Introduction
Rheumatoid arthritis (RA) is a multi-systemic autoimmune disease of unknown etiology that is characterized by hyperplastic synovial membrane capable of destroying adjacent articular cartilage and bone [
1,
2]. The pathology of RA synovial membrane includes infiltration of inflammatory leukocytes, proliferation of synovial cells and extensive angiogenesis, which is collectively referred to as the rheumatoid pannus [
2‐
4]. A critical phenomenon occurring in the early stages of synovial inflammation is angiogenesis [
4,
5], which commences with the activation of endothelial cells by a variety of stimuli, including pro-inflammatory cytokines and growth factors such as vascular endothelial growth factor (VEGF). The affected endothelial cells (ECs) then begin to digest the basement membrane, proliferate, migrate and eventually differentiate to form a tubular structure [
6].
Fas (also known as CD95) was initially discovered as a cell surface molecule that efficiently triggers death signals when bound to its ligand, Fas ligand (FasL) [
7,
8]. Fas is ubiquitously expressed, whereas FasL is principally expressed on activated T cells [
7], natural killer cells [
9], tumor cells [
10], and in immune privileged sites such as the eye [
11]. The Fas-FasL interaction plays a pivotal role in activation-induced cell death of T lymphocytes [
12], and is responsible for the cytotoxicity of T lymphocytes [
13,
14] and natural killer cells [
9]. Consequently, Fas and FasL are crucial components of lymphocyte homeostasis. In addition to the homeostatic regulation of the immune system, Fas and FasL are involved in tumor surveillance [
15,
16]. Moreover, Fas and FasL are thought to inhibit angiogenesis by inducing apoptosis of either ECs or leukocytes that provide angiogenic growth factors [
17‐
20], although one study reported an increase in angiogenesis by Fas and FasL [
21]. Similar to tumor necrosis factor (TNF)-α, FasL is cleaved from the cell surface by a metalloproteinase [
22]. The released form of FasL, soluble FasL (sFasL), was originally thought to induce apoptosis in a manner similar to membrane-associated FasL (mFasL) [
23]. However, there have been many subsequent reports upholding the differences between sFasL and mFasL regarding apoptosis induction [
24,
25]. Despite the numerous studies on the role of Fas and FasL in immune homeostasis, the effect of sFasL on the angiogenic process of RA remains to be determined.
In this study, we tested whether sFasL can regulate angiogenesis and apoptosis of rheumatoid synoviocytes. We demonstrate here that sFasL potently decreased VEGF165 production by RA fibroblast-like synoviocytes (FLSs) by inducing apoptosis in vitro. In addition, sFasL effectively inhibited VEGF165-induced migration and chemotaxis of ECs, although it did not affect tube formation by ECs. The effect of sFasL on ECs was not due to Fas-mediated cell death, since sFasL did not change either spontaneous or VEGF165-stimulated EC proliferation or survival. Moreover, sFasL strongly inhibited neovascularization in the Matrigel plug in vivo. Taken together, sFasL inhibits angiogenesis within RA synovium not only by inducing apoptosis of VEGF165-producing cells such as FLSs, but also by blocking VEGF165-induced migration of ECs, independent of Fas-mediated apoptosis.
