Resistin promotes angiogenesis in endothelial progenitor cells through inhibition of microRNA206: potential implications for rheumatoid arthritis
Chen-Ming Su1, Chin-Jung Hsu2,3, Chun-Hao Tsai2,4, Chun-Yin Huang4,5, Shih-Wei Wang6, and Chih-Hsin Tang1,7,8*
1Graduate Institute of Basic Medical Science, China Medical University, Taichung, Taiwan
2Department of Orthopedic Surgery, China Medical University Hospital, Taichung, Taiwan.
3School of Chinese Medicine, College of Chinese Medicine, China Medical University, Taichung, Taiwan
4Graduate Institute of Clinical Medical Science, China Medical University, Taichung, Taiwan
5Department of Orthopedic Surgery, China Medical University Beigang Hospital, Yun-Lin County, Taiwan
6Department of Medicine, Mackay Medical College, New Taipei City, Taiwan
7Department of Pharmacology, School of Medicine, China Medical University, Taichung, Taiwan
8Department of Biotechnology, College of Health Science, Asia University, Taichung, Taiwan
*Corresponding author Chih-Hsin, Tang PhD
Graduate Institute of Basic Medical Science, China Medical University No. 91, Hsueh-Shih Road, Taichung, Taiwan
Tel: (886) 4-22052121 Ext. 7726. Fax: (886) 4-22333641.
E-mail: [email protected]
Running title: Resistin induces EPCs migration and tube formation.
Key words: Resistin, Endothelial progenitor cell, Rheumatoid arthritis, Collagen- induced arthritis, and microRNA
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Abstract
Endothelial progenitor cells (EPCs) promote angiogenesis and are therefore key contributors to a wide variety of angiogenesis-related autoimmune diseases such as rheumatoid arthritis (RA). However, the signaling mechanisms through which these progenitor cells influence RA pathogenesis remain unknown. The aim of this study was to examine whether resistin plays a role in the pathogenesis of and angiogenesis associated with RA by circulating EPCs. We found that levels of resistin in synovial fluid and tissue from patients with RA and from mice with collagen-induced arthritis were overexpressed and promoted the homing of EPCs into the synovium, thereby inducing angiogenesis. EPCs isolated from healthy donors were used to investigate the signal transduction pathway underlying EPC migration and tube formation after treatment with resistin. We found that resistin directly induced a significant increase in expression of vascular endothelial growth factor (VEGF) in EPCs. We also found that the expression of microRNA-206 (miR-206) was negatively correlated with the expression of resistin during EPC-mediated angiogenesis. Notably, the increased expression of VEGF was associated with decreased binding of miR-206 to the VEGF- A 3ʹ untranslated region through protein kinase C delta–dependent AMP-activated protein kinase signaling pathway. Moreover, blockade of resistin reduced EPC homing into synovial fluid and angiogenesis in vivo. Taken together, our study is the first to demonstrate that resistin promotes EPCs homing into the synovium during RA angiogenesis via a signal transduction pathway that involves VEGF expression in primary EPCs. These findings provide support for resistin as a therapeutic target for the patients with RA.
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Introduction
Rheumatoid arthritis (RA), one of the most common autoimmune diseases, is characterized by the infiltration of proinflammatory cytokines into synovial fluid leading to pannus formation and joint destruction [1]. Proangiogenic cells such as endothelial stem cells are crucial to the development and progression of RA as they promote the creation of angiogenic ventilation shafts which consequently cause synovitis by forming new capillaries in the joints [2, 3].
Bone marrow-derived endothelial stem cellsare characterized by their ability to self-renew and develop into intermediate stem cell–endothelial progenitor cells (EPCs) [4]. EPCs possess cellular subpopulations with different functional capacities, including proliferation, migration, and recruitment in response to angiogenesis.
Circulating EPCs, a cell population characterized by the CD133+/CD34+/VEGFR2+ phenotype, are mobilized from the bone marrow into the bloodstream and are involved in neovascularization by becoming incorporated within neovessels [5-7].
Although EPCs are critical components of the earliest phases of tumor neovascularization [8], it is still unclear whether EPCs are involved in the initiation of RA by causing deterioration in the joints.
Resistin, a small cysteine-rich adipose-derived peptide hormone, is specifically expressed and secreted by adipocytes and is linked to both inflammation [9, 10] and insulin resistance [11]. Previous research has shown that increased serum resistin levels in patients with insulin resistance correspond to increased levels of protein kinase C delta (PKC-) signaling [12, 13]. A number of studies have demonstrated that resistin levels in serum and synovial fluid are elevated in patients with RA [14- 16] and that resistin levels might correlate with markers of inflammation. Thus, elevated resistin may be a predictor of RA.
Increased resistin expression has been shown to be correlated with increased expression of vascular endothelial growth factor (VEGF), one of the most important angiogenic switch molecules [17]. Accumulating evidence has revealed that VEGF is regulated by by microRNAs (miRNAs) at multiple levels during angiogenesis [18, 19]. These miRNAs are small non-coding RNA molecules that map to intronic locations within protein-coding genes and function in RNA silencing and post- translational regulation of gene transcription. Some miRNAs have been shown to be possible effectors of autoimmune diseases and to be involved in both physiological and pathological angiogenesis, as they regulate several genes that participate in both functions [20, 21]. However, miRNAs have hundreds of putative targets, and it is challenging to determine the physiological functions of individual miRNA-target interactions.
