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. 2016 Nov 7;291(52):26913–26921. doi: 10.1074/jbc.M116.741967

Smooth Muscle Cell Proangiogenic Phenotype Induced by Cyclopentenyl Cytosine Promotes Endothelial Cell Proliferation and Migration*

Rui Tang 1, Gui Zhang 1, Shi-You Chen 1,1
PMCID: PMC5207196  PMID: 27821588

Abstract

Vascular smooth muscle cells (SMCs) and endothelial cells (ECs) are in close contact with blood vessels. SMC phenotypes can be altered during pathological vascular remodeling. However, how SMC phenotypes affect EC properties remains largely unknown. In this study, we found that PDGF-BB-induced synthetic SMCs suppressed EC proliferation and migration while exhibiting increased expression of anti-angiogenic factors, such as endostatin, and decreased pro-angiogenic factors, including CXC motif ligand 1 (CXCL1). Cyclopentenyl cytosine (CPEC), a CTP synthase inhibitor that has been reported previously to inhibit SMC proliferation and injury-induced neointima formation, induced SMC redifferentiation. Interestingly, CPEC-conditioned SMC culture medium promoted EC proliferation and migration because of an increase in CXCL1 along with decreased endostatin production in SMCs. Addition of recombinant endostatin protein or blockade of CXCL1 with a neutralizing antibody suppressed the EC proliferation and migration induced by CPEC-conditioned SMC medium. Mechanistically, CPEC functions as a cytosine derivate to stimulate adenosine receptors A1 and A2a, which further activate downstream cAMP and Akt signaling, leading to the phosphorylation of cAMP response element binding protein and, consequently, SMC redifferentiation. These data provided proof of a novel concept that synthetic SMC exhibits an anti-angiogenic SMC phenotype, whereas contractile SMC shows a pro-angiogenic phenotype. CPEC appears to be a potent stimulator for switching the anti-angiogenic SMC phenotype to the pro-angiogenic phenotype, which may be essential for CPEC to accelerate re-endothelialization for vascular repair during injury-induced vascular wall remodeling.

Keywords: adenosine receptor, endothelial cell, migration, proliferation, vascular smooth muscle cells, cyclopentenyl cytosine, smooth muscle phenotype

Introduction

Pathological vascular remodeling is one of the major obstacles limiting the long-term clinical efficacy of cardiovascular intervention, including angioplasty, bypass surgery, and transplantation arteriopathy (1, 2). Endothelium denudation caused by injury promotes a series of pro-inflammatory responses (3, 4) that further impair endothelial cell (EC)2 and smooth muscle cell (SMC) functions (5–9). EC dysfunction may alter EC proliferation and migration, causing delayed endothelium repair with an increased risk of thrombosis (10, 11). It is unknown, however, whether the SMC phenotype affects EC properties during vascular remodeling and endothelium recovery.

SMC is known to exhibit a remarkable phenotypic plasticity (12–14). SMCs within adult arteries express contractile SMC markers, ion channels, and signaling molecules related to their contractile function (15, 16). Injury-induced growth factors such as PDGF-BB suppress SMC marker expression, leading to phenotype alteration to synthetic SMC (13, 14, 17). This phenotypic modulation (18) is an essential event in vascular remodeling/neointima formation (19–21).

SMCs and ECs interact under various physiological and pathological conditions (22–25). During vascular development, e.g. vasculogenesis, EC tube formation recruits supporting cells, including SMCs, to form functional blood vessels (26). Conversely, growth-arrested pericytes or SMCs inhibit capillary EC growth in a cell-cell contact-dependent manner in vitro (27). SMCs may also interact with ECs through β-catenin-related pathways and thus impact inflammatory responses of ECs (28). It is unknown, however, whether the proliferating SMCs seen under pathological conditions affect arterial EC proliferation and migration, which are essential for vascular repair following mechanical injury.

Our previous studies have shown that cyclopentenyl cytosine (CPEC), a CTP synthase (CTPS) inhibitor, suppresses neointima formation while promoting re-endothelialization (29). Because CPEC does not induce EC proliferation and migration (29), the underlying mechanisms controlling CPEC-accelerated re-endothelialization remain to be determined. Also, although CPEC affects SMC proliferation, it is unclear whether CPEC affects the SMC phenotype. In this study, we found that PDGF-BB-induced synthetic SMCs exhibit an anti-angiogenic property and thus inhibit EC proliferation/migration. However, CPEC induces SMC redifferentiation to a contractile phenotype that shows a pro-angiogenic property, as evidenced by induction of pro-angiogenic factors and inhibition of anti-angiogenic factors. Of importance, CPEC-induced SMCs stimulate EC proliferation and migration via a pro-angiogenic paracrine effect.

