Osteoprotegerin Causes Apoptosis of Endothelial Progenitor Cells by Induction of Oxidative Stress

Authors


Abstract

Objective

Elevated serum osteoprotegerin (OPG) levels represent an independent risk factor for atherosclerotic disease, although the underlying mechanism is not clear. The aim of this study was to investigate the association of serum OPG levels and circulating endothelial progenitor cell (EPC) numbers, and to explore the effect of OPG on EPC apoptosis and its underlying mechanisms.

Methods

Flow cytometry was used to enumerate EPCs in the peripheral blood of 91 patients with systemic lupus erythematosus (SLE). Cultured EPCs, isolated from peripheral blood, were challenged with OPG, and apoptosis was evaluated by TUNEL staining. Expression of apoptosis-related proteins was measured by real-time quantitative polymerase chain reaction (qPCR) and Western blotting. Reactive oxygen species (ROS) were detected by flow cytometry, and the expression of NADPH oxidase (NOX) and MAP kinases (MAPK) was measured by qPCR and Western blotting.

Results

The serum OPG level was independently associated with reduced numbers of EPCs in patients with SLE. In vitro treatment with OPG significantly induced apoptosis of EPCs; this effect was mediated by syndecan 4. OPG-induced apoptosis was abolished by the ROS scavenger N-acetylcysteine and the NOX inhibitor diphenyleniodonium. OPG increased ROS production through activation of NOX-2 and NOX-4 and triggered phosphorylation of ERK-1/2 and p38 MAPK. Quenching of ROS by knockdown of NOX-2 or NOX-4 transcripts inhibited phosphorylation of ERK-1/2 and p38 MAPK. Moreover, inhibitors of ERK-1/2 and p38 MAPK decreased ROS production and subsequent EPC apoptosis, indicating a feed-forward loop between NOX and MAPK to amplify ROS production related to apoptosis.

Conclusion

Elevated OPG levels increase apoptosis of EPCs by induction of oxidative stress.

Systemic lupus erythematosus (SLE) is an autoimmune disorder with a strong predilection for atherosclerotic cardiovascular disease (CVD) ([1]). The pathogenesis of atherosclerosis in SLE is not completely elucidated but might be attributable to profound endothelial cell dysfunction ([2]). In a previous study, elevated levels of circulating apoptotic endothelial cells were observed in patients with active SLE and strongly correlated with endothelial cell dysfunction ([3]). This indicates that diffuse vascular damage occurs in SLE and participates in the promotion of atherosclerosis, because apoptotic endothelial cells are prothrombotic ([4]), and endothelial cell denudation is known to initiate plaque formation ([5]). However, the overall integrity of endothelium may depend not only on the extent of injury but also on the endogenous capacity for repair, which involves the recruitment of endothelial progenitor cells (EPCs) from circulating blood ([6]).

Reduced circulating EPC counts are associated with an increased risk of CVD and vascular complications ([7, 8]). In patients with SLE, the circulating EPC count was shown to be decreased compared with that in healthy subjects ([9]). Previous studies have suggested an increased propensity of EPCs to undergo apoptosis as the underlying cause of reduced numbers of EPCs in SLE. Indeed, the percentage of apoptotic EPCs is significantly higher in patients with SLE compared with healthy controls ([10]). Evidence is emerging that several factors associated with traditional and disease-related risk elicit apoptosis of EPCs. These include C-reactive protein, oxidized low-density lipoprotein, and interferon-α ([11-13]).

Elevated levels of osteoprotegerin (OPG) have been observed in chronic inflammatory diseases such as atherosclerosis, rheumatoid arthritis, and SLE ([14-16]). This increase has been suggested to reflect a compensatory response to enhanced osteoclast activity or negative bone remodeling balance occurring in the setting of chronic inflammation. However, recent studies have demonstrated the pathogenetic role of OPG in atherogenesis. These actions are supported by the finding of OPG in atherosclerotic human vessels ([17]) and the up-regulation of OPG by proinflammatory cytokines, such as tumor necrosis factor α (TNFα) or interleukin-1β, in endothelial cells and smooth muscle cells ([18, 19]). Conversely, in vitro treatment with OPG is able to promote the proliferation of vascular smooth muscle cells ([20]) and increase the proadhesive activity of TNFα by enhancing the endothelial expression of vascular cell adhesion molecule 1, intercellular adhesion molecule 1, and E-selectin, which are considered biomarkers of endothelial cell dysfunction ([21]). Thus, these cell types are thought to be implicated as the target as well as the cellular source of OPG. Furthermore, a pathologic role of OPG was also supported by an in vivo study showing that administration of OPG promoted leukocyte rolling and adhesion to rat mesenteric postcapillary venules ([22]). Moreover, high serum OPG levels are associated with endothelial dysfunction in patients with type 2 diabetes ([23]) and with markers of oxidative stress in patients undergoing dialysis ([24]).

On the basis of these considerations, we investigated the association between the OPG level and the number of EPCs, defined by CD34+/vascular endothelial growth factor receptor 2 (VEGFR-2)+ cells, and determined the underlying mechanisms of OPG involved in the reduction of EPC counts. However, because of the low number of CD34+/VEGFR-2+ cells in the peripheral circulation, we chose to isolate EPCs by culturing the mononuclear cell fraction of peripheral blood in order to collect in vitro qualitative data. In parallel, we sorted CD34+ cells from the peripheral blood of healthy control subjects and patients with SLE to complement the results obtained using cultured EPCs.