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A previous study showed that chemokine-induced angiogenesis is an early and critical event in RA pathogenesis [2]. However, the role resistin plays in the development of and angiogenesis associated with autoimmune diseases such as RA remains unclear. In the present study, we investigated whether highly expressed resistin in patients with RA upregulates the function of human circulating EPCs in RA-associated degeneration and explored the signaling mechanisms involved in this process. We conclude by discussing the possible significance of our results.
Materials and Methods
Materials
Anti-mouse and anti-rabbit horseradish peroxidase-conjugated IgG, and rabbit polyclonal antibodies specific for -actin, PKC-, phosphorylated PKC-, AMP- activated protein kinase 1 (AMPK1), phosphorylated AMPK1, CD31, and CD133 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). A mouse monoclonal antibody and enzyme immunoassay kit specific for resistin were purchased from R&D Systems (Minneapolis, MN, USA). Recombinant human resistin was purchased from PeproTech (Rocky Hill, NJ, USA). Anti-resistin neutralized antibody was purchased from Fisher Scientific (Thermo Fisher Scientific Inc., NY, USA). ON-TARGETplus small interfering RNA (siRNA) targeting VEGF, PKC-, and AMPKα1, and ON-TARGETplus non-targeting siRNA (control) were purchased from Dharmacon Research (Lafayette, CO, USA). Synthesized sequences of a human miRNA-206 (miR-206) mimic, a miR-206 inhibitor, and a negative control miRNA were purchased from GeneDireX Inc. (Las Vegas, NV, USA).
Customized miRNA array primers were purchased from System Biology Ireland (Galway, Ireland). Rottlerin, Compound C, and adenosine-9-β-D-arabino-furanoside (Ara A) were purchased from Calbiochem (San Diego, CA, USA). All other chemicals were purchased from Sigma-Aldrich (St. Louis, MO, USA).
Human synovial fluids and tissues
We obtained approval for this study from the local ethics committee and all subjects gave written informed consent to participate. Abnormal synovial fluids and synovial tissues were obtained from 10 patients (aged 40–60 years) during total knee arthroplasty for RA, and normal synovial fluids and tissues were obtained from 4 patients during arthroscopy for trauma/joint derangement. The protocol for this study was approved by the Institutional Review Board of the China Medical University Hospital (DMR-103-059), and informed consent was obtained from each donor.
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Primary synovial fibroblasts from RA patients and normal patients were isolated, cultured, and characterized as previously described [22]. Briefly, human primary synovial fibroblasts were isolated in a solution of type II collagenase for 16–18 h and then filtered through 70-m nylon filters. The cells from passages 3 to 9 were used for the experiments.
Isolation of human circulating EPCs
EPCs were derived from healthy donors who gave informed consent before enrollment, and ethical approval was granted by the Institutional Review Board of Mackay Medical College (New Taipei City, Taiwan; reference number: P1000002).
EPCs were isolated and purified as previously described [23, 24]. Briefly, mononuclear cells were isolated from peripheral blood (80 mL) using the Ficoll- Paque™ plus (Amersham Biosciences, Uppsala, Sweden) centrifugation method.
Then, EPCs were separated from the isolated mononuclear cells using the MicroBead Kit and the MACS™ Cell Separation System (all from Miltenyi Biotec, Bergisch Gladbach, Germany). EPCs were maintained and characterized as follows: briefly, EPCs were seeded on gelatin-coated dishes containing MV2 medium, SupplementMix (PromoCell, Heidelberg, Germany) and 20% non-heat-inactivated defined fetal bovine serum (FBS; HyClone, Logan, UT, USA) and incubated for 3 days; the medium was changed every 3 days. For characterization, labeled EPCs were examined and characterized with CD34+/CD133+/VEGFR2+ antibodies by using a FACSCaliburTM flowcytometer and CellQuestTM software (both from BD Biosciences, San Jose, CA, USA) [25, 26].
Collagen-induced arthritis mouse model
Male C57BL/6J mice (aged 8–10 weeks) were purchased from the National Laboratory Animal Centre in Taipei. Mice were maintained under conditions consistent with the guidelines established by the Animal Care Committee of the China Medical University. A collagen-induced arthritis (CIA) mouse model was established according to published protocols [27]. Briefly, 0.1 mL of an emulsion containing 100
g of bovine type II collagen (CII; Sigma) dissolved at a concentration of 2 mg/mL in 0.1 M acetic acid and complete Freund’s adjuvant was injected intradermally into the base of the tail. Two weeks after the primary immunization, a booster injection of 100
g CII dissolved and emulsified 1:1 with incomplete Freund’s adjuvant was administered into the hind leg. The incidence of arthritis in the CIA mice was very high within 6 weeks after the first immunization, and 95% of the mice developed severe arthritis.
The clinical severity of arthritis in each knee was measured in a blinded manner 146
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with a plethysmometer (Marsap, Mumbai, India) once weekly for 4 weeks. Upon sacrifice on day 42, the phalanges and knee joints were removed immediately and fixed in 4% paraformaldehyde for micro computed tomography (micro-CT) analysis.
