Lysophosphatidic acid mediates migration of human mesenchymal stem cells stimulated by synovial fluid of patients with rheumatoid arthritis

Biochimica et Biophysica Acta 1801 (2010) 23–30

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Biochimica et Biophysica Acta j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / b b a l i p

Lysophosphatidic acid mediates migration of human mesenchymal stem cells stimulated by synovial fluid of patients with rheumatoid arthritis Hae Young Song a, Mi Jeong Lee a, Min Young Kim a, Kyung Hye Kim a, Il Hwan Lee a, Sang Hun Shin a, Jung Sub Lee b,⁎, Jae Ho Kim a,⁎ a b

Medical Research Center for Ischemic Tissue Regeneration, the Medical Research Institute, School of Medicine, Pusan National University, Busan, Republic of Korea Department of Orthopaedic Surgery, School of Medicine, Pusan National University, Busan, Republic of Korea

a r t i c l e

i n f o

Article history: Received 20 March 2009 Received in revised form 17 July 2009 Accepted 26 August 2009 Available online 3 September 2009 Keywords: Lysophosphatidic acid Mesenchymal stem cell Migration Synovial fluid Arthritis

a b s t r a c t Migration of mesenchymal stem cells plays a key role in regeneration of injured tissues. Rheumatoid arthritis (RA) is a chronic inflammatory disease and synovial fluid (SF) reportedly contains a variety of chemotactic factors. This study was undertaken to investigate the role of SF in migration of human bone marrow-derived mesenchymal stem cells (hBMSCs) and the molecular mechanism of SF-induced cell migration. SF from RA patients greatly stimulated migration of hBMSCs and the SF-induced migration was completely abrogated by pretreatment of the cells with the lysophosphatidic acid (LPA) receptor antagonist Ki16425 and by small interfering RNA- or lentiviral small hairpin RNA-mediated silencing of endogenous LPA1/Edg2. Moreover, SF from RA patients contains higher concentrations of LPA and an LPA-producing enzyme autotoxin than normal SF. In addition, SF from RA patients increased the intracellular concentration of calcium through a Ki16425-sensitive mechanism and pretreatment of the cells with the calmodulin inhibitor W7 or calmodulin-dependent protein kinase II inhibitor KN93 abrogated the SF-induced cell migration. These results suggest that LPA-LPA1 plays a key role in the migration of hBMSCs induced by SF from RA patients through LPA1-dependent activation of calmodulin-dependent protein kinase II. © 2009 Elsevier B.V. All rights reserved.

1. Introduction Rheumatoid arthritis (RA) is a chronic inflammatory disease characterized by the destruction of articular cartilage and adjacent bone tissues [1] and disordered synovial microenvironment, including infiltration of inflammatory cells, hyperplasia of stromal cells, and tissue scarring [2]. The pathological events are mediated by a complex interplay of pro-inflammatory cytokines and mediators produced in the joint tissues or synovium of patients with RA [3]. It has been reported that synovial fluid (SF), which nourishes articular cartilage and lubricates articular joint surfaces [4], contains various growth

Abbreviations: RA, rheumatoid arthritis; SF, synovial fluid; MSCs, mesenchymal stem cells; LPA, lysophosphatidic acid; ATX, autotaxin; [Ca2+]i, intracellular concentration of calcium; hBMSCs, human bone marrow-derived mesenchymal stem cells; αMEM, α-minimum essential medium; PTX, pertussis toxin; PDGF-BB, platelet-derived growth factor-BB; 1-oleoyl-LPA, 1-oleoyl-sn-glycero-3-phosphate; S1P, sphingosine-1phosphate; OA, osteoarthritis; RT-PCR, reverse transcription–polymerase chain reaction; siRNA, small interfering RNA; shRNA, small hairpin RNA; RA-SF, SF from RA patients; CaMK II, calmodulin-dependent protein kinase II ⁎ Corresponding authors. J.H. Kim, Department of Physiology, School of Medicine, Pusan National University, Yangsan 626-813, Gyeongsangnam-do, Republic of Korea. J.S. Lee, Department of Orthopaedic Surgery, School of Medicine, Pusan National University, Busan 602-739, Republic of Korea. E-mail addresses: [email protected] (J.S. Lee), [email protected] (J.H. Kim). 1388-1981/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.bbalip.2009.08.011

factors and chemotactic factors which play a key role in the pathogenesis of RA [5]. Mesenchymal stem cells or multipotent stromal cells (MSCs) can be isolated from a variety of tissues, including bone marrow, adipose tissue, peripheral blood, articular cartilage, and synovial tissue [6–10]. MSCs possess self-renewal capacity, long-term viability, and differentiation potential toward diverse cell types, such as adipogenic, osteogenic, chondrogenic, and myogenic lineages [7,8,11,12], suggesting potential application of MSCs for regenerative medicine. MSCs have been reported to reside in the SF of patients with arthritis [13] and accumulate in the synovium of collagen-induced arthritis animal model [14]. However, the molecular identities of the factors involved in the chemotactic migration of MSCs into the synovium have not been clarified. Lysophosphatidic acid (LPA) is a naturally occurring bioactive lipid belonging to the family of phospholipid growth factors, present in micromolar concentrations in serum and biological fluids and in higher concentrations at sites of inflammation [15]. It has been reported that SF from RA patients contains LPA and autotaxin (ATX) [16], which is involved in the generation of LPA by hydrolyzing lysophosphatidylcholine [17]. ATX has been reported to exist in body fluids under pathological conditions, including ascetic fluid from ovarian cancer patients [18], serum from patients with chronic hepatitis C [19], and synovial fluid from RA patients [16,20], suggesting a potential role of LPA and ATX in the pathogenesis of RA. LPA is involved in a variety of