Results

PDGF-BB-induced Synthetic SMC Suppressed EC Proliferation and Migration

PDGF-BB is a potent and known SMC mitogen that induces SMC proliferation and migration (29–31). PDGF-BB treatment also results in a synthetic SMC phenotype, as shown by the reduction of SMC contractile proteins such as smooth muscle myosin heavy chain (SMMHC), smooth muscle α-actin (α-SMA), SM22α, and calponin (CNN1) (Fig. 1, A and B) (32). Because SMC has been shown to affect EC proliferation in a co-culture system (27), we sought to determine whether synthetic SMC has a pro- or anti-angiogenic property. Therefore, we first detected whether synthetic SMC expresses pro-angiogenic or anti-angiogenic factors. As shown in Fig. 1C, PDGF-BB-induced synthetic SMC exhibited decreased expression of several well characterized pro-angiogenic factors, including CCL2 (33), CXCL1 (34), CRY61 (35), G-CSF (36), IGF-1 (37), IL-1β (38), and IL-6 (39), and increased expression of the anti-angiogenic factors endostatin (EST) (40) and TSP1 (41). Accordingly, synthetic SMC culture medium inhibited the proliferation and migration of ECs (Fig. 1, D and F). These data indicate that synthetic SMC exhibits an anti-angiogenic phenotype.

FIGURE 1.

FIGURE 1.

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PDGF-BB-induced synthetic SMC suppressed EC proliferation and migration. A, PDGF-BB (20 ng/ml, 24 h) down-regulated SMC marker protein expression. B, quantification of protein expression shown in A by normalizing to α-tubulin. Ct, control. C, PDGF-BB (20 ng/ml, 12 h) down-regulated the mRNA expression of pro-angiogenic factors and up-regulated the expression of anti-angiogenic factors in cultured SMCs. D, PDGF-BB-conditioned SMC medium suppressed EC proliferation. E, PDGF-BB-conditioned SMC medium suppressed EC migration. Migrating ECs are indicated by arrowheads. F, quantification of EC migration shown in E. Ct-SMCS, control SMC culture medium; BB-SMCS, PDGF-BB-conditioned SMC medium. *, p < 0.05; **, p < 0.01; n = 3.

CPEC Induced Synthetic SMC Redifferentiation into a Contractile SMC Phenotype

Our previous studies showed that CPEC inhibits SMC proliferation (29). Because synthetic SMC displays an anti-angiogenic phenotype, and CPEC promotes re-endothelialization without affecting EC proliferation, we sought to determine whether CPEC alters the SMC phenotype. As shown in Fig. 2A, CPEC induced SMC marker gene expression in both vehicle and PDGF-BB-treated synthetic SMCs. In fact, CPEC induced the expression of SMC contractile proteins such as SMMHC and SM22α in PDGF-BB-treated SMCs in a dose- (Fig. 2, B and C) and time-dependent manner (Fig. 2, D and E), suggesting that CPEC induces the redifferentiation of synthetic SMC to a contractile SMC phenotype. Because contractile SMCs have a spindle-shaped morphology, we tested whether CPEC treatment also alters SMC morphology. As shown in Fig. 2F, CPEC induced SMCs to assume an elongated and spindle-shaped morphology resembling the SMC phenotype observed in the media layer of the artery. To further verify the CPEC-induced SMC differentiation, we tested whether CPEC induces SMC progenitor cells to express SMMHC. As shown in Fig. 2, G and H, CPEC stimulated a dose-dependent expression of SMMHC in human embryonic stem cell-derived mesenchymal stem cells (hMSC). These data indicate that CPEC can induce the contractile phenotype from either synthetic SMC or SMC progenitors.

FIGURE 2.

FIGURE 2.