Micro-CT scanning of the knees was performed using an in vivo micro-CT scanner (Skyscan 1176; Bruker Corp., Kontich, Belgium) at 9 m resolution and with a rotation step of 0.30 degree per image. Scanning was done at 50 keV of X-ray voltage, 500 A of current, and 885 ms of exposure time/image with a 0.5 mm aluminum filter. Reconstruction of sections was carried out with GPU-based scanner software (NRecon, Bruker Corp.). In addition, the grayscale was based on the Hounsfield unit, and the validated calcium standards were scanned as the density reference. The three-dimensional microstructural volumes from the micro-CT scans were analyzed using Skyscan software (CTAn, Bruker Corp.). To evaluate bone focal erosion in the knee joint, quantification of volumetric bone mineral density (BMD) was performed in bone areas defined by manually drawn regions of interest (8 mm/913 slices from the proximal calcaneus to the metatarsals).
EPC tube formation
Matrigel (BD Biosciences, Bedford, MA, USA) was dissolved at 4°C, added to 48- well plates at 100 µL/well, and incubated at 37°C for 30 min. EPCs (2 × 104/100 µL) were resuspended in MV2 serum-free medium with the indicated concentration of resistin, and then added to the wells. VEGF (50 ng/mL) and PBS were used as the positive and negative control, respectively. After 6–8 h of incubation at 37°C, EPC tube formation was assessed by microscopy, and each well was photographed at 200×
magnification under a light microscope. The number of tube branches and the total tube length were calculated and quantified using MacBiophotonics Image J software.
Immunohistochemistry
Paraffin-embedded plug sections were prepared, mounted on silane-coated slides, deparaffinized in xylene, rehydrated in a graded alcohol series, and washed in deionized water. After antigen retrieval (boiling in a microwave for 30 min in 10 mM sodium citrate, pH 6.0), the intrinsic peroxidase activity was blocked by incubation with 3% hydrogen peroxide. Nonspecific antibody-binding sites were blocked using 3% bovine serum albumin (BSA) in PBS. Plug sections were then incubated with appropriately diluted primary rabbit polyclonal anti-mouse resistin, VEGF, CD31, or CD133 antibodies (Abcam, MA, USA) at 4°C overnight. After 3 washes in PBS, the secondary antibody (biotin-labeled goat anti-rabbit IgG) was applied for 1 h at room temperature. Staining was detected with 3,3ʹ-diaminobenzidine tetrahydrochloride (DAB) and the sections were then counterstained with hematoxylin and eosin and 184
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observed under a light microscope. Some specimens were also stained with Safranin O-fast Green for bone erosion.
Quantification of mRNA and miRNA by real-time quantitative polymerase chain reaction amplification
Total RNA was extracted from EPCs with a TRIzol kit (MDBio, Taipei, Taiwan) and miRNAs were quantified using the Mir-XTM miRNA First Strand Synthesis Kit (Clontech Laboratories, Inc., CA, USA), as per the manufacturers’ protocols. The RNA quality and yield of each total RNA sample was determined via A260 measurements using a NanovueTM Spectrophotometer (GE Healthcare, WI, USA).
Complementary DNA was derived from 1 μg of total RNA using an M-MLV RT kit (Invitrogen, CA, USA), according to the manufacturer’s recommendations. Real-time quantitative polymerase chain reaction (qPCR) analysis was carried out with the KAPA SYBR® FAST qPCR Kit (Applied Biosystems, CA, USA). The cycling conditions were as follows: polymerase activation for 10 min at 95°C followed by 40 cycles at 95°C for 15 s and 60°C for 60 s. Relative normalization of gene expression was performed using endogenous GAPDH as the internal control for mRNA or snRNA U6 as the internal control for miRNA. Relative expression levels were calculated using the comparative threshold cycle (CT), which is the number of the cycle at which the transcript was detected [28]. All target gene primers were purchased from Applied Biosystems and are listed in Supplementary Table S1.
Enzyme-linked immunosorbent assay
Human primary EPCs were cultured in 24-well plates, treated with resistin, and then incubated in a humidified incubator at 37°C for 24 h. To examine the downstream signaling pathways involved in resistin treatment, cells were pretreated with various inhibitors for 30 min before resistin (3 ng/mL) was added. After incubation, the supernatant was collected as conditioned medium (CM) and stored at −80°C until the assay was performed. VEGF in the CM was assayed using a VEGF-A enzyme-linked immunosorbent assay (ELISA) kit (R&D Systems) according to the procedure described by the manufacturer.
Transwell migration assay
All EPC migration assays were performed using Transwell inserts (8 μm pore size;
Costar, NY, USA) in 24-well plates. EPCs were pretreated with the indicated concentrations of inhibitors (Rottlerin, Compound C, or Ara A) for 30 min or transfected with various siRNAs for 16–18 h; then, the cells were treated with resistin for another 24 h. EPCs (5 × 103 in 200 μL of medium with 10% FBS) were then 222
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seeded in the upper chamber, and 300 μL of the same medium containing the appropriate concentration of resistin was placed in the lower chamber. Cells on the upper side of the Transwell membrane were examined and counted under a microscope. Each experiment was performed in triplicate and was repeated at least 3 times.
Western blot analysis
Proteins were resolved using sodium dodecyl sulfate–polyacrylamide gel electrophoresis and transferred to Immobilon polyvinyldifluoride membranes (Millipore, MA, USA). The membranes were blocked with 4% BSA for 1 h at room temperature and then probed with rabbit antibodies against human phosphorylated PKC-, PKC-, phosphorylated AMPK1, or AMPK1, (1:1,000) for 1 h at room temperature. After 3 washes, the blots were incubated with a donkey anti-rabbit peroxidase-conjugated secondary antibody (1:1000) for 1 h at room temperature. The blots were then developed via enhanced chemiluminescence and visualized using a Fujifilm LAS-3000 chemiluminescence detection system (Fujifilm, Tokyo, Japan).