Materials and methods
Culture of RA synoviocytes and collection of synovial fluids
The RA FLSs were prepared from the synovial tissues of ten RA patients that were undergoing total joint replacement surgery, as described previously [
26]. The mean age of the RA patients (9 females and 1 male) was 48.3 years. Eight patients had a positive rheumatoid factor. Osteoarthritis (OA) FLSs, isolated from 5 female OA patients (mean age 66.2 years), were used as a control. Synovial tissues were minced into 2 to 3 mm pieces, and treated for 4 hours with 4 mg/ml of type I collagenase (Worthington Biochemical, Freehold, NJ, USA) in DMEM at 37°C in a 5% CO
2 atmosphere. Dissociated cells were then resuspended in DMEM, supplemented with 10% FCS, 2 mM glutamine, penicillin and streptomycin, and then plated in 75 cm
2 flasks. After overnight culturing, the non-adherent cells were removed and adherent cells cultivated in DMEM plus 10% FCS. Cultures were kept at 37°C in a 5% CO
2 atmosphere, and the medium was replaced every 3 days. At confluence, the cells were passed by diluting 1:3 with fresh medium and re-cultured until used. Synoviocytes, from passages 4 through 8, were used for each experiment. The FLSs were washed in DMEM and then incubated for an additional 24 hours in serum-free DMEM supplemented with insulin-transferrin-selenium A (ITSA; Life Technologies, Grand Island, NY, USA). The cells (3 × 10
4 cells/well) were seeded in triplicate into 24-well plates (Nunc, Roskilde, Denmark) in serum-free DMEM (supplemented with ITSA) without or with transforming growth factor (TGF)-β (PeproTech, London, UK) in the presence of sFasL (10 to 100 ng/ml; MBL, Nagoya, Japan). After the indicated hours of incubation, cell viability was tested by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay, and the cell-free media were collected to measure VEGF
165 concentration in the culture supernatants. Synovial fluid was obtained with informed consent from RA and OA patients with joint effusion, and stored at -20°C in a refrigerator. All samples were obtained according to the guidelines approved by the Ethics Committee of the Catholic University of Korea.
ELISA for sFas and VEGF165
The amount of sFasL and VEGF
165 was measured by ELISA, as previously described [
27]. Recombinant human sFasL and VEGF
165 (R & D, Minneapolis, MN, USA) were used as a calibration standard. A standard curve was drawn by plotting the optical density versus the log of the concentration of sFas and VEGF
165.
Isolation and culture of endothelial cells
The ECs were isolated from normal-term umbilical cord vein by collagenase digestion, and then grown to confluence in 75 cm2 flasks containing M199 medium (Life Technologies) supplemented with 20% FCS, 100 U/ml penicillin, 100 μg/ml streptomycin, and 2 mM L-glutamine. Cultures were kept at 37°C in a CO2 incubator, and the medium was changed every 2 to 3 days until confluence was reached. ECs were passed with 0.2% collagenase and 0.02% EDTA (Life Technologies); cells from passages 2 to 3 were used in this study.
Determination of endothelial cell proliferation and viability
The proliferation and survival of ECs were assessed by [3H]-thymidine incorporation assay and MTT assay, respectively. Briefly, ECs were incubated for 24 hours in DMEM supplemented with 1% FCS. The medium was replaced with fresh DMEM/ITSA supplemented with 1% FCS, and the cells were incubated without or with 10 ng/ml of VEGF165 in the presence of sFasL (1 to 100 ng/ml) for 24 hours. For the determination of the proliferation rate of ECs, 1 μCi of [3H]-thymidine was added to each of the wells prior to the final 6 hours of culturing, and the incorporated radioactivity was counted with a scintillation counter.
Wounding migration and tube formation assay
The wounding migration and tube formation activity of ECs were measured as described previously [
28]. In brief, ECs plated at confluence on 60 mm culture dishes were wounded with pipette tips, and then treated with VEGF
165 (20 ng/ml) in M199 medium, supplemented with 1% FCS and 1 mM of thymidine. After 12 hours of incubation, migration was quantified by counting the cells that had moved beyond the reference line. For the tube formation assay, ECs were seeded on a layer of previously polymerized Matrigel (BD Biosciences, San Jose, CA, USA) with VEGF
165 (20 ng/ml). After 18 hours of incubation, the cell morphology was visualized via phase-contrast microscopy and photographed. The degree of tube formation was quantified by measuring the length of tubes in 5 randomly chosen low-power fields (×40) from each well using image-Pro Plus v4.5 (Media Cybernetics, San Diego, CA, USA).