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CPEC induced redifferentiation of synthetic SMCs to the contractile SMC phenotype. A, CPEC treatment (100 nm, 12 h) induced SMC marker mRNA expression. Ct, vehicle treatment; BB, PDGF-BB treatment (20 ng/ml); CP, CPEC; BB-CP, PDGF-BB with CPEC. B, CPEC induced contractile protein expression in PDGF-BB-treated SMC in a dose-dependent manner. The cells were treated with CPEC for 24 h. C, quantification of protein expression shown in B by normalizing to GAPDH. D, CPEC (1 μm) induced SMC contractile protein expression in a time-dependent manner. E, quantification of protein expression shown in D by normalizing to α-tubulin. F, CPEC (1 μm, 24 h) induced a spindle-shaped contractile SMC morphology. Scale bar = 25 μm. G, CPEC (1 μm, 24 h) induced SMMHC expression in human mesenchymal stem cells. H, quantification of protein expression shown in G by normalizing to α-tubulin. *, p < 0.05; **, p < 0.01; n = 3.

CPEC-conditioned SMC Culture Medium Promoted EC Proliferation and Migration

Because synthetic SMC exhibited an anti-angiogenic effect (Fig. 1, C–F), we sought to determine whether CPEC-induced contractile SMCs have a pro-angiogenic property. Thus, we tested whether CPEC-treated SMCs express pro- or anti-angiogenic factors. As shown in Fig. 3, A and B, although PDGF-BB blocked expression of the pro-angiogenic factors CXCL1, CYR61, and G-CSF and induced the anti-angiogenic factors EST and TSP1, CPEC reversed the effect of PDGF-BB, as shown by stimulating expression of the pro-angiogenic factors while inhibiting expression of the anti-angiogenic factors. Importantly, CPEC-conditioned SMC culture medium promoted EC proliferation and migration (Fig. 3, C–E). These results suggest that CPEC-induced contractile SMCs have a pro-angiogenic property.

FIGURE 3.

FIGURE 3.

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CPEC-conditioned SMC culture medium promoted EC proliferation and migration. A, CPEC (100 nm, 12 h) up-regulated mRNA expression of the pro-angiogenic factors CXCL1, CYR61, and G-CSF in PDGF-BB-treated SMCs. Ct, control; BB, PDGF-BB; BB-CP, PDGF-BB with CPEC. B, CPEC (100 nm, 12 h) down-regulated mRNA expression of the anti-angiogenic factors EST and TSP-1 in PDGF-BB-treated SMCs. C, CPEC-conditioned SMC medium promoted EC proliferation. The cells were cultured in a 24-well plate for 4 days. Ct-SMCS, control SMC medium; CP-SMCS, CPEC-conditioned SMC medium; BB-SMCS, PDGF-BB-conditioned SMC medium; BB-CP-SMCS, both PDGF-BB- and CPEC-conditioned SMC medium. D, CPEC-conditioned SMC medium promoted EC migration. Migrating ECs are indicated by arrowheads. E, quantification of EC migration shown in D. *, p < 0.05; **, p < 0.01; n = 3.

CPEC Induced the Pro-angiogenic Effect of SMCs via EST and CXCL1

Because CXCL1 and EST exhibited the most dramatic changes in SMCs (Figs. 1C and 3, A and B), we tested whether CPEC induces the paracrine angiogenic effects of SMCs via EST or CXCL1. PDGF-BB induced EST and down-regulated CXCL1 protein expression in synthetic SMCs (Fig. 4, A and B), consistent with their mRNA expression (Figs. 1C and 3, A and B). CPEC, however, reversed the effect of PDGF-BB, as shown by the inhibition of EST and the increase in CXCL1 protein expression in PDGF-BB-treated SMCs (Fig. 4, C and D). CPEC also regulated EST and CXCL1 protein expression in a time-dependent manner (Fig. 4, E and F). To determine whether CPEC affects EST and CXCL1 secretion in addition to their expression, we detected the EST and CXCL1 protein levels in SMC culture medium via ELISA. As shown in Fig. 4, G and H, CPEC suppressed EST and increased CXCL1 secretion into SMC culture medium with PDGF-BB treatment.

FIGURE 4.

FIGURE 4.