Plasmid construction and luciferase assays
The 3ʹ-untranslated region (3ʹUTR) of human VEGF-A contains a miR-206 binding site. A DNA fragment containing VEGF 3ʹUTR (wt-VEGF 3ʹUTR) and a fragment containing a version of the VEGF 3ʹUTR in which the miR-206 binding site had been mutated (mut-VEGF 3ʹUTR) were purchased from Invitrogen. Each fragment was subcloned into the luciferase reporter vector pGL2-Control (Promega, Madison, WI, USA), upstream of the vector’s promoter. All constructs were confirmed by sequencing using the Applied Biosystems 3730xl DNA Analyzer (Thermo Fisher Scientific Inc., NY, USA). EPCs, grown to 80% confluence, were transiently cotransfected with a luciferase reporter construct (wt-VEGF 3ʹUTR or mut-VEGF 3ʹUTR) and the miR-206 mimic, the miR-206 inhibitor, or the control miRNA using Lipofectamine 2000 according to the manufacturer's instructions (Invitrogen). After treatment with resistin for 24 h, luminescence from treated EPCs lysates was measured using a microplate luminometer. Luciferase activity was normalized to that of the cotransfected -galactosidase expression vector.
Chick chorioallantoic membrane (CAM) assay
In vivo angiogenetic activity was determined using a chick chorioallantoic membrane (CAM) assay as previously described [29]. Briefly, 5-day-old fertilized chick embryos (6 eggs/group) were incubated at 37°C in an 80% humidified atmosphere. On developmental day 8, PBS (vehicle) or resistin was resuspended in Matrigel and 260
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placed onto the CAMs for another 3 days. The CAMs were then examined by microscopy and photographed. Angiogenesis was quantified by counting the number of blood vessel branches. All animal work was performed in accordance with a protocol approved by the China Medical University (Taichung, Taiwan) Institutional Animal Care and Use Committee.
In vivo Matrigel plug assay
The Matrigel plug angiogenesis assay was performed as previously described [30].
Thirty 4-week-old male nude mice were randomized into 3 groups and subcutaneously injected with 0.15 mL Matrigel containing PBS (vehicle) or resistin (3 ng/mL). On day 7, the Matrigel plugs were harvested; some were fixed with 3%
paraformaldehyde for at least 2 days, and then embedded in paraffin and subsequently processed for immunostaining of VEGF, CD31, and CD133; others were evaluated by Drabkin’s method (Drabkin’s Reagent Kit; Sigma) to quantify the hemoglobin content.
Statistical analysis
All quantified results were calculated using GraphPad Prism 5.0 software and are presented as the mean ± SD of at least 3 experiments. Statistical comparison of 2 groups was performed using the Student’s t-test. Statistical comparisons of more than 2 groups were performed using two-factor analysis of variance with the Bonferroni post hoc test or the Mann-Whitney U test, as appropriate. In all cases, P < 0.05 was considered statistically significant.
Results
Resistin is highly expressed in synovial fluid and tissues of RA patients
Resistin has been shown to be associated with the overexpression of proinflammatory cytokines such as IL-6, TNF-, and IL-1 in the progression of RA [31, 32]. Thus, we first examined whether resistin is highly expressed in patients with RA. We found that resistin protein and resistin mRNA levels were significantly higher in synovial fluids and tissues from patients with RA than in healthy patients (Figure 1A, B). Resistin protein levels were also significantly higher in primary synovial fibroblasts from RA patients than in normal synovial fibroblasts (Figure 1C). A previous study demonstrated preferential homing of EPCs to the inflamed synovium [33]. Therefore, we conducted a migration assay combined with anti-resistin antibody to investigate the homing abilities of EPC in synovial fluids from patients with and without RA 298
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following resistin treatment (Figure 1D) and found that the observed effects in RA synovial fluid were attributable to highly expressed resistin. In addition, we examined whether resistin had similar functions in the CIA mice, one of the most commonly used animal models for evaluating the efficacy of RA therapies [34]. We found that CIA mice had significantly greater bone erosion and worse BMD in the knee joints than mice in the control group on day 42 (Figure 1E and F). Safranin O staining also showed significantly greater bone erosion in knee joints of CIA mice (Figure 1G).
Double immunofluorescent staining of the synovium with EPC markers CD34 and CD133 (Figure 1H) and immunostaining of tissue sections for resistin, EPC markers (CD133 and VEGFR2), and vessels markers (CD31 and VEGF) revealed increased expression in CIA mice (Figure 1I). These observations suggest that EPC infiltration and VEGF production are involved in arthritic development in CIA mice. Taken together, these data demonstrate that resistin plays a significant role in angiogenesis during the arthritic process.
Resistin promotes VEGF-dependent EPC migration and tube formation
Since VEGF is one of the most potential angiogenic mediators and induces the formation of new capillaries from pre-existing vessels, angiogenesis may occur and progress in RA pathogenesis [2, 35]. Although resistin has been shown to directly enhance angiogenesis in numerous diseases [17, 36], the mechanism by which it promotes angiogenesis and EPC homing is not clear. Therefore, we used circulating EPCs to investigate whether resistin plays a role in the expression of VEGF.