PATIENTS AND METHODS

Patient population

Ninety-one patients with SLE (9 men and 82 women) who fulfilled the American College of Rheumatology revised criteria for the classification of SLE ([25]) were recruited from the Division of Rheumatology at Yeouido St. Mary's hospital (Seoul, Korea). Fifty age- and sex-matched healthy control subjects (6 men and 44 women) were also included. Patients were excluded if any of the following features were present: preexisting overt coronary artery disease, transient ischemic attack, stroke, congestive heart failure, renal failure (defined by a serum creatinine level of ≥3.0 mg/dl), pregnancy, or active infection. In patients with SLE, clinical and laboratory data were obtained at the time of sampling. Serum OPG levels were determined by a commercial enzyme-linked immunosorbent assay (ELISA) kit (R&D Systems). All subjects gave written consent before entering the study, which was approved by the institutional review board of the Catholic University of Korea (no. SC08TISI0159).

Enumeration of circulating EPCs by flow cytometry

Quantification of circulating EPCs was performed by flow cytometry, as described previously ([26]). Briefly, mononuclear cells (MNCs) were isolated from peripheral blood by Ficoll density-gradient centrifugation. Cells were incubated with fluorescein isothiocyanate (FITC)–conjugated anti-human CD34 antibody (BD Biosciences) and allophycocyanin (APC)–conjugated anti-human VEGFR-2 antibody (R&D Systems). Isotype-identical antibodies served as controls. After incubation, cells were analyzed with a FACSCalibur flow cytometer (BD Biosciences). Each analysis consisted of 300,000 events. The percentages of positive cells were converted to cells per milliliter of blood, using the complete blood cell count.

Isolation and identification of EPCs and CD34+ cells

Peripheral blood MNCs from healthy volunteers and patients with SLE were isolated by density-gradient centrifugation and plated on culture dishes coated with fibronectin (BD Biosciences). Cells were cultured in endothelial growth medium 2 (Cambrex). After 3 days of culture, nonadherent cells were removed, and attached cells (5 × 105/ml) were cultured on new fibronectin-coated plates in endothelial basal medium 2 (EBM-2) supplemented with 2% fetal bovine serum (FBS) and stabilized for 24 hours. To confirm EPC phenotype, cells were stained with DiI-labeled acetylated low-density lipoprotein (DiI-AcLDL; Biomedical Technologies) followed by counterstaining with FITC-labeled Ulex europaeus agglutinin type I (UEA-I; Sigma-Aldrich). Double-positive cells were analyzed by flow cytometry. EPCs were further identified by demonstrating the expression of VEGFR-2 (R&D Systems), von Willebrand factor (vWF; Abcam), CD14 (eBioscience), and CD34 (BD Biosciences).

CD34+ cells were isolated from the peripheral MNCs of healthy subjects and patients with SLE, using a CD34 MicroBead kit (Miltenyi Biotech) according to the manufacturer's instructions. The purity of isolated CD34+ cells was >95%, as measured by flow cytometry.

TUNEL assay

After culturing for 3 days, adherent cells were harvested and seeded in 8-well Lab-Tek Chamber Slides (Nunc) coated with fibronectin in EBM-2 supplemented with 2% FBS and stabilized for 24 hours. EPCs were then incubated with recombinant OPG (R&D Systems) for the indicated periods of time. In some experiments, EPCs were pretreated with caspase 3 inhibitor (Z-DEVD-FMK; Calbiochem), heparinase I (Sigma-Aldrich), chondroitinase ABC (Sigma-Aldrich), anti–syndecan 2/anti–syndecan 4 (Santa Cruz Biotechnology), diphenyleneiodonium chloride (DPI; Sigma-Aldrich), N-acetylcysteine (NAC; Sigma-Aldrich), SB203580 (Sigma-Aldrich), PD98059 (Sigma-Aldrich), and LY294002 (Sigma-Aldrich) for 1 hour before incubation with OPG. EPCs were then treated with varying concentrations of OPG for 24 hours. Apoptosis of EPCs was evaluated by using an In Situ Cell Death Detection Kit (Roche).

Measurement of intracellular reactive oxygen species (ROS).

ROS formation was measured by the detection of 2′,7′-dichlorofluorescein diacetate (H2DCFDA), using a flow cytometer. EPCs were plated on 24-well plates coated with fibronectin in EBM-2 supplemented with 2% FBS and stabilized for 24 hours. EPCs were then treated with varying concentrations of OPG for 24 hours. For some experiments, EPCs were preincubated in the presence of DPI, NAC, SB203580, PD98059, or LY294002 for 1 hour before treatment with OPG. In selected wells, CD34+ cells isolated from the peripheral blood of healthy subjects and patients with SLE were treated with OPG. Cells were then incubated in H2DCFDA solution (10 μM; Invitrogen) for 20 minutes. After incubation, cells were washed with PBS twice and subjected to flow cytometric analysis.