Chemotaxis assay of endothelial cells
The chemotactic migration of ECs was assayed using a Transwell chamber with 6.5 mm diameter polycarbonate filters (8 μm pore size). In brief, the filter's lower surface was coated with 10 μg of gelatin. VEGF165 (10 ng/ml), which was prepared in M199 medium containing 1% FCS, was placed in the lower wells. The ECs that were incubated in M199 with 1% FCS for 6 hours or overnight were trypsinized and suspended at a final concentration of 1 × 106 cells/ml in M199 containing 1% FCS. Various concentrations of sFasL or soluble CD40 ligand (sCD40L; R & D) were added to the upper wells with 100 μl of cell suspensions. The chamber was incubated at 37°C for 4 hours. The cells were fixed and stained with hematoxylin and eosin. Non-migrating cells on the filter's upper surface were removed by wiping with a cotton swab. Chemotaxis was quantified by counting the cells that migrated to the lower side of the filter by optical microscopy at ×200 magnification. Eight random fields were counted for each assay. Each sample was assayed in duplicate.
Western blot analysis for phospho-Akt and phospho-ERK
ECs were incubated for 24 hours in DMEM with 1% FCS, and then VEGF165 (20 ng/ml) plus various concentrations of sFasL (1 to 50 ng/ml) were added to the cells for the indicated times. The treated ECs were then washed twice in PBS, dissolved in sample buffer (50 mM Tris-HCl, 100 mM NaCl, 0.1% SDS, 1% NP-40, 50 mM NaF, 1 mM Na3VO4, 1 μg/ml aprotinin, 1 μg/ml pepstatin, and 1 μg/ml leupeptin), boiled, separated via SDS-PAGE, and transferred to nitrocellulose membranes. After immunoblot analysis with anti-phospho-ERK1/2 (Thr202/Tyr204) or anti-phospho-Akt (Ser473), the membranes were stripped and re-incubated with β-actin antibody in order to detect total protein amounts.
Mouse Matrigel plug assay
Matrigel (500 μl) containing VEGF
165 (500 ng/ml) and heparin (9 U/ml) were injected subcutaneously with or without sFasL (100 ng/ml) into the abdomen of C57BL/6 mice (7 weeks of age), as described previously [
28]. After 14 days, the skins of the mice were pulled back to expose the Matrigel plugs, which remained intact. After noting and photographing any quantitative differences, hemoglobin levels were measured by the Drabkin method, using a Drabkin reagent kit 525 (Sigma, St. Louis, MO, USA). The hemoglobin concentration was calculated from the parallel assay of a known amount of hemoglobin. The Matrigel plugs were fixed in 4% formalin, embedded with paraffin, and stained using hematoxylin and eosin.
Statistical analysis
Data are expressed as the mean ± standard deviation (SD). Comparisons of the numerical data between groups were performed by paired or unpaired Mann-Whitney U-test. P values less than 0.05 are considered statistically significant.
Discussion
FasL is a type II membrane protein of approximately 280 amino acids that belongs to the TNF/nerve growth factor family, which includes TNF, lymphotoxin, CD40L, and TRAIL. FasL induces apoptotic death in cells expressing its receptor, Fas [
17,
35]. FasL undergoes proteolytic cleavage in its extracellular domains, resulting in the release of sFasL [
22]. In general, sFasL is less potent at inducing apoptosis than membrane-bound Fas, as shown in a variety of cell types [
24,
25,
36]. Elevated levels of sFasL are found in sera from patients with atherosclerosis [
37], leukemia [
38], and acute graft-versus host disease [
39]. The concentration of sFasL in the synovial fluid is higher in patients with severe RA than in those with mild RA or OA [
27]. Moreover, local injection of sFasL into the affected joints suppresses experimental arthritis in rats [
40], suggesting it has therapeutic potential in RA.