Treatment of EPCs with resistin (0.03–3.0 ng/mL) for 24 h induced VEGF mRNA expression in a concentration-dependent manner (Figure 2A). Resistin stimulation also led to a concentration-dependent increase in VEGF protein expression (Figure 2B), and induced a concentration-dependent increase in EPC migration and tube formation (Figure 2C–E). These results indicate that activation of EPC migration and tube formation is critical for resistin-mediated VEGF expression. In contrast, transfection with VEGF siRNA notably abolished resistin-increased EPC migration and tube formation (Figure 2C–E). Taken together, these data suggest that resistin facilitates VEGF-dependent EPC migration and tube formation.
The effect of resistin on EPCs is mediated by the PKC-/AMPK signaling pathway
Previous research has shown that increased serum resistin levels are associated with increased PKC- signaling, which is involved in angiogenesis [13, 37, 38]. To verify the involvement of PKC- in the response of EPCs to resistin, PKC-
phosphorylation was examined in EPCs after resistin stimulation (Figure 3A). We 335
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then pretreated EPCs with the PKC- inhibitor Rottlerin or transfected cells with PKC- siRNA or control siRNA. We found that both Rottlerin and the PKC- siRNA abolished resistin-induced VEGF production (Figure 3B, C) as well as EPC migration and tube formation (Figure 3D–F). A previous study showed that PKC-δ is upstream of AMPK and is required for the activation of AMPK in phenylephrine preconditioning [39]. In the present study, we found that resistin stimulation enhanced AMPK phosphorylation in a time-dependent manner (Figure 4A). We also examined whether AMPK is a downstream effector of PKC-δ. Pretreatment with a PKC-δ inhibitor or PKC-δ siRNA abolished resistin-promoted AMPK phosphorylation (Figure 4B). As shown in Figures 4C and D, levels of VEGF mRNA and VEGF protein were reduced in cells pretreated with the AMPK inhibitors Ara A and Compound C and in cells that underwent transfection with AMPK1 siRNA. To ascertain whether resistin activated EPC migration and tube formation via the AMPK signaling pathway, we used AMPK inhibitors or AMPK1 siRNA. The results show that AMPK activation is required for the EPC response to resistin (Figure 4E–G).
Taken together, these results indicate that resistin increases VEGF expression and EPC migration and tube formation via activation of the PKC-δ-dependent AMPK signaling pathway.
Resistin induces EPC migration and tube formation by downregulating miR-206 via the PKC-δ/AMPK signaling pathway
Recently, miRNAs have been shown to be powerful regulators in autoimmune diseases; however, the targets of most miRNAs remain unknown and the roles of miRNAs in biological processes are not clearly understood [40]. We used a customized miRNome microRNA Profilers Kit, which contains 384 human miRNAs, and found that approximately one-fifth of the miRNAs examined were influenced by resistin. To specifically study miRNAs affected by resistin, we selected miRNAs for which expression changed by at least two-fold in the presence of resistin. We found that eight of the miRNAs were upregulated and three were significantly downregulated when EPCs were treated with resistin (see Supplementary Figure S1).
Next, we integrated the results of the miRNAs array with 3 online computational algorithms (TargetScan, PicTar, and miRanda) to identify conserved miRNAs that targeted VEGF-A and found 1 candidate miRNA: miR-206.
We then evaluated the miR-206 expression profile in EPCs. Treatment of human circulating EPCs with resistin resulted in a significant, concentration-dependent decrease in miR-206 levels (Figure 5A). To determine whether miR-206 directly regulates VEGF, we transiently transfected EPCs with a miR-206 mimic, a miR-206 inhibitor, or a negative control miRNA and found that transfection with the miR-206 373
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mimic inhibited the induction of VEGF mRNA and VEGF protein in EPCs that had been exposed to resistin. In contrast, transfection with the miR-206 inhibitor increased the expression of VEGF mRNA and VEGF protein in response to resistin (Figure 5B–
D). In addition, resistin-induced EPC migration and tube formation were attenuated by the miR-206 mimic but were largely enhanced by the miR-206 inhibitor (Figure 5E–G). To further investigate the specificity for which miR-206 targets VEGF 3ʹUTR, we transfected two firefly luciferase reporter plasmids, wt-VEGF 3ʹUTR and mut- VEGF 3ʹUTR, into EPCs and then analyzed cell extracts for luciferase activity. The results showed that resistin notably increased luciferase activity in cells transfected with the wt-VEGF 3ʹUTR luciferase reporter plasmid but not in those transfected with the mut-VEGF 3ʹUTR luciferase reporter plasmid (Figure 5H), suggesting that resistin promotes the expression of the VEGF 3ʹUTR in the absence of the miR-206 binding site. Furthermore, pretreatment with Rottlerin, Compound C, or Ara A or transfection with siRNAs against PKC-δ or AMPK reversed the resistin-mediated downregulation of miR-206 expression in EPCs (Figure 5I), indicating that the PKC-δ/AMPK- dependent regulation of miR-206 expression is involved in the resistin-induced signaling pathway. These data therefore show that resistin downregulated the expression of miR-206 and then promoted VEGF expression and EPC migration as well as tube formation via the PKC-δ/AMPK signaling pathway.