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CPEC induced a pro-angiogenic paracrine effect of SMCs via EST and CXCL1. A, PDGF-BB (20 ng/ml, 24 h) down-regulated CXCL1 and up-regulated EST protein expression in SMCs. B, quantification of protein expression shown in A by normalizing to α-tubulin. Ct, control. C—F, CPEC up-regulated CXCL1 and down-regulated EST protein expression in PDGF-BB-treated SMC in dose-dependent (C and D, 24-h treatment) and time-dependent (E and F, 100 nm CPEC) manners. The protein expression in C and E was normalized to α-tubulin (D) and GAPDH (F), respectively. G, CPEC (1 μm, 24 h) suppressed EST secretion in SMC culture medium. EST protein levels were normalized to total cellular proteins (ng/ml/1 μg total cellular proteins). H, CPEC (1 μm, 24 h) promoted CXCL1 secretion in SMC culture medium. CXCL1 protein levels were normalized to total cellular proteins (ng/ml/1 μg of total cellular proteins). I and J, recombinant EST (100 ng/ml) or CXCL1-neutralizing antibody (CXCL1nAb, 1 μg/ml)) suppressed EC migration (I) and proliferation (J) induced by CPEC-conditioned SMC medium (BB-CP-SMCS). Ct-SMCS, control SMC medium; BB-SMCS, PDGF-BB-conditioned SMC medium; BB-CP-SMCS, both PDGF-BB- and CPEC-conditioned SMC medium. *, p < 0.05; **, p < 0.01; n = 3.

To test whether CXCL1 and EST play roles in the pro-angiogenic effect of CPEC-induced SMCs on EC proliferation and migration, we added recombinant EST or CXCL1-neutralizing antibody (to block CXCL1 function) in CPEC-conditioned SMC culture medium. As shown in Fig. 4, I and J, recombinant EST (Fig. 4I) or CXCL1-neutralizing antibody (Fig. 4J) effectively suppressed the EC proliferation and migration promoted by the paracrine effects of contractile SMCs that was induced by CPEC, suggesting that CXCL1 and EST mediated the pro-angiogenic effect of the CPEC-induced SMC phenotype.

CPEC Promoted SMC Redifferentiation through Adenosine Receptor (ADOR) Activation

CPEC is an inhibitor of CTPS. However, blockade of CTPS1 expression by its shRNA did not increase the expression of the contractile SMC marker SM22α (Fig. 5, A and B), suggesting that CPEC induced SMC redifferentiation through a CTPS-independent mechanism. In addition to inhibiting CTPS, CPEC can deplete the intercellular CTP pool. Thus, we tested whether CPEC induces SMC redifferentiation by altering the CTP pool. Cytidine feeding has been shown to increase the intercellular CTP concentration (42), so we added cytidine to CPEC-treated SMCs to counter the CPEC effect. Surprisingly, combination treatment of CPEC and cytidine did not attenuate the CPEC effect; rather, it led to higher SM22α expression (Fig. 5, C and D). This increased effect as a result of the addition of cytidine prompted us to hypothesize that CPEC may act as a cytosine derivate to stimulate SMC redifferentiation. In support of this hypothesis, other endogenous cytosine derivatives such as cytosine, cytidine, and CMP, exhibited similar effects in inducing SM22α expression in SMCs (Fig. 5, E and F).

FIGURE 5.

FIGURE 5.

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CPEC prompted SMC redifferentiation by serving as a cytidine derivate mimic. A, CTPS1 knockdown by shRNA did not increase SMC marker SM22α expression. B, quantification of protein expression shown in A by normalizing to α-tubulin level. C, cytidine (1 μm, 24 h) significantly increased CPEC-induced SMC contractile protein SM22α expression. Ct, control; NS, non-serum. D, quantification of protein expression shown in C by normalizing to α-tubulin. E, cytosine derivatives (1 μm each, 24 h) induced SM22α expression. F, quantification of protein expression shown in E by normalizing to α-tubulin. *, p < 0.05; **, p < 0.01; n = 3.

Previous studies have shown that adenosine receptors can function as receptors for nucleotide mimics (43). CPEC up-regulated the expression of ADORA1 and ADORA2a in both control and PDGF-BB-treated SMCs (Fig. 6A). CPEC also enhanced ADORA2a protein expression (Fig. 6, B and C). Moreover, the ADORA downstream signal was also activated by CPEC treatment, as evidenced by the increased intercellular cAMP level (Fig. 6D) and the phosphorylation of cAMP response element-binding protein (CREB) (Fig. 6, E and F). Importantly, both the ADORA1 inhibitor CPDX and the ADORA2a inhibitor KW-6002 suppressed CPEC-induced SMC redifferentiation (Fig. 6, G and H), indicating that CPEC regulates SMC redifferentiation through ADORA signaling. Notably, KW-6002 displayed a more potent effect than CPDX (Fig. 6, G and H), suggesting that ADORA2a may share more responsibility than ADORA1 in mediating CPEC function.