Small interfering RNA (siRNA) transfection

Syndecan 4, NADPH oxidase 2 (NOX-2), NOX-4, and control siRNA were purchased from Santa Cruz Biotechnology. Briefly, EPCs were seeded in fibronectin-coated 24-well plates at 5 × 105 cells per well in EBM-2 supplemented with 2% FBS and grown to 60–80% confluence. Cells were then transfected with siRNA using Lipofectamine 2000 reagent (Invitrogen) according to the manufacturer's protocol. After overnight stabilization, transfected cells were incubated with OPG for the indicated periods of time and then subjected to real-time polymerase chain reaction (PCR), flow cytometry, and Western blot analysis.

Real-time quantitative PCR analysis

Total RNA was extracted from the EPCs, CD34+ cells, and CD34+/VEGFR-2+ cells using an RNeasy Mini Kit (Qiagen). The RNA was reverse transcribed with Moloney murine leukemia virus reverse transcriptase and random hexaprimers (Promega). The 80–150-bp primers for each gene were purchased from Bioneer. Quantitative assessment of target messenger RNA (mRNA) levels was performed by real-time PCR with a Bio-Rad CFX96 Real-Time PCR Detection System. The quantity of mRNA was calculated using the 2−ΔΔCt method, and GAPDH was used to normalize total RNA quantities.

Immunoprecipitation assay

Binding of OPG to syndecan 4 was performed by immunoprecipitation assay as described previously ([27]). Briefly, cultured EPCs (2 × 108) were washed 3 times with PBS and were either left untreated or were treated with OPG for 1 hour at 4°C. The cells were incubated with 5 mM BS3 (Pierce) for crosslinking of OPG with the receptor. After 30 minutes, the crosslinking reaction was quenched with 15 mM Tris for 15 minutes. Cell lysates were immunoprecipitated with monoclonal anti–syndecan 4 antibody (Santa Cruz Biotechnology) and control mouse IgG. The immunoprecipitates and cell lysates were analyzed by immunoblotting with rabbit polyclonal antibodies against OPG and syndecan 4 (Santa Cruz Biotechnology).

Western blot analysis

Rabbit monoclonal antibodies against caspase 3, total/phospho–p38 MAPK, and total/phospho–ERK-1/2 were purchased from Cell Signaling Technology. Mouse monoclonal antibodies against Bax and Bcl-2, rabbit polyclonal antibodies against syndecan 2 and syndecan 4, and rabbit polyclonal antibody against NOX-4 were purchased from Santa Cruz Biotechnology. Rabbit polyclonal antibody against NOX-2 was purchased from Millipore. Cellular proteins from EPCs under various treatments were resolved by 10% sodium dodecyl sulfate–polyacrylamide gel electrophoresis and probed with different primary antibodies as specified above. Horseradish peroxidase–conjugated secondary antibodies (anti-mouse or anti-rabbit) were used in conjunction with an Amersham ECL Chemiluminescent Detection system (Amersham).

Statistical analysis

Comparisons of numerical data between groups were performed by Student's t-test or analysis of variance, and comparisons of categorical data between groups were performed by chi-square test or Fisher's exact test, as appropriate. Correlations between 2 variables were determined using Spearman's rank correlation coefficient. P values less than 0.05 were considered significant.

RESULTS

Inverse correlation between serum OPG levels and numbers of circulating EPCs

Ninety-one patients with SLE and 50 healthy control subjects were enrolled in the study. The demographic and clinical characteristics of these groups are shown in Supplementary Table 1, available online at http://www.ribjd.com/data. As we observed previously ([16]), serum OPG levels were significantly higher in patients with SLE compared with control subjects. The number of EPCs, as determined by coexpression of CD34 and VEGFR-2, was significantly lower in patients with SLE (median 44 cells/ml, interquartile range [IQR] 17–89) compared with control subjects (median 86 cells/ml, IQR 70–112; P < 0.01) (see Supplementary Figure 1, available online at http://www.ribjd.com/data). Importantly, the number of EPCs was inversely correlated with serum OPG levels (γ = −0.309, P = 0.014). Multiple regression analysis showed that the serum OPG level was an independent predictor of reduced numbers of EPCs in a model including age, hypertension, serum creatinine, lipid profile, and drug treatment, all of which may affect the serum OPG level and EPC numbers (β = −0.427, P = 0.018).

OPG-induced apoptosis of EPCs and CD34+ cells

Having observed an inverse correlation between OPG levels and EPC counts, we sought to determine whether a decreased number of EPCs was a consequence of apoptosis induced by OPG. Because of the low number of CD34+/VEGFR-2+ cells in peripheral blood, we isolated EPCs by culturing the MNC fraction of peripheral blood derived from healthy subjects and patients with SLE, as described previously ([8, 28]). EPCs were characterized as adherent cells that were double-positive for DiI-acLDL uptake and UEA-1 lectin binding. These cells also expressed the endothelial markers VEGFR-2 and vWF, the monocyte-lineage cell marker CD14, and, to some degree, the stem cell marker CD34 (see Supplementary Figure 2, available online at http://www.ribjd.com/data).