Resistin is associated with EPC angiogenesis in vivo
To determine the effects of resistin on VEGF expression and EPC tube formation in vivo, we investigated angiogenesis using the CAM assay. PBS (vehicle) or resistin (3 ng/mL) were resuspended in Matrigel and then placed onto the surface of the CAMs for clear observation. Interestingly, CAMs incubated with resistin had significantly more new capillaries than CAMs incubated with PBS (Figure 6A).
Quantification of the vessels at the surface of the CAMs is shown in Figure 6B. To further confirm that EPCs are recruited by resistin in vivo, we subcutaneously injected 0.15 mL Matrigel containing PBS or resistin (3 ng/mL) into the flanks of nude mice and found that the Matrigel plugs containing resistin resulted in a greater degree of microvessel formation than plugs containing PBS (Figure 6C). In addition, we quantified the hemoglobin content of the Matrigel plugs using Drabkin’s method (Figure 6D). When we evaluated paraffin-embedded plug sections by immunohistochemistry, we found that levels of VEGF, CD31, and CD133 were higher in resistin-treated Matrigel plugs than in Matrigel plugs containing PBS (Figure 6E). To further demonstrate the role of resistin in RA angiogenesis, we tested the effect of resistin on tube formation by exposing synovial fluid from RA patients and control subjects to blockade of resistin (Figure 6F, G) and found that resistin 411
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played a crucial role in RA angiogenesis in vitro. In addition, we investigated in synovial fluid from RA patients with or without exposure to blockade of resistin using the CAM assay and Matrigel plug assay, respectively (Figure 6H, J) and quantified vessels count and hemoglobin content, respectively (Figure 6I, K). We found that blockade of resistin attenuated angiogenesis in RA synovial fluid in vivo, indicating that highly expressed resistin plays an important role in RA angiogenesis. Taken together, the results suggest that resistin significantly promotes EPC migration and tube formation in vivo, implicating that resistin is a crucial modulator of the pathogenesis of RA.
Discussion
RA is characterized by peripheral symmetrical polysynovitis associated with the accumulation of body fat in patients with RA [41]. The abnormal accumulation of adipose tissue in rheumatoid joints might implicate certain factors in the regulation of peripheral polysynovitis in RA, as both bone marrow and white adipose tissues secrete a wide variety of proteins known as adipokines that participate in the regulation of processes involved in metabolism, immunity, and inflammation [42].
Previous studies have characterized resistin as a potent proinflammatory adipokine that appears to have a critical role in RA pathogenesis [9, 43]. In our study, we found that high levels of resistin in patients with RA drive the function of circulating EPCs during RA processes.
EPCs are associated with the induction and modulation of VEGF during RA pathogenesis [44-46], and angiogenesis has recently become a possible therapeutic target for patients with the disease [47, 48]. As widespread pattern, it is not yet clear however, whether circulating EPCs are associated with the regulation and mechanistic function of resistin in RA pathogenesis. Several studies have demonstrated that EPCs are connected with abnormal angiogenesis via the induction and modulation of VEGF during RA pathogenesis [44-46]. In our study, there are limitations of the present study that EPCs from RA patients are too rare and hard to acquire. However, we were only able to provide indirect evidence of the role EPCs play in the pathogenesis of RA because we used EPCs derived from mononuclear cells that had been isolated from peripheral blood of healthy patients rather than EPCs from RA patients. In addition, irrespective of whether resistin plays a key role in inflammatory or neovascular- signaling pathways, angiogenesis as a target of therapeutic intervention in RA has become a current possibility [47, 48]. On the other hand, although it is possible to observe most of the histopathological characteristics of RA in CIA mice [49], EPC infiltration through the synovial membrane in CIA mice has not been reported 449
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previously. Our study is the first to identify the role resistin plays in promoting the homing of EPCs to the synovium and the subsequent production of VEGF during angiogenesis in RA.
The VEGF-related activation of PKC- has been previously observed, and PKC-
may regulate a number of pathways required for angiogenesis [50, 51]. Although different isoforms of PKC have been shown to regulate various cellular molecular responses [52], Gliki et al found that Rottlerin was not specific for PKC- [53]. Here, we additionally used PKC- siRNA to confirm PKC- function in EPCs. Our findings indicate that one of the earliest steps in sprouting angiogenesis requires the PKC-
and the downstream AMPK signaling pathway in resistin-promoted EPC migration and tube formation.
As miRNAs are able to modulate target gene translation via binding to the 3ʹUTR of mRNAs and thus affecting multiple protein-encoding genes at the posttranscriptional level, they are implicated in the control of a wide range of biological functions [54]. Therefore, we investigated whether miRNAs were involved in angiogenesis following resistin stimulation of EPCs. We first found that of 384 human miRNAs in a customized microRNA Profiler Kit, approximately one-fifth were influenced by resistin in EPCs. We subsequently identified 11 miRNAs that underwent at least a two-fold change in expression in response to resistin. Both clinical and experimental studies have shown that miRNAs appear to negatively regulate angiogenic events, including neovascularization and VEGF production [55, 56]. Therefore, we looked for miRNAs that were predicted by multiple algorithms (TargetScan, miRDB, and PicTar) to downregulate VEGF-A, and identified miR-206 as a possible candidate.
Recent research has provided growing evidence for the importance of miR-206 in development and proliferation of adenocarcinoma and lipid metabolism [57, 58].