FIGURE 6.

FIGURE 6.

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CPEC triggered SMC redifferentiation by activating ADORA signaling. A, CPEC (100 nm, 12 h) induced ADORA1 and ADORA2α mRNA expression. B, CPEC (1 μm, 24 h) induced ADORA2a protein expression. C, quantification of protein expression shown in B by normalizing to GAPDH. D, CPEC (1 μm, 15 min) increased the intercellular cAMP level (nanograms/10E7 cells) in SMCs. E, CPEC (1 μm) induced CREB phosphorylation in SMCs. F, quantification of protein expression shown in E by normalizing to GAPDH. G, the ADORA1 inhibitor CPDX (10 nm) and ADORA2a inhibitor KW-6002 (10 nm) suppressed CPEC-induced (1 μm, 24 h) SMC marker expression. H, quantification of protein expression shown in G by normalizing to α-tubulin. Ct, control (vehicle treatment); CP, CPEC treatment; CP+DX, CPEC with CPDX; CP+DW, CPEC with KW-6002. *, p < 0.05; **, p < 0.01; n = 3.

CPEC Triggered SMC Redifferentiation through ADOR Downstream Akt Signaling

It is well established that Smad3 activation is critical for contractile protein expression and SMC differentiation (44). However, CPEC did not enhance Smad3 expression or phosphorylation (Fig. 7, A and B), suggesting that CPEC induces SMC redifferentiation through a Smad-independent mechanism. Indeed, CPEC induced Akt phosphorylation in a dose-dependent manner in both control and PDGF-BB-treated SMCs (Fig. 7, A and B). CPDX or KW-6002 suppressed CPEC-induced Akt phosphorylation (Fig. 7, C and D), suggesting that CPEC activates Akt signaling via ADORA1 and ADORA2a. Because Akt signaling is involved in CREB activation (45), which further regulates SMC differentiation (46), we tested whether CPEC induced SMC redifferentiation through the ADORA-Akt-CREB axis. Thus, we tested whether PI3/Akt signaling mediated CPEC-induced CREB phosphorylation. As shown in Fig. 7, E and F, the blockade of PI3K/Akt signaling by its inhibitor LY-294002 attenuated CPEC-enhanced CREB phosphorylation as well as SMC marker SM22α expression (Fig. 7, G and H), indicating that CPEC induced SMC redifferentiation through the ADORA-Akt-CREB axis.

FIGURE 7.

FIGURE 7.

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CPEC induced SMC redifferentiation through the ADORA-Akt-CREB axis. A, CPEC induced Akt phosphorylation in a dose-dependent manner. Note that Smad3 phosphorylation was suppressed in a high concentration of CPEC (1 μm), probably because of inhibition of Smad3 protein expression. Ct, control. B, quantification of protein expression shown in A by normalizing to the GAPDH level. BB, PDGF-BB; CP, CPEC. C, both ADORA1 (CPDX) and ADORA2a (KW-6002) inhibitors suppressed CPEC-induced (1 μm) Akt phosphorylation. D, quantification of Akt phosphorylation shown in C by normalizing to the total Akt level. CP+DX, CPEC with CPDX; CP+KW, CPEC with KW-6002. E, the PI3K/Akt pathway inhibitor LY294002 (10 nm) suppressed CPEC-induced (1 μm) CREB phosphorylation. F, quantification of CREB phosphorylation shown in E by normalizing to the total CREB level. G, the PI3K/Akt pathway inhibitor LY294002 (10 nm) suppressed SM22α protein expression enhanced by CPEC (1 μm). H, quantification of protein expression shown in G by normalized to α-tubulin level. BB+CP, PDGF-BB with CPEC; BB+CP+LY, treatment with PDGF-BB, CPEC, and LY294002. **, p < 0.01; n = 3.