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physiological and pathophysiological responses including wound healing, production of angiogenic factors, chemotaxis, neointima formation, tumor cell invasion, metastasis, and cell cycle progression [21,22]. The biological functions of LPA are mediated through several G protein-coupled receptors, i.e., LPA1/Edg2, LPA2/Edg4, LPA3/Edg7, LPA4/p2y9/GPR23, LPA5/GPR92, GPR87, and p2y10 [23–26]. Activation of LPA receptors mediates the biological responses through activating multiple signaling pathways involving intracellular concentration of calcium ([Ca2+]i), ERK, and phosphatidylinositol-3-kinase [21,22]. We have previously reported that LPA in malignant ascites from patients with ovarian cancer induces migration of human adipose tissue-derived MSCs through LPA1-dependent mechanism [27], suggesting a key role of LPA in the migration of MSCs. However, it is still elusive whether LPA plays a key role in the migration of MSCs induced by SF from RA patients. In the present study, we sought to explore whether SF from patients with RA can induce migration of human bone marrowderived MSCs (hBMSCs). We demonstrated for the first time that LPA plays a pivotal role in the migration of hBMSCs stimulated by SF from RA patients through LPA1 receptor-mediated activation of calmodulin-dependent protein kinase II (CaMK II). 2. Materials and methods 2.1. Materials α-Minimum essential medium (α-MEM), phosphate-buffered saline (PBS), trypsin, fetal bovine serum, M-MLV reverse transcriptase, and Lipofectamine plus™ reagent were purchased from Invitrogen (Carlsbad, CA). Pertussis toxin (PTX) was from BIOMOL (Plymouth Meeting, PA). Human platelet-derived growth factor-BB (PDGF-BB) was purchased from R&D Systems (Minneapolis, MN). Fluo-4-AM was from Molecular Probes, Inc. (Eugene, OR). 1-Oleoyl-sn-glycero-3phosphate (1-oleoyl-LPA), sphingosine-1-phosphate (S1P), fatty acidfree bovine serum albumin, W7, KN92, KN93, and Ki16425 were purchased from Sigma-Aldrich (St. Louis, MO). Universal LPA assay kit was purchased from Echelon Biosciences, Inc. (Salt Lake City, UT).

were maintained at 37 °C in a humidified atmosphere containing 5% CO2 in growth medium (α-MEM, 10% fetal bovine serum, 100 units/ ml of penicillin, 100 μg/ml of streptomycin) until they reached confluence. The primary hBMSCs were subcultured in tissue culture dishes at a concentration of 2000 cells/cm2. The hBMSCs were c-kit, CD34, and CD45 negative and greater than 90% CD29, CD44, CD90, and CD105 positive were used in the experiments. 2.4. Cell migration assay Migration of hBMSCs was assayed using a Boyden chamber apparatus, as previously described [29]. Briefly, hBMSCs were harvested with 0.05% trypsin containing 0.02% EDTA, washed once, and suspended in α-MEM at a concentration of 2 × 105 cells/ml. A polycarbonate membrane filter with 8-μm pores of the disposable 96-well chemotaxis chamber (Neuro Probe, Inc., Gaithersburg, MD) was precoated overnight with 20 μg/ml rat-tail collagen at room temperature, an aliquot (50 μl) of hBMSCs suspension was loaded into the upper chamber, and test reagents were then placed in the lower chamber, unless otherwise specified. For elucidation of signaling pathways involved in the LPA-induced migration, the cells were preincubated with pharmacological inhibitors for 15 min before loading. After exposure of the cells to either LPA or SF in the absence or presence of inhibitors for 12 h at 37 °C, the filters were then disassembled, and the upper surface of each filter was scraped free of cells by wiping it with a cotton swab. The number of cells that had migrated to the lower surface of each filter was determined by counting the cells in four places under microscopy at ×100 magnification after staining with hematoxylin and eosin. 2.5. Reverse transcription–polymerase chain reaction (RT-PCR)

SF was obtained with the patient's consent, as approved by the Institution Review Board of Busan National University Hospital. SF was obtained from 11 patients with RA (7 male, 4 female; mean age of 59± 10 years) and 10 patients with osteoarthritis (OA) (5 male, 5 female; 61 ± 13 years) during therapeutic arthrocentesis. SF from normal donors was obtained postmortem from 10 organ donors (4 male, 6 female; mean age of 55± 15 years) without joint diseases. SF was transferred to heparin-treated tubes, transported immediately to the laboratory, and centrifuged at 3000 ×g for 10 min at 4 °C to remove possible inflammatory cells and blood cells. Aliquots of the supernatants were used immediately or stored at −80 °C for future analysis. SF aliquots were either used immediately or subjected to one freeze–thaw cycle. To prevent ATX-mediated generation of LPA during preparation of SF, all steps were carried out at 4 °C. To denature proteins in SF, an aliquot (200 μl) of SF or 1-oleoyl-LPA was heated at 95 °C for 5 min and centrifuged at 15,000 rpm for 5 min to remove denatured proteins, and the supernatants were collected.