Carlos et al. reported that miR-1/miR-206 negatively regulated angiogenesis in zebrafish [59]. However, a role for miR-206 in linking EPCs and RA pathogenesis has not previously been identified. In this study, we used a miR-206 mimic or inhibitor and a luciferase reporter containing a miR-206 binding site on the VEGF 3ʹUTR to verify the direct effects of miR-206 on resistin-elevated VEGF regulation. We found that resistin, via the downregulation of miR-206 targeting of VEGF 3ʹUTR, functions as a critical angiogenic participant and a regulator of circulating EPCs. In addition, we performed in vivo Matrigel implants and CAM assays to investigate the effects of EPCs on recruitment of cytokines such as VEGF and angiogenesis in articular microenvironments. Our results suggest that resistin when present in high levels in synovial fluid from RA patients promotes EPCs migration and tube formation.
In conclusion, our study is the first to identify the role of EPC homing to the 486
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synovium during RA angiogenesis and the signaling pathway involved in resistin- induced VEGF expression in primary EPCs. We found that resistin, which is highly expressed in patients with RA, causes EPC homing to the synovium and VEGF- dependent tube formation through the negative regulation of miR-206 via the PKC- δ/AMPK signaling pathway. EPC infiltration into the synovium results in subsequent angiogenesis. Thus, resistin may be a novel therapeutic target in patients with RA.
Our findings may open a new aspect on the nature of the signal transduction of progenitor cells and provide a better understanding of the mechanisms underlying RA pathogenesis.
Acknowledgments
This work was supported by grants from the Ministry of Science and Technology of Taiwan (103-2628-B-039-002-MY3, 102-2632-B-715-001-MY3, and 101-2314-B- 039-002-MY3) and China Medical University, Taiwan (CMU 103-S-06).
Conflict of interest
The authors declare that there are no conflicts of interest.
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FIGURE LEGENDS
Figure 1. Resistin is highly expressed in synovial fluid and tissue from patients with rheumatoid arthritis and from mice with collagen-induced arthritis. (A, B) Synovial fluid or tissue from 10 patients with rheumatoid arthritis (RA) and normal (NL) synovial fluid from 4 healthy patients undergoing arthroscopy for trauma/joint derangement were measured for levels of resistin protein or mRNA by enzyme-linked immunosorbent assay or real-time quantitative polymerase chain reaction, respectively. (C) NL primary cells and human synovial fibroblasts from RA patients were studied for resistin expression by Western blot. (D) Circulating endothelial progenitor cells (EPCs) isolated from human peripheral blood were examined in a migration assay using RA or NL synovial fluids with or without resistin neutralizing antibody. (E) Representative micro computed tomography images of the knees of mice with collagen-induced arthritis (CIA) on day 42 (F, femur; T, tibia; P, patella; M, meniscus). Scale bar: 1mm. (F) Quantification of volumetric bone mineral density (BMD) from femur and tibia. (G) Knee joint sections were stained with hematoxylin and eosin (H &E) and safranin O. Scale bar: 10 m (H) Tissue sections of synovium from control and CIA mice were detected with CD34 (CD34-positive cells, red) and CD133 (CD133‐positive cells, green) by using double immunofluorescent staining;
DAPI (blue) was used for nuclear staining. Arrows indicate tri-fluorescent tagged EPCs in the synovium. Scale bar: 5 m. (I) Knee joint tissue sections from control and CIA mice were immunohistochemically stained for resistin, CD133, vascular endothelial growth factor receptor 2 (VEGFR2), CD31, and VEGF. Arrows indicate dye-tagged targets in the synovium of the CIA mice. Scale bar: 5 m and 3 m (inset). Bars show the mean ± S. E. M. from 3 individual experiments; *P < 0.05 versus NL (A, B, and D) or control (F); #P< 0.05 versus RA fluids (D).
Figure 2. Resistin promotes vascular endothelial growth factor–dependent endothelial progenitor cell migration and tube formation. (A) Treatment of endothelial progenitor 708
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cells (EPCs) with resistin (resistin; 0.03–3 ng/mL) for 24 h induced vascular endothelial growth factor (VEGF) mRNA expression in a concentration-dependent manner. (B) Resistin stimulation increased VEGF protein expression in a concentration-dependent manner, as shown by enzyme-linked immunosorbent assay.
(C, D) Transfection with small interfering RNA (siRNA) targeting VEGF or with control siRNA for 16–18 h abolished the resistin-induced increase in EPC migration and tube formation. (E) The number of tube branches and the total tube length were calculated and quantified. Bars show the mean ± S. E. M. from 3 individual experiments; *P < 0.05 versus control; #P< 0.05 versus resistin (3 ng/mL) (C, E).
Figure 3. The protein kinase C delta-signaling pathway is involved in resistin-induced endothelial progenitor cell migration and tube formation. (A) Endothelial progenitor cells (EPCs) were incubated with resistin (3 ng/mL) for the indicated time intervals, and then protein kinase C delta (PKC-) phosphorylation was analyzed by Western blot. (B) The effect of small interfering RNA (siRNA) targeting PKC- (2 M) on EPCs vs. that of control siRNA, as analyzed by Western blot (upper graph). The upregulation of vascular endothelial growth factor (VEGF) mRNA in response to resistin was reversed by pretreatment with the PKC- inhibitor Rottlerin or PKC-
siRNA, as shown by real-time quantitative polymerase chain reaction (lower graph).