CPEC Induced Neointimal SMC Redifferentiation, Inhibited EST and Enhanced CXCL1 Expression, and Promoted Re-endothelialization in Vivo

CPEC did not promote EC proliferation and migration in vitro (29), but CPEC promoted EC proliferation/migration via the paracrine effect of CPEC-induced contractile SMCs. To test whether CPEC-induced contractile SMCs produce angiogenic factors under pathological conditions, we used rat carotid artery balloon injury model to mimic vascular injury in vivo and use an osmotic pump to infuse saline or CPEC into rats undergoing the artery injury. As shown in Fig. 8, A and B, infusion of CPEC significantly attenuated neointima formation and promoted redifferentiation of the neointimal SMCs, as indicated by expression of the SMC marker SM22α compared with the saline-treated artery. Importantly, neointimal SMCs in CPEC-treated arteries showed reduced expression of the anti-angiogenic factor EST and a significant increase in pro-angiogenic CXCL1 (Fig. 8, C–F). Consequently, CPEC treatment promoted re-endothelialization, as demonstrated by CD31 staining (Fig. 8, G and H), consistent with our previous finding (29). These data suggest that CPEC induces a contractile/pro-angiogenic phenotype in neointimal SMCs, which promotes EC proliferation/migration, resulting in accelerated re-endothelialization.

FIGURE 8.

FIGURE 8.

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CPEC promoted re-endothelialization through inducing neointima SMC redifferentiation in vivo. Rat left carotid arteries were injured with a 2F Fogarty arterial embolectomy balloon catheter. 1 mg/kg/day CPEC was infused into the rat through an osmotic minipump that was implanted on the day of artery injury. 14 days later, the arteries were removed, and immunohistochemistry staining with the indicated antibodies was performed. A and B, CPEC significantly increased contractile SMC marker SM22α expression in neointimal SMCs. C and D, CPEC suppressed expression of the anti-angiogenic factor EST in neointimal SMCs. E and F, CPEC induced expression of pro-angiogenic factor CXCL1 expression in neointimal SMCs. G and H, CPEC accelerated re-endothelialization. The endothelium was stained by CD31. The expression of SM22α, EST, and CXCL1 was quantified by measuring mean intensity per area in 10 different fields and is shown as -fold changes. The re-endothelization was quantified by averaging the CD31-positive cells in 10 different fields and is shown as -fold changes. *, p < 0.05; **, p < 0.01; n = 10.

Discussion

EC proliferation and migration are key events during vascular repair following injury. The mechanisms underlying re-endothelialization are thought to be attributable primarily to the intrinsic factors or signaling of ECs. This study indicates that SMC phenotypes play critical roles in EC properties. Proliferative or synthetic SMCs inhibit EC proliferation and migration, whereas CPEC-induced contractile SMCs stimulate EC proliferation/migration, which is due to the production of pro-angiogenic factors and the blockade of anti-angiogenic factors within the redifferentiated SMCs. Therefore, contractile SMCs exhibit a pro-angiogenic phenotype, whereas synthetic SMCs display an anti-angiogenic phenotype. This concept is also supported by a previous study showing that serum-treated SMCs suppress EC replication, although the mechanism has not been determined (47). It is likely that serum-treated SMCs act similarly as PDGF-BB-treated SMCs, i.e. producing anti-angiogenic factors to block EC proliferation.

The pro-angiogenic function of CPEC-induced SMCs is likely to be a unique property of contractile SMCs converted from neointima/proliferative SMCs during vascular repair following pathological remodeling. Thus, it may not be relevant to physiological angiogenesis because mature SMCs inhibits excessive EC proliferation and migration during vascular development (27). Accordingly, CPEC-induced SMCs may not be identical to mature SMCs in the blood vessel that is formed during the vascular development, although CPEC can also induce SMC differentiation from mesenchymal progenitors.

Interestingly, although CPEC is an inhibitor of CTPS, it does not cause SMC phenotypic alteration by its function in CTPS activity or depletion of the CTP pool. Instead, CPEC plays a new role by acting as a cytosine derivate to induce SMC redifferentiation. Importantly, other cytosine derivatives can also induce SMC redifferentiation, although their effects are much less compared with CPEC (Fig. 5E). Combined treatment of CPEC and cytidine significantly increases SMC marker gene expression, suggesting that CPEC and other cytosine derivatives have a synergistic effect in inducing SMC redifferentiation. Collectively, it appears that CPEC promotes vascular repair through two independent mechanisms, i.e. blocking SMC proliferation via inhibition of CTPS activity and prompting a contractile/pro-angiogenic SMC phenotype by acting as a cytosine derivate. Both functions appear to be critical for promoting re-endothelialization and vascular repair.