Total cellular RNA was extracted by the Trizol method (Invitrogen, Carlsbad, CA). For RT-PCR analysis, aliquots of 2 μg each of RNA were subjected to cDNA synthesis with 200 U of M-MLV reverse transcriptase and 0.5 μg of oligo (dT) 15 primer (Promega, Madison, WI). The cDNA in 2 μl of the reaction mixture was amplified with 0.5 U of GoTaq DNA polymerase (Promega, Madison, WI) and 10 pmol each of sense and antisense primers as follows: LPA1 (384 bp product): sense 5′TCTTCTGGGCCATTTTCAAC-3′, antisense 5′-TGCCTRAAGGTGGCGCTCAT-3′; LPA2 (780 bp product): sense 5′-CCTACCTCTTCCTCATGTTC-3′, antisense 5′-TAAAGGGTGGAGTCCATCAG-3′; LPA3 (450 bp product): sense: 5′-GGAATTGCCTCTGCAACATCT-3′, antisense 5′-GAGTAGATGATGGGGTTCA-3′; LPA4 (200 bp product): sense 5′-TACAACTTCAACCGCCACTG-3′, antisense 5′-ATCCAGACACCAGCACACAC-3′; LPA5/GPR92 (261 bp product): sense 5′-GTGCTGATGGTGATGGTGCT-3′, antisense 5′-TGTGAAGGAAGACAGAGAGTGG-3′; GPR87 (536 bp product): sense 5′-CCGTATGAGGTGAATGGACA-3′, antisense 5′-CCAAGGAACACGATGGAAGT-3′; p2y10 (434 bp product): sense 5′-TCTTCTTCATCTGCTTCACTCC-3′, antisense 5′-CTCTGCCTTCACCATCACAC-3′; glyceraldehyde3-phosphate dehydrogenase (GAPDH): sense 5′-TCCATGACAACTTTGGTATCG-3′, antisense 5′-TGTAGCCAAATTCGTTGTCA-3′. The thermal cycle profile was as follows: denaturation at 95 °C for 30 s, annealing at 54–55 °C for 30 s depending on the primers used, and extension at 72 °C for 40 s. Each PCR reaction was carried out for 30 cycles, and PCR products were size fractionated on 1.2% ethidium bromide/agarose gel and photographed under UV transillumination.

2.3. Cell culture

2.6. Transfection with small interfering RNA

After informed consent, heparinized bone marrow cells were obtained from different individuals undergoing total hip arthroplasty and hBMSCs were isolated as previously described [28]. To isolate hBMSCs, mononuclear cells from bone marrow were separated by centrifugation in a Ficoll–Hypaque gradient (density = 1.077 g/cm3; Sigma) and seeded at a concentration of 1 × 106 cells/cm2. Cultures

Small interfering RNA (siRNA) duplexes were synthesized, desalted, and purified by Samchully Pharm. Co. Ltd. (Siheung, GyeongGi, Korea) as follows: LPA1, 5′-GGACUUGGAAUCACUGUUUUU-3′ (sense) and 5′AAACAGUGAUUCCAAGUCCUU-3′ (antisense). Nonspecific control siRNA (D-001206-13-05) was purchased from Dharmacon, Inc. (Chicago, IL). For siRNA experiments, hBMSCs were seeded on 60-mm

2.2. Collection of synovial fluid

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dishes at 70% confluence, and they were then transfected with siRNAs by using the Lipofectamine plus™ reagent according to the manufacturer's instructions. Briefly, Lipofectamine plus™ reagent was incubated with serum-free medium for 15 min, and respective siRNAs were then added to the mixtures. After incubation for 15 min at room temperature, the mixtures were diluted with serum free medium and added to each well. The final concentration of siRNAs in each well was 100 nM. After incubation of hBMSCs to serum-free medium containing siRNAs for 4 h, the cells were cultured in growth medium for 24 h, and the expression levels of LPA1 and GAPDH were then determined by reverse transcription–polymerase chain reaction analysis. 2.7. Lentiviral small hairpin RNA (shRNA) transduction pLKO.1-puro lentiviral vectors expressing LPA1 shRNA (TRCN0000011368) or nontarget control shRNA (SHC002) were purchased from Sigma-Aldrich. The functional sequence in the LPA1 shRNA lentiviral vector is “CCGGCCTTCTGAAGACTGTGGTCATCTCGAGATGACCACAGTCTTCAGAAGGTTTTT” to target the LPA1 gene sequence (CCTTCTGAAGACTGTGGTCAT). To generate lentiviral particles, HEK293FT cells were co-transfected with the shRNA lentiviral plasmid (pLKO.1-puro) and ViraPower Lentiviral packaging mix (pLP1, pLP2, pLP-VSV-G; Invitrogen) using Lipofectamine 2000 (Invitrogen) and the culture supernatants containing lentivirus were harvested at 48 h after transfection. For lentiviral transduction, hBMSCs were treated with the shRNAexpressing lentivirus in the presence of 5 μg/ml polybrene (SigmaAldrich) and stable cell lines expressing shRNA were generated by selection with puromycin (5 μg/ml). To ensure shRNA-mediated silencing of LPA1 expression, the mRNA levels of LPA1 and GAPDH were determined by RT-PCR analysis.