First, we examined the effects of OPG on apoptosis of EPCs. As shown in Figure 1A, recombinant human OPG caused a dose-dependent increase in apoptosis of EPCs from healthy subjects, which was completely abrogated by pretreatment with caspase 3 inhibitor (Z-DEVD-FMK). As expected, OPG increased the expression of cleaved caspase 3 in a dose-dependent manner, and pretreatment with caspase 3 inhibitor completely suppressed the increased expression of cleaved caspase 3 (Figure 1C). Interestingly, EPCs from patients with SLE showed significantly increased baseline apoptosis compared with those from healthy controls (P < 0.01) (Figure 1B), which is in agreement with a previous report ([10]). OPG also enhanced the apoptosis of EPCs from patients with SLE (P < 0.01). The percentage of OPG-induced EPC apoptosis was significantly higher in patients with SLE than in healthy controls, albeit the fold increase in OPG-induced apoptosis was lower in patients than in controls (1.8-fold versus 3.7-fold; P < 0.05). These results suggested enhanced activation of apoptosis in EPCs from patients with SLE, because these cells had already been exposed to apoptotic stresses and were close to achieving full activation of apoptosis in vivo.

Figure 1.

A, Effect of osteoprotegerin (OPG) on endothelial progenitor cell (EPC) apoptosis. EPCs were treated with OPG (0–20 ng/ml) for 24 hours in the absence or presence of 20 μM caspase 3 inhibitor (INH), and apoptotic cells were identified by TUNEL staining. Left, Representative images showing dark brown staining of apoptotic nuclei in OPG-treated EPCs. Right, Dose-dependent induction of EPC apoptosis by OPG. ∗ = P < 0.01 and ∗∗ = P < 0.001 versus control; ∗∗∗ = P < 0.01 versus 20 ng/ml OPG. B, Apoptosis of EPCs from patients with systemic lupus erythematosus (SLE) and normal healthy controls (NHC). ∗ = P < 0.01 versus OPG-untreated SLE; ∗∗ = P < 0.001 and ∗∗∗ = P < 0.01 versus OPG-untreated NHC. C and D, Effects of OPG on the expression of apoptosis-related proteins, as determined by Western blotting. Cultured EPCs and CD34+ cells were treated with 0–20 ng/ml OPG and 10 ng/ml OPG, respectively, for 24 hours. ∗ = P < 0.01 versus control; ∗∗ = P < 0.05 versus 20 ng/ml OPG. Bars show the mean ± SD of triplicate experiments (n = 3 donors).

It has been known that the ratio of Bax to Bcl-2 (Bax/Bcl-2 ratio) is critical for the induction of apoptosis, and that an increasing Bax/Bcl-2 ratio leads to activation of caspase 3 and determines the susceptibility of a cell to undergo apoptosis ([29, 30]). Thus, we examined the expression of Bax and Bcl-2 in EPCs, using real-time PCR and Western blot analysis. Consistent with the TUNEL assay results shown in Figure 1B, EPCs from patients with SLE showed lower basal expression of Bcl-2 but higher basal expression of Bax and caspase 3 as well as an elevated Bax/Bcl-2 ratio compared with control (Figure 2A). After OPG stimulation, the expression of Bcl-2 was decreased at both the mRNA and protein levels, but the expression of Bax and caspase 3 was up-regulated (Figures 1C and 2C). As a result, the Bax/Bcl-2 expression ratio was augmented in EPCs treated with OPG. Similar results were observed when EPCs from patients with SLE were treated with OPG (data not shown).

Figure 2.

A and B, Basal expression of Bcl-2, Bax, Bax/Bcl-2, and caspase 3 mRNA in cultured EPCs (A) and CD34+ cells (B) from patients with SLE (n = 8) and healthy control subjects (n = 8). C and D, Changes in Bcl-2, Bax, Bax/Bcl-2, and caspase 3 mRNA expression in EPCs (C) and CD34+ cells (D) treated with OPG. Cultured EPCs and CD34+ cells from healthy controls (n = 3) were treated with OPG (0–20 ng/ml) and 10 ng/ml OPG, respectively. Real-time polymerase chain reaction was performed to determine Bcl-2, Bax, and caspase 3 mRNA expression levels in cultured EPCs (A and C) and in CD34+ cells (B and D). Data are presented as the fold change relative to control. Bars show the mean ± SD of triplicate experiments. ∗ = P < 0.05, ∗∗ = P < 0.01 versus healthy controls; † = P < 0.05, ‡ = P < 0.01 versus control (no OPG). See Figure 1 for definitions.

To further assess whether OPG also induces apoptosis of CD34+ cells, CD34+ cells from the peripheral blood of healthy donors (n = 3) and patients with SLE (n = 3) were enriched by performing immunomagnetic cell sorting with anti-CD34 antibody, followed by incubation with OPG. As observed in EPCs, OPG decreased Bcl-2 expression and concomitantly increased Bax expression, the Bax/Bcl-2 expression ratio, and caspase 3 expression in CD34+ cells (Figures 1D and 2D). These data unequivocally demonstrated that apoptosis induced by OPG occurs not only in cultured EPCs but also in circulating progenitor cells expressing CD34.