CPEC induces SMC redifferentiation via the ADOR-PI3K/Akt-CREB axis. ADOR is a class of G protein-coupled purinergic receptors using free nucleotides (mainly adenosine and uridine) as ligands (48). SMCs express ADORs. Functionally, these receptors are involved in cell contraction and ion channel activities related to cAMP signaling (49–51). Our results demonstrate for the first time that ADORA1 and ADORA2a may also serve as receptors for CPEC, which activates both cAMP and Akt; cAMP and Akt further promote CREB phosphorylation, leading to SMC redifferentiation. The results showing that PI3K/Akt inhibitor blocks CPEC-induced SMC redifferentiation are consistent with previous findings that the PI3K/Akt pathway is essential for maintaining the differentiated SMC phenotype (52) and that cAMP-mediated Akt signaling is necessary for SMC contraction (49–51). Therefore, PI3K/Akt signaling may regulate the SMC phenotype via both CREB-dependent and CREB-independent mechanisms.

In summary, we have demonstrated that PDGF-BB-induced synthetic SMC exhibits an anti-angiogenic phenotype inhibiting EC proliferation/migration, whereas CPEC-induced contractile SMCs shows a pro-angiogenic phenotype promoting EC proliferation/migration. CPEC induces SMC redifferentiation through activating the ADORA-Akt-CREB cascade (Fig. 8).

Experimental Procedures

Reagents and Cell Culture

Rat aortic SMCs and ECs were cultured by the enzyme digestion method from rat thoracic aortae as described previously (53, 54). SMCs were maintained at 37 °C in a humidified 5% CO2 incubator in DMEM containing 10% FBS, 4.5 g/liter glucose, 4.5 g/liter sodium pyruvate, 2 mm l-glutamine, 100 units/ml penicillin, and 100 μg/ml streptomycin. Primary ECs were maintained at 37 °C in a humidified 5% CO2 incubator in DMEM containing 20% fetal bovine serum, 100 units/ml penicillin, 100 μg/ml streptomycin, 2 mm l-glutamine, 10 mm non-essential amino acids, 1 mm sodium pyruvate, 25 mm HEPES, 100 μg/ml heparin, 100 μg/ml endothelial cell growth supplement. Cells within passage 3 were used for all experiments in this study. The SMC and EC phenotypes of the cultured cells were confirmed by expression of smooth muscle α-actin and CD31, respectively. hMSCs were obtained from ArunA Biomedical (Athens, GA). hMSCs were cultured in α-minimal essential medium (Cellgro, Fisher Scientific, Pittsburgh, PA) with 10% mesenchymal stem cell-qualified fetal bovine serum (Hyclone) and 2 mm l-glutamine (Hyclone).

SMCs were cultured in complete medium with or without treatment of CPEC and/or PDGF-BB. 48 h after the treatment, 3 × 106 cells were reseeded in 6-cm culture dishes. After the SMC monolayer was formed, cells were washed with warm PBS three times and incubated with serum-free DMEM for 12 h, followed by collection of conditioned medium that was filtered through a 0.22-μm filter, and stored at −80 °C.

CPEC (compound 375575) was obtained from the Open Chemical Repository of the NCI, National Institutes of Health Developmental Therapeutics Program (DTP). Cytosine, cytidine, CMP, CDP, CTP, CPDX, and KW6002 were purchased from Sigma-Aldrich (St. Louis, MO). PDGF-BB was purchased from R&D Systems (Minneapolis, MN).

SM22α (ab10135), CNN1 (ab46794), and CXCL1 (ab86436) antibodies were purchased from Abcam (Cambridge, MA). Smad3 (9523S), p-Smad3 (9520S), Akt (4691S), and pAkt (9271S) antibodies were purchased from Cell Signaling Technology (Danvers, MA). EST (abc60) antibody was purchased from EMD Millipore (Billerica, MA). GAPDH (G8795) antibody was purchased from Sigma-Aldrich. EST proteins and CXCL1-neutralizing antibody were purchased from R&D Systems.

Animals

Male Sprague-Dawley rats weighing 450–500 g were purchased from Harlan. All animals were housed under conventional conditions in the animal care facilities and received humane care in compliance with the Principles of Laboratory Animal Care formulated by the National Society for Medical Research and the Guide for the Care and Use of Laboratory Animals. Animal surgical procedures were approved by the Institutional Animal Care and Use Committee of the University of Georgia.