(C) Pretreatment with Rottlerin or transfection with PKC- siRNA abolished resistin- induced VEGF production, as shown by the enzyme-linked immunosorbent assay.
(D–F) Resistin-promoted EPC migration and tube formation were reduced by pretreatment with Rottlerin or transfection with PKC- siRNA. (E) Quantification of tube formation. Data represent mean ± S. E. M. from 3 individual experiments;
*P < 0.05 versus control; #P < 0.05 versus resistin alone or control siRNA (B–D, F).
Figure 4. The role of AMP-activated protein kinase α 1 in resistin-induced endothelial progenitor cell migration and tube formation. (A) Endothelial progenitor cells (EPCs) were incubated with resistin (3 ng/mL) for the indicated time intervals, and then phosphorylated AMP-activated protein kinase 1 (AMPK1) was measured by Western blot. (B) EPCs were pretreated with Rottlerin for 30 min or transfected with small interfering RNA (siRNA) targeting PKC- for 16–18 h and then stimulated with resistin; subsequently, phosphorylated AMPK1 was measured by Western blot.
(C) EPCs were transfected with AMPK1 siRNA or control siRNA (2 M), and then the level of AMPK1 protein expression was measured by Western blot (upper graph). EPCs were pretreated with the AMPK inhibitors Ara A or Compound C, or transfected with AMPK1 siRNA and stimulated with resistin; then, levels of VEGF mRNA were assessed by real-time quantitative polymerase chain reaction (lower 745
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graph). (D) ELISA assay indicated that resistin-mediated vascular endothelial growth factor (VEGF) production was abolished by pretreatment with Ara A or Compound C, or transfection with AMPK1 siRNA. (E–G) Resistin-induced EPC migration and tube formation were reduced by pretreatment with Ara A and Compound C or transfection with AMPK1 siRNA, as shown by quantification of tube formation (G).
Data represent mean ± S. E. M. from 3 individual experiments; *P < 0.05 versus control; #P < 0.05 versus resistin alone or control siRNA (C–E, G).
Figure 5. Resistin induces endothelial progenitor cell migration and tube formation by downregulating microRNA-206 expression via the protein kinase C delta/AMP- activated protein kinase-signaling pathway. (A) Endothelial progenitor cells (EPCs) were incubated with resistin (1–3 ng/mL) for 24 h, and then microRNA (miR)-206 expression was examined by real-time quantitative polymerase chain reaction (qPCR).
(B–D) EPCs were transfected with an miR-206 mimic (10 nM), an miR-206 inhibitor (10 nM), or a negative control (NC) miR (10 nM) followed by resistin treatment; then, vascular endothelial growth factor (VEGF) expression was evaluated by Western blot (B), qPCR (C), and enzyme-linked immunosorbent assay (D). (E–G) Resistin-induced EPC migration and tube formation were attenuated by transfection with a miR-206 mimic and significantly enhanced by transfection with an miR-206 inhibitor. (H) EPCs were transfected with indicated luciferase plasmids for 24 h and then treated with resistin, after which the relative luciferase activity was measured. (I) Pretreatment with Rottlerin, Compound C, or Ara A, or transfection with small interfering RNA (siRNA) targeting either protein kinase C delta (PKC-δ) or AMP- activated protein kinase (AMPK) reversed the resistin-mediated downregulation of miR-206 expression in EPCs. Data represent mean ± S. E. M. from 3 individual experiments; *P < 0.05 versus control; #P < 0.05 versus resistin alone or NC miR (C–
E, G).
Figure 6. Resistin is associated with endothelial progenitor cell–mediated angiogenesis in vivo. (A) Chick chorioallantoic membrane (CAM) assay using 5-day- old fertilized chick embryos (6 eggs/group): PBS (vehicle) or resistin (3 ng/mL) were resuspended in Matrigel and placed onto the CAMs, which were allowed to develop for another 3 days. The CAMs were then examined by microscopy and photographed.
Scale bar: 2 mm. (B) Vessels on the CAMs were calculated and quantified. (C, D) Matrigel plugs containing PBS (vehicle) and resistin (3 ng/mL) were subcutaneously injected into the flanks of nude mice. After 7 days, the plugs were photographed (C) and hemoglobin levels were quantified and normalized to those obtained with PBS (D). (E) Specimens from the plugs were immunostained with antibodies against 783
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VEGF, CD31, and CD133. Scale bar: 5 m. (F) Micrographs of tube formation by endothelial progenitor cells in normal (NL) and RA fluid. (G) The number of tube branches and the total tube length were calculated and quantified. (H) CAM assay using 5-day-old fertilized chick embryos (6 eggs/group): NL fluid, RA fluid, or RA fluid plus resistin neutralized antibody was resuspended in Matrigel and placed onto the CAMs, which were allowed to develop for another 3 days. The CAMs were then examined by microscopy and photographed. Scale bar: 2 mm. (I) Vessels on the CAMs were calculated and quantified. (J) After 7 days incubation, matrigel plugs containing NL fluid, RA fluid, or RA fluid plus resistin neutralized antibody were photographed. (K) Matrigel plugs were quantified with hemoglobin levels and normalized to those obtained with NL fluid. Data represent mean ± S. E. M. from 3 individual experiments; *P < 0.05 versus vehicle (B and D) or NL fluids (G, I, and K);
#P < 0.05 versus RA fluids (G, I, and K).
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