20 μm (for E–H). See Figure 1 for definitions.

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Proliferative response.

The addition of the PDGF-DD core protein to cultured synovial fibroblasts purified from RA SM significantly stimulated the proliferation in a dose-dependent manner (maximum 5.5-fold), starting at a concentration of 50 ng/ml (Figure 4A). This demonstrated that fibroblasts from RA synovial tissue show a proliferative response to this PDGF isoform.

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Figure 4. A, Proliferative response of synovial fibroblasts from rheumatoid arthritis (RA) patients (n = 3) to stimulation with platelet-derived growth factor D (PDGF-D) core protein, as measured by bromodeoxyuridine (BrdU) incorporation. Values are the mean ± SEM. B, Influence of PDGF-D core protein on the expression of mRNA for matrix metalloproteinase 1 (MMP-1) in synovial fibroblasts from RA patients (n = 3), as quantified by real-time reverse transcription–polymerase chain reaction. Bars show the mean expression, and circles show individual expression. + = P ≤ 0.05 versus unstimulated control, by Mann-Whitney U test.

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Stimulation of MMP-1 expression.

The influence of PDGF-D on the expression of a matrix-remodeling molecule (MMP-1) was also investigated. At a concentration of 100 ng/ml, recombinant PDGF-DD core protein significantly increased the expression of MMP-1 mRNA in RA synovial fibroblasts by 2.5-fold (Figure 4B).

DISCUSSION

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

PDGFs are thought to play an important role in the synovial hyperplasia that is characteristic of RA. These effects are triggered by binding to PDGFR α and β chains, which results in activation of protooncogenes and subsequent proliferation of synoviocytes. This study is the first to demonstrate that, besides the classic PDGF-A and PDGF-B, the recently discovered isoforms PDGF-C and PDGF-D are expressed in the SM of RA and OA patients and that PDGF-D protein is detected at significantly higher levels in RA than in OA.

Interestingly, there was a discrepancy between the expression of mRNA and protein for PDGF-D in the SM of RA patients. However, similar discrepancies have been reported for the proteinase inhibitor maspin (23) and the jun/fos protooncogenes (24). These discrepancies were observed exclusively in SM from RA patients but not OA patients, suggesting disease-specific alterations of posttranscriptional processes.

PDGF-C and PDGF-D were detected in the synovial lining layer, diffuse infiltrates, and stroma of the SM, according to the expression previously reported for PDGF-A and PDGF-B (13, 15); this leads to the assumption that cells of these compartments are able to express all isoforms of PDGF. Double-labeling experiments with specific markers showed expression of both PDGF-C and PDGF-D by synovial fibroblasts and macrophages, which is consistent with recent findings of PDGF-C in dermal and lung fibroblasts (25) and in infiltrating inflammatory CD11b+ cells in coxsackievirus B3–induced chronic myocarditis (26).

PDGF-C and PDGF-D, although functional analogues of PDGF-A (2) and PDGF-B (3, 4), represent a new subfamily of growth factors that require proteolytic activation for receptor binding. This cleavage of the CUB domain results in the appearance of a 23-kd (PDGF-C) and a 32-kd (PDGF-D) protein under reducing conditions, representing the “core” growth factor domain monomers (2–4, 25). Although the respective bands were usually absent in the case of PDGF-C, active PDGF-D could be detected in all SM samples investigated (at the predicted molecular mass of 32 kd) and showed a significantly higher expression in RA SM. This is consistent with the increased proteolytic activity in the RA SM (for review, see ref. 27), in particular, with respect to the activity of plasmin, thrombin, and tissue plasminogen activator or urokinase plasminogen activator, which are reported to activate PDGF-C (2, 28) and PDGF-D (29), respectively.

Since little is known about the detailed functional roles of PDGF-C and PDGF-D, their importance in the pathogenesis of arthritic diseases remains speculative. Similar to PDGF-B, PDGF-D is important for cellular transformation and support of tumor development, probably due to exclusive or preferential signaling through PDGFRβ (8, 30). A similar function has been ascribed to the PDGFRα agonist PDGF-C (5, 6). Although signaling through PDGFRα alone is not able to efficiently transform fibroblastic cells (1), the transforming action of PDGF-C may be explained by the activation of PDGFRα/β heterodimers (25). Because both PDGFRs are expressed in the rheumatoid synovium (14), it is conceivable that binding of PDGF-D, and possibly also PDGF-C, to their receptors contributes to the proliferation and semitransformation of cells that is observed during pannus formation in RA (11). This hypothesis is further supported by the finding that the PDGF-D core protein stimulated the proliferation of synovial fibroblasts, as was previously shown for vascular fibroblasts transfected with the PDGF-D gene (31).

Recent reports indicate the involvement of both factors in fibrosis (2) or tissue remodeling (32), processes characterized by the synthesis of matrix molecules and the activity of MMPs. Indeed, in the present study, stimulation with PDGF-D increased the expression of MMP-1 mRNA in RA synovial fibroblasts. Similar results for PDGF-D have been reported in the case of MMP-2 in vascular smooth muscle cells (31), for MMP-9 in renal carcinoma cells (33), as well as for PDGF-C with respect to MMP-1 expression in dermal fibroblasts (32). Taken together, these findings indicate the relevance of the protease-activated PDGF isoforms in tissue remodeling, a process of major importance during the progression of arthritic diseases.

Acknowledgements

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

Drs. A. Roth, R. Fuhrmann, and R. Winter (all from the Orthopedic Clinic, Waldkrankenhaus “Rudolf Elle,” Eisenberg, Germany) are gratefully acknowledged for providing synovial tissue and Dr. E. Palombo-Kinne for critical revision of the manuscript. We thank Prof. F.-D. Böhmer (Institute of Molecular Cell Biology, Jena, Germany) for support and Prof. U. Eriksson (Ludwig Institute for Cancer Research, Stockholm, Sweden) for providing the PDGF-D core protein.

REFERENCES

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