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2.8. Measurement of LPA concentration in SF The LPA level was determined by a commercially available LPA assay kit (Universal LPA assay kit, Echelon Biosciences) according to the manufacturer's instructions. In brief, each well of a microtiter plate was blocked by adding blocking solution and washed with icecold PBS four times. Samples or LPA standards were mixed with biotinylated anti-LPA antibody and the mixtures (100 μl/well) were transferred to the microtiter plate. After incubation for 1 h at 4 °C, the plate was washed with ice-cold PBS and streptavidin horseradish peroxidase solution was added to each well of the microtiter plate. After 1 h, the plate was washed, and then TMB substrate was added to each well and incubated for 4 min at room temperature. The reaction was stopped by the addition of 1N H2SO4 to each well and the absorbance of each solution at 450 nm was determined by using a PowerWavex microplate spectrophotometer (Bio-Tek Instruments, Winooski, VT). The concentration of LPA in the samples is determined by comparison to the standard curve. 2.9. Western blotting SF was resolved by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE), and the separated proteins were transferred onto a nitrocellulose membrane, and then stained with 0.1% Ponceau S solution. After blocking with 5% nonfat milk, the membranes were immunoblotted with anti-ATX monoclonal antibody (3D1), which was kindly provided by Dr. Junken Aoki (Tohoku University, Japan), and the bound antibodies were visualized with horseradish peroxidaseconjugated secondary antibody using an enhanced chemiluminescence system.

Fig. 1. RA-SF patients stimulate migration of hBMSCs. Cell migration was determined by a Boyden chamber apparatus as described in Materials and methods. (A) hBMSCs (1 × 104 cells/50 μl in serum-free α-MEM) were loaded into the upper chamber and serum-free α-MEM containing indicated concentrations of RA-SF (case 1) was placed in the lower chamber. The number of cells migrated across a polycarbonate membrane filter was counted after 12 h. (B) hBMSCs were exposed to serum-free medium in the absence (w/o) or presence (RA) of 5% RA-SF for indicated time periods. Data represent mean ± SD (n = 4). ⁎, p b 0.01 by two-way ANOVA and Scheffe's post hoc test. (C) hBMSCs were exposed to serum-free medium in the absence (w/o) or presence of 5% SF from patients with RA or OA or normal donors (each 10 cases) for 12 h. Data represent mean ± SD (n = 3). (D) Statistical analysis of cell migration. Box and whisker plots represent median values, 25–75% range, and 10–90% range. ⁎, p b 0.05 vs. normal; #, p b 0.05 vs. OA by one-way ANOVA and Scheffe's post hoc test.

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Fig. 2. Role of Gi in the RA-SF-induced migration of hBMSCs. (A) To denature protein factors, SF from RA patients or normal donors was heated at 95 °C for 5 min. hBMSCs were then exposed to heated or unheated SF from RA patients or normal donors (each 5% concentration) for 12 h, and the number of migrating cells was counted under microscopy. (B) hBMSCs were pretreated with serum-free medium containing vehicle or 100 ng/ml PTX for 24 h, loaded into upper chamber, and exposed to 0.5 μM 1-oleoyl-LPA or 5% SF from RA patients or normal donors for 12 h for determination of cell migration. Data represent mean ± SD (n = 4). ⁎, p b 0.05; ⁎⁎, p b 0.01 by two-way ANOVA and Scheffe's post hoc test.

2.10. Measurement of intracellular calcium concentration Spatially averaged photometric [Ca2+] measurement from single cells was performed with the fluorescent Ca2+ indicator fluo-4-AM. In brief, hBMSCs grown on 32-mm dish were incubated with serum-free α-MEM for 24 h, loaded with 5 μM fluo-4-AM for 40 min at 37 °C in buffer A [135 mM NaCl, 4 mM KCl, 1 mM MgCl2, 2 mM CaCl2, 10 mM glucose, and 20 mM N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid (HEPES), pH 7.3], and washed twice with Hanks' balanced salt solution without phenol red and Ca2+. The fluo-4-AM-loaded hBMSCs were treated with agonists in the absence or presence of Ki16425. A Leica TCS-SP2 laser scanning confocal microscope (Leica Microsystems, Germany) was used to visualize Ca2+-mediated fluorescence in the cells. Fluo-4 was excited with 488-nm line of an argon laser, and fluo-4 fluorescence was collected between 510 and 525 nm. Scanning was performed every 1 s for the indicated times, and the ratio of fluorescence intensity to initial fluorescence intensity (F/F0) was calculated at each point for quantitative measurement. 2.11. Statistical analysis The results of multiple observations are presented as mean ±SD. For multivariate data analysis, group differences were assessed with two-way ANOVA, followed by post hoc comparisons tested with Scheffe's method. 3. Results 3.1. SF derived from RA patients induces migration of hBMSCs In order to explore whether SF from RA patients (RA-SF) can regulate migration of hBMSCs, SF was loaded into the lower chamber