Rat Carotid Artery Injury Model and Immunohistochemistry Staining

Rat carotid artery balloon injury was performed using a 2F Fogarty arterial embolectomy balloon catheter (Baxter Edwards Healthcare) as described previously (53). 14 days later, the balloon-injured arteries were perfused with saline, fixed with 4% paraformaldehyde, embedded in paraffin, and sectioned. For immunohistochemistry staining, sections were rehydrated, permeabilized with 0.01% Triton X-100 in PBS, blocked with 10% goat serum, and incubated with primary antibodies overnight at 4 °C, followed by incubation with HRP-conjugated secondary antibody. The sections were counterstained with hematoxylin.

Quantitative RT-PCR (qPCR)

Total RNA was extracted from primary cultured SMCs using TRIzol reagent (Invitrogen) and reverse-transcribed to cDNA using the iScriptTM cDNA synthesis kit (Bio-Rad). qPCR was performed on a Stratagene Mx3005 qPCR thermocycler (Agilent Technologies, La Jolla, CA) as described previously (55). Primer sequences for qPCR are listed in supplemental Table S1. Cyclophilin was used as an internal control. The primer efficiency was verified by the dissociation curve in qPCR reactions. The -fold change for each target gene was calculated as 2−ΔΔCt.

Western Blotting

Western blotting was performed as described previously (55).

EC Proliferation Assay

ECs were seeded in 24-well plates at 1 × 104/well in EC complete medium. After 24 h, the medium was changed to DMEM containing 20% SMC-conditioned medium, 2% FBS, and 2 mm l-glutamine and cultured for another 4 days. Fresh medium was changed every other day, and cell numbers were counted every day.

Transwell Assay

The transwell assay was carried out according to the instructions of the manufacturer (Corning Inc., Corning, NY). Primary ECs were seeded onto the transwell upper chamber inserts (1 × 105/insert). The inserts were then put back into the receiver plate filled with 80% endothelial culture medium plus 20% SMC-conditioned medium and incubated at 37 °C in a humidified 5% CO2 incubator for 12 h. After incubation, the transwell inserts were washed with PBS three times and fixed with 4% paraformaldehyde for 10 min at room temperature, followed by washing with PBS. Cells were stained with Wright-Giemsa for acquiring images or DAPI for cell counting. Images of migrating cells were captured using a dissection microscope (Olympus).

ELISA

CXCL1 and EST levels in SMC-conditioned medium were measured with commercial rat CXCL1 (MBS824537) and Endostatin ELISA kits (MBS730385) from MyBioSource Inc., respectively. SMC-conditioned medium was collected, filtered through a 0.22-μm filter, and stored at −80 °C for later use. Purified rat CXCL1 and EST were serially diluted and served as standards. Intercellular cAMP levels were measured with a commercial cAMP ELISA kit (EIA-CAMP-1) from RayBiotech, Inc. ELISA assays were performed according to the protocol of the manufacturer.

Statistical Analysis

Each experiment was repeated at least three times. All values are presented as mean ± S.E. Comparisons of parameters among groups were made by one-way analysis of variance, and comparisons of different parameters between each group were made by a post hoc analysis using a Bonferroni test. p < 0.05 was considered statistically significant.

Author Contributions

R. T. and S. Y. C. conceived and coordinated the study. R. T. designed and performed the experiments shown in Figs. 1–8. G. Z. provided technical assistance and contributed to the experiments shown in Figs. 2 and 4–6. R. T. and S. Y. C. analyzed the results and wrote the paper. All authors analyzed the results and approved the final version of the manuscript.

Supplementary Material

Supplemental Data
supp_291_52_26913__index.html (1,019B, html)

Acknowledgments

We thank the Developmental Therapeutics Program of the NCI, National Institutes of Health for providing CPEC.

*

This work was supported, in whole or in part, by National Institute of Health Grants HL123302 and HL119053. The authors declare that they have no conflicts of interest with the contents of this article. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

Inline graphic

This article contains supplemental Table S1.

2
The abbreviations used are:
EC
endothelial cell
SMC
smooth muscle cell
CPEC
cyclopentenyl cytosine
SMMHC
smooth muscle myosin heavy chain
EST
endostatin
hMSC
human embryonic stem cell-derived mesenchymal stem cell
ADOR
adenosine receptor
CREB
cAMP response element-binding protein; 1,3-dipropyl-8-phenylxanthine
qPCR
quantitative RT-PCR.

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