Role of syndecan 4 in OPG-induced EPC apoptosis

The heparin-binding domain of OPG has the potential to interact with numerous proteoglycans on cell surfaces. Previous studies have demonstrated that OPG binds to monocytes via cell surface heparin sulfate proteoglycans, such as syndecan, and thus facilitates chemotaxis of monocytes ([31]). To determine the role of heparin sulfate proteoglycans in OPG-induced apoptosis, EPCs were pretreated with heparinase I and chondroitinase ABC. As shown in Figure 3A, the apoptotic effects of OPG were completely blocked by pretreatment with heparinase I but not by chondroitinase ABC, suggesting the involvement of syndecan 2 and syndecan 4 in apoptosis.

Figure 3.

A, Effects of heparinase I (Hep) and chondroitinase ABC (Chon) on OPG-induced EPC apoptosis. EPCs were preincubated with heparinase I (20 mU/ml) or chondroitinase ABC (20 mU/ml) for 1 hour and then treated with OPG (20 ng/ml) for 24 hours. Apoptosis was determined by TUNEL staining. B, Expression of syndecan 1 (SDC-1) SDC-2, SDC-3, and SDC-4 in EPCs from 3 healthy donors, as determined by reverse transcription–polymerase chain reaction (RT-PCR) and Western blotting. Surface expression of SDC-2 and SDC-4 was confirmed by flow cytometry. C, Differential regulation of SDC-2 and SDC-4 by OPG. EPCs were treated with OPG, and the expression of SDC-2 and SDC-4 was determined by real-time PCR and Western blotting. Inset shows a representative Western blot. D, Involvement of SDC-4 in OPG-induced apoptosis. EPCs were pretreated with 5 μg/ml of control IgG, anti–SDC-2, or anti–SDC-4 antibody for 1 hour and then incubated with OPG (20 ng/ml) for 24 hours. Bars in A, C, and D show the mean ± SD of triplicate experiments (n = 3 donors). ∗ = P < 0.05, ∗∗ = P < 0.001 versus control; ∗∗∗ = P < 0.01 versus OPG. E, Binding of OPG to SDC-4. Cell lysates of EPCs treated with or without OPG were prepared as described in Patients and Methods and immunoprecipitated (IP) with anti–SDC-4 antibody and control mouse IgG. The immunoprecipitates and cell lysates were analyzed by immunoblotting (IB) with anti-OPG and anti–SDC-4 antibodies. Results are representative of 3 independent experiments. See Figure 1 for other definitions.

Furthermore, we studied the expression of syndecan family members on resting and stimulated EPCs from 3 different healthy donors. Both syndecan 2 mRNA and syndecan 4 mRNA were expressed at high levels in unstimulated EPCs, and the protein expression of syndecan 2 and syndecan 4 was confirmed by Western blotting and fluorescence-activated cell sorting analysis (Figure 3B). OPG dose-dependently up-regulated syndecan 4 expression in EPCs, but it did not change syndecan 2 expression (Figure 3C). Preincubation of EPCs with anti–syndecan 4 antibody abolished the OPG-induced apoptosis, whereas neither anti–syndecan 2 antibody nor isotype-matched IgG had an effect (Figure 3D). In addition, immunoprecipitation assay revealed that OPG could coimmunoprecipitate syndecan 4, but control IgG did not immunoprecipitate any proteins (Figure 3E). Taken together, these data indicated that syndecan 4 acts as a receptor for OPG on EPCs, critically mediating EPC apoptosis upon OPG ligation.

Role of ROS in OPG-induced EPC apoptosis

Although ROS at low levels function as signaling molecules to regulate angiogenesis and vasculogenesis ([32]), excess amounts of ROS are toxic and linked to senescence and apoptosis of stem/progenitor cells ([33]). Thus, we investigated whether OPG induces intracellular ROS in EPCs. EPCs and CD34+ cells from patients with SLE had significantly higher basal production of ROS compared with those from healthy controls (Figures 4A and B) (P < 0.01 and P < 0.05, respectively). Treatment of EPCs with varying concentrations of OPG resulted in a dose-dependent increase in 2′,7′-dichlorofluorescein fluorescence (Figure 4A, bottom). ROS generation increased as early as 10 minutes after OPG treatment, peaked at 6 hours, and persisted at high levels up to 24 hours (data not shown). ROS production also increased in CD34+ cells treated with OPG (Figure 4B, bottom). Furthermore, pretreatment with the antioxidant NAC completely blocked OPG-induced EPC apoptosis (Figure 4C). Similar results were obtained following pretreatment with the NOX inhibitor DPI. Taken together, these results indicated that ROS formation induced by OPG ligation is causally linked to EPC apoptosis and also raised the possibility that NOX-derived ROS may be responsible for apoptosis.

Figure 4.

A and B, Basal and OPG-induced reactive oxygen species (ROS) production in EPCs (A) and CD34+ cells (B) from healthy control subjects and patients with SLE. Cultured EPCs and CD34+ cells from healthy control subjects (n = 3) were treated with 0–20 ng/ml OPG and 10 ng/ml OPG, respectively, for 24 hours. ROS production was measured by flow cytometry. Representative flow cytometry data are shown (top). C and D, Inhibition of EPC apoptosis (C) and ROS generation (D) by ROS scavengers. EPCs were preincubated with diphenyleneiodonium chloride (DPI; 0.2 μM) or N-acetylcysteine (NAC; 0.5 mM) for 1 hour and then incubated with OPG for 24 hours. Bars show the mean ± SD of triplicate experiments (n = 3 donors). ∗ = P < 0.01, ∗∗ = P < 0.001 versus control; ∗∗∗ = P < 0.01 versus OPG. DCF = 2,7-dichlorofluorescein; MFI = mean fluorescence intensity (see Figure 1 for other definitions).