of the Boyden apparatus, and migration of the cells from the upper chamber toward the lower chamber was determined. As shown in Fig. 1A, RA-SF dose-dependently stimulated migration of hBMSCs with a maximal stimulation at 10% concentration. Moreover, exposure of the cells to RA-SF time-dependently stimulated migration of hBMSCs (Fig. 1B). To assess whether RA-SF specifically increased migration of hBMSCs, we next compared the effects of SF from several patients with RA or OA and normal donors. As shown in Fig. 1C and D, RA-SF exhibited more potent stimulatory effects than SF from OA patients or normal donors. These results suggest that RA-SF possessed greater potential to recruit hBMSCs than SF from OA patients or normal donors. 3.2. Role of PTX-sensitive G proteins in the migration of hBMSCs induced by SF from RA patients To assess whether protein factors are responsible for the RASF-induced migration of hBMSCs, RA-SF was heated to 95 °C for 5 min for denaturation of protein factors. As shown in Fig. 2A, the stimulatory effects of RA-SF on the migration of hBMSCs were not abrogated by heating, suggesting that protein factors are not likely to be involved in the chemotactic migration of hBMSCs. To explore whether G protein-coupled receptors are involved in the migration of hBMSCs, we next examined the effects of PTX, a pharmacological inhibitor of Gi, in the RA-SF-induced migration of hBMSCs. Since we have reported that LPA stimulated migration of human adipose tissuederived MSCs through PTX-sensitive mechanism [27], we determined the effects of PTX on the LPA-induced migration of hBMSCs. As shown in Fig. 2B, LPA stimulated migration of hBMSCs, and preincubation of the cells with PTX completely abrogated the LPA-induced migration of hBMSCs. Furthermore, migration of hBMSCs induced by SF from

Fig. 3. LPA is responsible for RA-SF-induced migration of hBMSCs. (A) hBMSCs were pretreated with the indicated concentrations of Ki16425 for 15 min, and then exposed to serum-free medium containing 0.5 μM 1-oleoyl-LPA, 5% RA-SF (case 1), or 0.1 μM S1P for 12 h. (B) hBMSCs were pretreated with vehicle or 5 μM Ki16425 for 15 min, and then exposed to serum-free medium containing 0.5 μM 1-oleoyl-LPA or 10 ng/ml PDGF-BB for 12 h. (C) hBMSCs were pretreated with vehicle or 5 μM Ki16425, and then exposed to serum-free medium containing 5% SF from RA patients (four cases) for 12 h. The number of migrating cells was determined. Data represent mean ± SD (n = 4). ⁎, p b 0.01 by two-way ANOVA and Scheffe's post hoc test.

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Fig. 4. Role of LPA1 in LPA-induced migration of hBMSCs. (A) Expression levels of seven LPA receptors were determined by RT-PCR. (B) hBMSCs were transfected with either control siRNA (si-control) or LPA1-specific siRNA (si-LPA1). The mRNA levels of LPA1, LPA4, LPA5, and GAPDH were determined by RT-PCR. (C) The siRNA-transfected hBMSCs were exposed to vehicle (w/o), 0.5 μM 1-oleoyl-LPA, 5% RA-SF (case 1), or 0.1 μM S1P for 12 h, and then the number of migrating cells was determined. (D) hBMSCs were infected with either control shRNA (sh-control) or LPA1-specific shRNA (sh-LPA1) lentiviruses. The mRNA levels of LPA1, LPA4, LPA5, and GAPDH were determined by RT-PCR. (E) The shRNA-infected hBMSCs were exposed to vehicle (w/o), 0.5 μM 1-oleoyl-LPA, 5% RA-SF (case 1), or 0.1 μM S1P for 12 h, and then the number of migrating cells was determined. Data represent mean ± SD (n = 4). ⁎, p b 0.01 by two-way ANOVA and Scheffe's post hoc test.

normal or RA donors was also blocked by PTX treatment. These results suggest a pivotal role of PTX-sensitive G proteins in the migration of hBMSCs induced by not only LPA but also RA-SF. 3.3. SF from RA patients induces migration of MSCs through LPA receptor-dependent mechanism To explore whether LPA is involved in the migration of hBMSCs induced by RA-SF, we examined the effects of Ki16425, an antagonist specific for LPA1 and LPA3, on the migration of hBMSCs in response to either LPA or RA-SF. As shown in Fig. 3A, Ki16425 dose-dependently inhibited the migration of hBMSCs stimulated by either LPA or RA-SF. To ensure specificity of Ki16425, we next examined the effect of Ki16425 on cell migration induced by S1P and PDGF-BB, which have been reported to stimulate migration of MSCs [30,31]. In contrast to the potent inhibitory effect of Ki16425 on LPA-induced cell migration, S1P- or PDGF-BB-induced migration was not significantly affected by Ki16425 treatment (Fig. 3A and B). Furthermore, the migration of hBMSCs induced by SF from four different RA patients was markedly abrogated by pretreatment of the cells with Ki16425 (Fig. 3C). These results suggest that LPA plays a key role in the migration of hBMSCs induced by RA-SF.