Angiogenesis has been considered a critical step in the progression of chronic arthritis, as well as an early determinant in the development of RA [
41]. In this study, we first identified a novel mechanism for anti-angiogenesis in RA involving sFasL. sFasL decreased VEGF
165 production from RA FLSs by inducing apoptosis
in vitro. The apoptotic action of sFasL seems to be specific to RA FLSs because it minimally affected the viability of OA FLSs, which is consistent with the previous finding that OA FLSs are less sensitive to Fas-mediated apoptosis [
42]. In addition, sFasL drastically suppressed VEGF
165-induced migration and chemotaxis of ECs
in vitro and also blocked neovascularization
in vivo, although it did not alter the proliferation and tube formation of ECs, indicating that sFasL displays anti-angiogenic activity through at least two different mechanisms: induction of Fas-mediated cell death of VEGF
165-producing cells; and apoptosis-independent inhibition of EC migration and chemotaxis. These data, together with earlier reports [
15,
40], suggest that administration of sFasL could be effective in treating several angiogenesis-dependent diseases, such as cancer and chronic inflammatory diseases, explaining how sFasL protects against the development of experimental arthritis.
RA FLSs are susceptible to anti-Fas IgM and undergo apoptosis
in vitro [
43,
44]. By contrast, apoptotic cells are rarely observed in the RA synovium
in vivo [
43]. The effect of FasL on apoptosis of ECs is also still controversial; although some studies report that ECs are sensitive to FasL-induced apoptosis [
45,
46], others report the contrary [
47,
48]. In our experiments, sFasL was able to induce the apoptosis of RA FLSs and ECs in a culture condition without FCS (Figure
2b; data not shown). However, when 1% FCS was added to the medium, sFasL-mediated cell death was partially inhibited in the cultured FLSs or did not occur in ECs (Figures
2b and
3a), indicating that the apoptotic action of sFasL on VEGF-secreting cells, such as FLSs, was hampered by growth factors. Recently, we have demonstrated that VEGF
165 protects FLSs from apoptotic death by regulating Bcl-2 expression and Bax translocation [
49]. Therefore, it seems unlikely that sFasL greatly elicits apoptosis of FLSs and ECs in the joints with high levels of VEGF
165 or other growth factors. Instead, a mechanism by which sFasL induces anti-angiogenesis by blocking migration and chemotaxis of ECs may be more relevant in this specific condition.
Fas ligation activates a pro-inflammatory program in some cell types independently of apoptotic responses [
50‐
52]. For example, Fas stimulation increased expression of IκBα, matrix metalloproteinases and chemokines, and Fas-activated RA FLSs displayed increased chemotactic activity for monocytic cells [
50]. In our study, Fas ligation by sFasL did not affect the production of chemokines, such as IL-8 and monocyte chemoattractant protein-1, by ECs (data not shown), suggesting that Fas ligation does not trigger chemotactic signals for ECs. On the contrary, sFasL treatment blocked the migration and chemotaxis of ECs induced by VEGF
165 (Figure
4). Moreover, sFasL strongly inhibited VEGF
165-induced pAkt activity, but not pERK activity, in ECs (Figure
5). It is unclear how sFasL regulates VEGF
165-induced pAkt activity and then suppresses the chemotaxis and migration of ECs. Activation of the phosphatidylinositol 3-kinase/protein kinase Akt pathway mediates nitric oxide-induced EC migration and angiogenesis [
53]. Conversely, inhibition of the Akt pathway results in anti-angiogenic effects through the inhibition of EC migration in an apoptosis-independent manner [
54,
55]. Given that the activities of Flt-1 and its downstream target pAkt are critical to VEGF
165 signaling for chemotaxis [
53,
54], the anti-migratory action of sFasL may be mediated by blocking VEGF
165-induced upregulation of pAkt activity. Further study is required to determine the molecular mechanisms for sFasL regulation of pAkt activity linked to chemotaxis control.
Competing interests
The authors declare that they have no competing interests.
Authors' contributions
WK participated in the design of the study, drafted the manuscript and analyzed the data. SK helped to draft the manuscript. KH carried out cell cultures and ELISA, and helped to perform migration, chemotaxis, and tube formation assays of ECs. JK and SU assisted in cell cultures and performed migration, chemotaxis, and tube formation assays of ECs, and the in vivo Matrigel assay. JC participated in the design of the study and the analysis of data. CC conceived of the study, participated in its design and coordination, and helped to draft the manuscript. All authors read and approved the final manuscript.