Control of OPG-induced ROS generation by NOX

NOX is known to be one of the major sources of ROS in endothelial cells and stem/progenitor cells ([34]). Thus, we analyzed the expression levels of NOX isoforms in EPCs, using real-time PCR and Western blotting. After OPG stimulation, the expression of NOX-2 and NOX-4 mRNA in EPCs was significantly up-regulated, while the expression of NOX-1 and NOX-3 was unchanged (Figure 5A). The increased expression of NOX-2 and NOX-4 in OPG-treated EPCs was also confirmed at the protein level, as determined by Western blot analysis (Figure 5A). Additionally, as observed in EPCs, the mRNA and protein expression of NOX-2 and NOX-4 in CD34+ cells was also significantly increased after OPG treatment (Figure 5B).

Figure 5.

A and B, Activation of NADPH oxidase (NOX) isoforms in OPG-treated EPCs (A) and CD34+ cells (B). EPCs (n = 3) and CD34+ cells (n = 3) from healthy donors were treated with OPG (10 ng/ml), and expression of NOX isoforms was determined by real-time polymerase chain reaction and Western blot analysis. Insets show the protein expression of NOX-2 and NOX-4. C, Decreased OPG-induced reactive oxygen species (ROS) production by knockdown of NOX-2 or NOX-4 transcripts. EPCs (n = 3) were transfected with control, NOX-2, or NOX-4 small interfering RNA (siRNA) and then incubated with OPG. Inset shows expression of NOX-2 and NOX-4 mRNA in EPCs transfected with NOX-2 or NOX-4 siRNA. D, Down-regulation of NOX-2 and NOX-4 mRNA expression in EPCs transfected with syndecan 4 (SDC-4) siRNA. EPCs (n = 3) were transfected with SDC-4 siRNA and then treated with OPG for 3 hours. Inset shows expression of SDC-2 and SDC-4 mRNA in EPCs transfected with SDC-4 siRNA. Bars in A–D show the mean ± SD of triplicate experiments (n = 3 donors). ∗ = P < 0.05, ∗∗ = P < 0.01 versus control; ∗∗∗ = P < 0.05 versus OPG-treated control. E, Correlation between the expression of SDC-4 mRNA and NOX-2 or NOX-4 mRNA in circulating EPCs. EPCs expressing both CD34 and vascular endothelial growth factor receptor 2 (VEGFR-2) were isolated from the peripheral blood of 24 patients with SLE using microbead-conjugated anti–CD34/VEGFR-2 antibody and then were subjected to real-time polymerase chain reaction to quantify the expression of SDC-4, NOX-2, and NOX-4 mRNA. See Figure 1 for other definitions.

Although DPI is widely used as a NOX inhibitor, it may not be specific for NOX-2 or NOX-4 ([35]). Therefore, we further assessed whether NOX-2 and NOX-4 are directly involved in OPG-induced ROS production, by performing specific knockdown of NOX genes using their respective siRNAs. Transfection of NOX-2 or NOX-4 siRNA, but not control siRNA, curtailed mRNA expression (Figure 5C). Flow cytometric analysis also revealed that ROS generation was significantly lower in EPCs transfected with NOX-2 or NOX-4 siRNA than in those transfected with control siRNA (Figure 5C). These results suggested that NOX-2 and NOX-4 are the major NOX isoforms responsible for OPG-induced ROS generation and subsequent apoptosis of EPCs.

To investigate whether OPG-activated syndecan 4 controls EPC apoptosis via NOX-2/4, we evaluated the effect of syndecan 4 knockdown on NOX-2 and NOX-4 expression. As expected, syndecan 4 siRNA substantially abrogated the OPG-induced increase in NOX-2 and NOX-4 mRNA expression, to an extent similar to that of control siRNA (Figure 5D). These observations suggest that OPG binding to syndecan 4 promotes EPC apoptosis by regulating NOX-2 and NOX-4 expression. To further address the clinical relevance of these findings, we measured the expression of syndecan 4 and NOX isoforms in CD34+/VEGFR-2+ cells, which were freshly isolated from the peripheral blood of patients with SLE. The results showed that syndecan 4 mRNA expression correlated with NOX-2 and NOX-4 mRNA expression of CD34+/VEGFR-2+ cells from patients with SLE (Figure 5E), which is in parallel with the in vitro down-regulatory effect of syndecan 4 siRNA on NOX-2 and NOX-4 (Figure 5D).

Mediation of OPG-induced EPC apoptosis by p38 MAPK and ERK-1/2.