or LPA. The mRNA levels of LPA1, but not those of other LPA receptors, in hBMSCs were specifically down-regulated by transfection with LPA1-specific siRNAs (Fig. 4B). As shown in Fig. 4C, the LPA-induced migration of hBMSCs was abrogated by depletion of the endogenous LPA1. Furthermore, the migration of the cells stimulated by RA-SF was completely inhibited by knockdown of LPA1 expression. However, S1P-induced cell migration was not abrogated by siRNA-mediated depletion of LPA1. To support the involvement of LPA1 in the RASF-induced migration, we next depleted endogenous LPA1 using lentiviral transduction of LPA1-specific shRNA (sh-LPA1). As shown in Fig. 4D, LPA1 expression in hBMSCs was specifically attenuated by lentiviral transduction of sh-LPA1, but not by control shRNA. Consistently, migration of hBMSCs induced by either LPA or RA-SF, but not S1P, was abrogated by sh-LPA1-mediated silencing of endogenous

3.4. SF from RA patients induces migration of MSCs through LPA1-dependent mechanism LPA activates several G protein-coupled receptors including LPA1/ Edg2, LPA2/Edg4, LPA3/Edg7, LPA4/p2y7/GPR23, LPA5/GPR92/ GPR93, GPR87, and p2y10 [23–26]. Appropriately, we next examined the expression levels of these LPA receptors in hBMSCs using RT-PCR. As shown in Fig. 4A, LPA1, LPA4, and LPA5 were expressed in hBMSCs. Because LPA-induced migration of hBMSCs was abrogated by Ki16425, an antagonist for LPA1 and LPA3, these results suggested the possibility that LPA1, but not LPA3, might be involved in LPA-induced cell migration. To explore the involvement of LPA1 in the SF-induced migration of hBMSCs, we examined the effects of siRNA-mediated depletion of LPA1 on the migration that was stimulated by either RA-SF

Fig. 5. Increased levels of LPA and ATX in RA-SF. (A) ATX levels in SF samples (1 μl) from normal (four cases), RA (six cases), or OA (six cases) patients were analyzed by Western blotting with anti-ATX antibody. (B) The densities of ATX were quantified from the panel (A) using a scanning densitometer. Data represent mean ± SD. ⁎, p b 0.05. (C) LPA contents in SF samples from normal (four cases), RA (six cases), or OA (six cases) patients were determined as described in Materials and methods. Data represent mean ± SD. ⁎, p b 0.05; ⁎⁎, p b 0.01 by one-way ANOVA and Scheffe's post hoc test.

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Fig. 6. LPA in RA-SF induces elevation of [Ca2+]i in hBMSCs. Serum-starved hBMSCs were loaded with 5 μM fluo-4-AM for 40 min at 37 °C in the absence or presence of 5 μM Ki16425 (Ki), and then treated with serum-free medium containing 0.5 μM 1-oleoyl-LPA, 5% RA-SF (case 1), or 0.1 μM S1P in the absence or presence of 5 μM Ki16425. Ca2+-dependent fluorescence was measured every second for indicated time periods, and fluorescence intensities of more than 20 different cells from time-lapse images were quantified over time. Results are expressed as percentage of the control (0 s) and expressed as mean ± SD.

LPA1 (Fig. 4E). These results are consistent with the suggestion that LPA plays a key role in RA-SF-stimulated migration of hBMSCs through activation of LPA1.

increase of [Ca2+]i was not affected by Ki16425 treatment (Fig. 6). These results indicate that LPA plays a key role in the elevation of [Ca2+]i elicited by RA-SF.

3.5. Increased concentrations of LPA and ATX in the SF derived from RA patients

3.7. Calcium/calmodulin-dependent protein kinase is involved in the migration of hBMSCs induced by SF from RA patients

Because ATX has been reported to play a key role in the production of LPA [17], we next determined the levels of ATX in SF from patients. As shown in Fig. 5A, ATX was detected in SF from patients with RA or OA using Western blotting, while it was hardly detected in SF from normal donors. The ATX levels in RA-SF were significantly higher than those in SF from normal donors (Fig. 5B), suggesting that ATX is markedly enriched in RA-SF. To further confirm the involvement of LPA in the RA-SF-induced migration, we next determined the concentrations of LPA in the SF derived from patients with RA or OA and normal donors. The concentrations of the LPA levels were estimated to be 0.3 ± 0.05 μM (normal SF), 0.98 ± 0.12 μM (RA-SF), and 0.52 ± 0.07 μM (OA-SF), respectively (Fig. 5C) and the LPA levels in RA-SF were significantly increased compared with those of healthy controls. Moreover, the LPA levels in OA-SF were slightly higher than those of healthy controls, albeit not statistically significant.

In order to explore the role of Ca2+ in the LPA-induced migration, we examined whether calmodulin is involved in the migration of hBMSCs induced by RA-SF and LPA because calmodulin is a key mediator of Ca2+ signaling. Pretreatment of hBMSCs with the calmodulin antagonist W7 markedly abrogated LPA-induced cell migration (Fig. 7). Furthermore, W7 also inhibited the migration of hBMSCs stimulated by RA-SF, suggesting that Ca2+/calmodulin complex plays a pivotal role in the migration of hBMSCs induced by LPA that is a key component of RA-SF. To explore the molecular mechanisms by which calmodulin mediates the LPA-induced migration, we next examined the involvement

3.6. SF derived from RA patients increases intracellular calcium concentration through LPA1-dependent mechanism To ascertain whether LPA is an active component of RA-SF, we examined the effects of RA-SF and LPA on the intracellular concentration of calcium ([Ca2+]i) of hBMSCs. Spatially averaged photometric measurements of [Ca2+]i were performed with the fluorescent Ca2+ indicator fluo-4-AM as described in Materials and methods. Both LPA and RA-SF rapidly increased [Ca2+]i in hBMSCs and pretreatment of hBMSCs with Ki16425 completely inhibited the elevation of [Ca2+]i induced by LPA and RA-SF (Fig. 6). To ascertain the specificity of Ki16425 on the LPA-induced elevation of [Ca2+]i, we examined the effect of Ki16425 on the S1P-induced increase of [Ca2+]i. S1P treatment elicited acute increase of [Ca2+]i, whereas S1P-induced

Fig. 7. The role of calmodulin-dependent protein kinase II in the RA-SF-induced migration of hBMSCs. hBMSCs were pretreated with vehicle, 5 μM W7, 2.5 μM KN92, or 2.5 μM KN93 for 15 min, and then exposed to serum-free medium containing 0.5 μM 1-oleoyl-LPA or 5% RA-SF (case 1) in the absence or presence of the pharmacological inhibitors for 12 h. The number of migrating cells was counted and data represent mean ± SD (n = 4). ⁎, p b 0.01 by two-way ANOVA and Scheffe's post hoc test.

H.Y. Song et al. / Biochimica et Biophysica Acta 1801 (2010) 23–30

of calmodulin-dependent protein kinase II (CaMK II), a downstream mediator of calmodulin, in the LPA-induced migration of hBMSCs. As shown in Fig. 7, pretreatment of hBMSCs with the CaMK II-specific inhibitor KN93 completely abrogated the cell migration induced by RA-SF or LPA. However, KN92, an inactive analog of KN93, had no significant impact on the migration of hBMSCs. These results suggest that activation of CaM/CaMK II signaling pathway plays a key role in the migration of hBMSCs induced by LPA in RA-SF. 4. Discussion In the present study, we demonstrated for the first time that LPA is responsible for the RA-SF-stimulated migration of hBMSCs through LPA1-dependent mechanism. The migration of hBMSCs induced by RA-SF and LPA was completely abrogated by the LPA1/3 inhibitor Ki16425. Furthermore, siRNA- or lentiviral shRNA-mediated depletion of LPA1 specifically abolished the RA-SF-induced migration of hBMSCs. It has been reported that LPA1 is implicated in LPA-induced cell migration in a variety of cell types [32–35]. Together with the previous report that LPA stimulated migration of human adipose tissuederived MSCs through LPA1-dependent mechanism [27], these results support the notion that LPA1 plays a key role in the LPA-induced migration of hBMSCs. An increasing body of evidence suggests that migration of MSCs is regulated by various growth factors, chemokines, and lipid factors, including PDGF-BB, insulin-like growth factor, stromal-derived factor-1, monocyte chemoattractant protein-1, and S1P [30,36–39]. RA-SF has been reported to contain various growth factors and chemokines [40,41], implying their potential role in RA-SF-induced migration of hBMSCs. However, the involvement of protein factors in RA-SF-induced migration can be ruled out, because heat denaturation did not abrogate the RA-SF-stimulated migration of hBMSCs (Fig. 2A). Moreover, we showed that S1P- or PDGF-BB-induced migration was not affected by pharmacological inhibition or depletion of LPA1, indicating that S1P and PDGF-BB are not involved in the RA-SF-induced migration. Taken together, these results led us to suggest that LPA plays a pivotal role in the RA-SF-induced migration of human MSCs. We demonstrated that both RA-SF and LPA increased [Ca2+]i through a Ki16425-sensitive mechanism, whereas Ki16425 had no significant impact on S1P-induced elevation of [Ca2+]i, suggesting the involvement of LPA in the elevation of [Ca2+]i induced by RA-SF. Furthermore, pharmacological inhibition of CaM/CaMK II-dependent pathway inhibited migration of hBMSCs induced by either RA-SF or LPA. Activation of CaM/CaMK II pathway has been reported to stimulate migration of vascular smooth muscle cells [42–44]. Selective pharmacological inhibitors of CaMK II (KN62 or KN93) abrogated chemotactic migration of vascular smooth muscle cells. These results support the present study that CaM/CaMK II-dependent pathway contributes to the LPA-induced migration of hBMSCs, although the molecular mechanism by which CaMK II mediates the chemotactic migration of hBMSCs should be clarified further. In the present study, we demonstrate that RA-SF contains a higher concentration of LPA than SF from normal donors. The concentrations of LPA in RA-SF and normal SF were estimated to be 0.98 ± 0.12 μM and 0.3 ± 0.05 μM, respectively. Furthermore, we showed that the amounts of ATX in RA-SF were significantly higher than those in normal SF, suggesting that ATX may be responsible for the high level of LPA in RASF. Consistent with the results, existence of LPA and ATX in RA-SF has been reported [16]. The concentration of LPA in RA-SF was estimated to be 3.7 ± 2.2 μM by using a bioassay based on the ability of LPA to inhibit cAMP accumulation in LPA1-expressing RH7777 cells and the concentration of LPA in RA-SF was further increased after incubation of SF in the culture medium at 37 °C, suggesting ATX-mediated production of LPA [16]. Moreover, recent evidence suggests that RA-SF contains both ATX [16,20] and the LPA precursor lysophosphatidylcholine [16,45]. These results raised a possibility that LPA in RA-SF could