Finally, we attempted to determine the signaling pathways involved in OPG-induced EPC apoptosis. Pharmacologic inhibitors of p38 MAPK (SB203580) and ERK-1/2 (PD98059), but not phosphatidylinositol 3-kinase (PI3K) inhibitor (LY294002), almost completely diminished EPC apoptosis induced by OPG (Figure 6A), suggesting a crucial role for p38 MAPK and ERK-1/2 in OPG-induced EPC apoptosis. In agreement with this, OPG elicited a rapid increase in phospho–p38 MAPK and phospho–ERK-1/2, which peaked as early as 5 minutes and was still detectable at 30 minutes (Figure 6B). Moreover, such an increase was completely inhibited by pretreatment of EPCs with DPI or by transfection with NOX-2 or NOX-4 siRNA (Figures 6C and D), indicating that activation of p38 MAPK and ERK-1/2 is redox sensitive. Alternatively, treatment of EPCs with SB203580 and PD98059, but not LY294002, completely abolished OPG-induced ROS production (Figure 6E), indicating that OPG-induced activation of p38 MAPK and ERK-1/2 is also required for ROS production.

Figure 6.

A, Inhibition of OPG-induced EPC apoptosis by p38 MAPK and ERK-1/2 inhibitors. EPCs were preincubated with 2 μM PD98059 (PD), SB203580 (SB), or LY294002 (LY) for 1 hour and then stimulated with OPG (20 ng/ml) for 24 hours. ∗ = P < 0.001 versus control; ∗∗ = P < 0.01 versus OPG alone. B, Activation of p38 MAPK and ERK-1/2 by OPG. EPCs were treated with OPG (10 ng/ml), and the expression of p38 MAPK and ERK-1/2 was determined by Western blot analysis. C and D, Suppression of OPG-triggered phosphorylation of p38 MAPK and ERK-1/2 by diphenyleneiodonium chloride (DPI) or by silencing of NADPH oxidase 2 (NOX-2) or NOX-4 transcripts. EPCs were preincubated with DPI for 1 hour and then treated with OPG for 30 minutes (C). EPCs were transfected with control, NOX-2, or NOX-4 small interfering RNA (siRNA) and then treated with OPG for 30 minutes (D). Results of densitometric analysis of phosphorylated p38 MAPK and ERK-1/2 are shown. ∗ = P < 0.05, ∗∗ = P < 0.001 versus control; ∗∗∗ = P < 0.01 versus OPG-treated control. E, Reduced OPG-induced reactive oxygen species (ROS) production by inhibitors of p38 MAPK and ERK-1/2 (SB and PD, respectively) but not by phosphatidylinositol 3-kinase (PI3K) inhibitor (LY). EPCs were preincubated with the inhibitors for 1 hour before OPG treatment. ∗ = P < 0.01 versus control; ∗∗ = P < 0.01 versus OPG alone. Bars in A and C–E show the mean ± SD of triplicate experiments (n = 3 donors). See Figure 1 for other definitions.

DISCUSSION

In the present study, we confirmed the results of previous studies that the number of circulating CD34+/VEGFR-2+ EPCs is reduced in patients with SLE ([9, 10]). Moreover, we observed that serum OPG levels were significantly higher in patients with SLE than in healthy controls and observed an inverse correlation between circulating EPCs and serum OPG levels in patients with SLE, suggesting a possible toxic effect of OPG on EPCs. Importantly, this study is the first to provide evidence that OPG promotes EPC apoptosis. In parallel, caspase 3 expression and the intracellular Bax/Bcl-2 ratio were increased in OPG-treated EPCs. This suggests OPG-induced regulation of Bax/Bcl-2/capase 3 in the process of EPC death ([36]), which might explain, at least in part, the decreased number of EPCs in the peripheral blood of patients with SLE.

When OPG was added to CD34+ cells freshly isolated from the peripheral blood of healthy subjects and patients with SLE, a similar increase in caspase 3 expression and the Bax/Bcl-2 ratio was observed in OPG-treated CD34+ cells. Given that increased apoptosis of CD34+ hematopoietic stem cells in patients with SLE reflects the decreased CD34+/VEGFR-2+ EPC subpopulation ([9]), these results indicate that OPG is toxic to CD34+ hematopoietic stem cells as well as to EPCs, which are critical for vascular repair (e.g., putative hemangioblastic and myeloid EPCs). Interestingly, analogous to our findings of decreased numbers of EPCs in patients with SLE, CD34+/VEGFR-2+ EPC and myeloid EPC counts were markedly reduced in patients undergoing dialysis ([37]). Furthermore, apoptosis of myeloid EPCs was induced by treatment with uremic serum but not by treatment with the specific uremic toxins tested, suggesting that unidentified toxins present in serum could be deleterious for EPCs ([37]). Because elevated serum OPG levels predict cardiovascular events in patients undergoing hemodialysis ([38]), it is conceivable that elevated serum OPG levels in these patients might be the culprit involved in EPC apoptosis.

It has been demonstrated that OPG displays ligand-independent biologic activities through its heparin-binding domain ([30, 39]). In mouse ovarian granulosa cells, syndecan 4 was prominently expressed in atretic follicles undergoing apoptosis ([40]). Also, syndecan 1 shed from the surface of myeloma cells was shown to mediate apoptosis in myeloma cells ([41]). However, there have been no data with respect to the direct role of the syndecan family in EPC apoptosis. In this study, we showed that syndecan 2 and syndecan 4 were prominently expressed in unstimulated EPCs at both the protein and mRNA levels. However, EPC apoptosis was almost completely inhibited by anti–syndecan 4 antibody but not by anti–syndecan 2 antibody. Moreover, the binding of OPG to syndecan 4 was confirmed by coimmunoprecipitation assay, further indicating that syndecan 4 is a predominant receptor that mediates OPG-induced EPC apoptosis.