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be produced by ATX during preparation of SF. However, in the present study, all preparation steps were carried out at 4 °C to prevent ATXmediated generation of LPA during preparation of SF and we demonstrated the existence of LPA in freshly prepared RA-SF (Fig. 5). These results indicate that RA-SF naturally contains a high level of LPA, although it is still possible to speculate that LPA may be further produced by ATX during incubation of RA-SF-containing medium at 37 °C for cell migration. Similar to the present study that SF induced migration of hBMSCs, SF reportedly stimulated migration of human MSCs derived from subchondrial spongious bone marrow [46]. Although it has not been clarified whether BMSCs can be infiltrated into synovium of RA patients through LPA-dependent mechanism, multipotent population of MSCs has been reported to exist in not only SF of patients with RA or OA but also normal SF [13,47,48]. The phenotypes and functional properties of MSCs derived from SF were similar to those of BMSCs [47,49,50]. In addition, BMSCs have been reported to comprise 1.2% of normal fibroblast-like synoviocyte and the population of BMSCs was increased up to 33.7% of fibroblast-like synoviocytes from RA animal models, suggesting that influx of BMSCs into synovium during RA development [51]. Moreover, it has been proposed that synovial MSCs were derived from the adjacent synovium or the cartilage superficial layer [6,52]. SF stimulated expansion of MSCs from synovium of OA patients in tissue culture system by stimulating cell migration [53]. Furthermore, LPA1/3 receptor antagonists and autotaxin inhibitors reduced the SF-induced migration of synoviocytes [20]. These results led us to suggest that LPA in SF may promote homing of BMSCs into synovium, albeit it should be determined whether BMSCs can infiltrate into synovium of RA patients in vivo. Acknowledgements This work was supported by the MRC program of MOST/KOSEF (R13-2005-009). We highly acknowledge Dr. Junken Aoki (Tohoku University, Japan) for providing anti-ATX antibody. References [1] M. Feldmann, F.M. Brennan, R.N. Maini, Rheumatoid arthritis, Cell 85 (1996) 307–310. [2] C.D. Buckley, Michael Mason prize essay 2003. Why do leucocytes accumulate within chronically inflamed joints? Rheumatology (Oxford) 42 (2003) 1433–1444. [3] R. Scrivo, M. Di Franco, A. Spadaro, G. Valesini, The immunology of rheumatoid arthritis, Ann. N. Y. Acad. Sci. 1108 (2007) 312–322. [4] P. Ghosh, D. Guidolin, Potential mechanism of action of intra-articular hyaluronan therapy in osteoarthritis: are the effects molecular weight dependent? Semin. Arthritis Rheum. 32 (2002) 10–37. [5] Z. Szekanecz, R.M. Strieter, S.L. Kunkel, A.E. Koch, Chemokines in rheumatoid arthritis, Springer Semin. Immunopathol. 20 (1998) 115–132. [6] C. De Bari, F. Dell'Accio, P. Tylzanowski, F.P. Luyten, Multipotent mesenchymal stem cells from adult human synovial membrane, Arthritis Rheum 44 (2001) 1928–1942. [7] D.J. Prockop, Marrow stromal cells as stem cells for nonhematopoietic tissues, Science 276 (1997) 71–74. [8] M.F. Pittenger, A.M. Mackay, S.C. Beck, R.K. Jaiswal, R. Douglas, J.D. Mosca, M.A. Moorman, D.W. Simonetti, S. Craig, D.R. Marshak, Multilineage potential of adult human mesenchymal stem cells, Science 284 (1999) 143–147. [9] P.A. Zuk, M. Zhu, P. Ashjian, D.A. De Ugarte, J.I. Huang, H. Mizuno, Z.C. Alfonso, J.K. Fraser, P. Benhaim, M.H. Hedrick, Human adipose tissue is a source of multipotent stem cells, Mol. Biol. Cell 13 (2002) 4279–4295. [10] S. Alsalameh, R. Amin, T. Gemba, M. Lotz, Identification of mesenchymal progenitor cells in normal and osteoarthritic human articular cartilage, Arthritis Rheum. 50 (2004) 1522–1532. [11] F.P. Barry, J.M. Murphy, Mesenchymal stem cells: clinical applications and biological characterization, Int. J. Biochem. Cell Biol. 36 (2004) 568–584. [12] B. Short, N. Brouard, T. Occhiodoro-Scott, A. Ramakrishnan, P.J. Simmons, Mesenchymal stem cells, Arch. Med. Res. 34 (2003) 565–571. [13] E.A. Jones, A. English, K. Henshaw, S.E. Kinsey, A.F. Markham, P. Emery, D. McGonagle, Enumeration and phenotypic characterization of synovial fluid multipotential mesenchymal progenitor cells in inflammatory and degenerative arthritis, Arthritis Rheum 50 (2004) 817–827. [14] L. Marinova-Mutafchieva, R.O. Williams, K. Funa, R.N. Maini, N.J. Zvaifler, Inflammation is preceded by tumor necrosis factor-dependent infiltration of mesenchymal cells in experimental arthritis, Arthritis Rheum 46 (2002) 507–513. [15] A. Tokumura, Physiological and pathophysiological roles of lysophosphatidic acids

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