Syndecan 4 works as a coreceptor of various growth factors, including fibroblast growth factor 2, and its expression is considerably enhanced by stimulation of growth factors ([42]). In the current study, the expression of syndecan 4, but not syndecan 2, was up-regulated by OPG treatment. Therefore, it is possible that increased syndecan 4 expression may further augment apopotic cell death by providing more binding sites for OPG. This assumption is bolstered by the results of a previous study showing that overexpression of syndecan 4 in rat embryonic fibroblasts enhances platelet-derived growth factor–induced ROS generation and MAPK activation ([43]), both of which are processes that have been implicated in apoptosis induction in EPCs ([44, 45]). These observations support the concept that syndecan 4 is an indispensable receptor for OPG binding and is likely to be a key regulator of the apoptotic action of OPG in EPCs.

The present study provides compelling evidence that the mechanism by which OPG promotes apoptosis of EPCs is tightly associated with ROS production. As shown in Figure 4, a ROS scavenger (NAC) and a NOX inhibitor (DPI) almost completely reversed OPG-induced EPC apoptosis. Given the OPG-induced regulation of Bcl-2/Bax/caspase 3, this further suggests that increased ROS production may act as an upstream regulator for mitochondrial membrane depolarization, inducing the release of cytochrome c and caspases, which leads to eventual EPC apoptosis. Moreover, the effect of OPG on ROS production was mediated by robust up-regulation of NOX-2 and NOX-4, because inhibition of ROS generation by NOX-2 and NOX-4 silencing completely abrogated EPC apoptosis. These findings are consistent with previous studies that demonstrated the involvement of these isoforms in EPC dysfunction and impaired angiogenesis ([46, 47]). For instance, overproduction of NOX-2–derived ROS contributes to impaired angiogenesis and EPC dysfunction in mice with type 1 diabetes, and this impaired angiogenesis is rescued in NOX-2–knockout mice ([46]). On the other hand, NOX-4 is a major source of ROS in EPC apoptosis induced by homocysteine. This apoptosis is reversed by treatment with atorvastatin through its inactivation of NOX-4 ([47]). Likewise, OPG-induced NOX-4 up-regulation is also suppressed by atorvastatin treatment (data not shown).

Oxidative stress induces the activation of MAPKs such as ERK, p38, and JNK, which have been implicated in the apoptotic response in several cells ([48, 49]). Consistently, we observed that OPG-induced EPC apoptosis was associated with up-regulation of p38 and ERK phosphorylation, and apoptosis was reversed by p38 MAPK and ERK inhibitors but not by PI3K inhibitor. In addition, the ERK/p38 MAPK pathway could act downstream of ROS to mediate OPG-induced EPC apoptosis, because quenching of ROS by either DPI treatment or siRNA silencing of NOX-2/NOX-4 decreased the phosphorylation of ERK and p38 MAPK. Conversely, inhibition of ERK and p38 MAPK using PD98059 and SB203580, respectively, markedly attenuated ROS production induced by OPG, suggesting that a feed-forward cycle may exist to amplify ROS generation related to OPG-induced EPC apoptosis. This notion is supported by previous observations that p38 MAPK could regulate ROS generation through the induction of NOX activity ([50]).

Our work underscores the importance of the OPG–syndecan 4–ROS axis in inducing EPC death, which is highly associated with CVD. To summarize, we first demonstrated that OPG promotes apoptosis of EPCs via engagement of syndecan 4. The apoptotic effect of OPG was mediated by the redox-sensitive ERK/p38 MAPK signaling pathway through activation of NOX-2 and NOX-4. This finding corroborates the clinical observation of an inverse association between serum OPG levels and circulating EPC counts in patients with SLE, suggesting that the atherogenic effects of OPG are attributable in part to its direct role in increasing apoptosis of EPCs. The relevance of our findings may extend beyond SLE to other conditions associated with high production of OPG, which is potentially involved in triggering premature atherosclerosis.

AUTHOR CONTRIBUTIONS

All authors were involved in drafting the article or revising it critically for important intellectual content, and all authors approved the final version to be published. Dr. Cho had full access to all of the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.

Study conception and design. J-Y. Kim, Y-J. Park, K-J. Kim, J-J. Choi, W-U. Kim, C-S. Cho.

Acquisition of data. J-Y. Kim, Y-J. Park, K-J. Kim, J-J. Choi, W-U. Kim, C-S. Cho.

Analysis and interpretation of data. J-Y. Kim, Y-J. Park, K-J. Kim, J-J. Choi, W-U. Kim, C-S. Cho.

Acknowledgments

We are grateful to Min-Jeung Kang and Dr. Eugene C. Yi (Department of Molecular Medicine and Biopharmaceutical Sciences, Seoul National University) for assistance with the immunoprecipitation assay.

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