p115 RhoGEF activates the Rac1 GTPase signaling cascade in MCP1 chemokine–induced vascular smooth muscle cell migration and proliferation

Abstract

Although the involvement of Rho proteins in the pathogenesis of vascular diseases is well studied, little is known about the role of their upstream regulators, the Rho guanine nucleotide exchange factors (RhoGEFs). Here, we sought to identify the RhoGEFs involved in monocyte chemotactic protein 1 (MCP1)–induced vascular wall remodeling. We found that, among the RhoGEFs tested, MCP1 induced tyrosine phosphorylation of p115 RhoGEF but not of PDZ RhoGEF or leukemia-associated RhoGEF in human aortic smooth muscle cells (HASMCs). Moreover, p115 RhoGEF inhibition suppressed MCP1-induced HASMC migration and proliferation. Consistent with these observations, balloon injury (BI) induced p115 RhoGEF tyrosine phosphorylation in rat common carotid arteries, and siRNA-mediated down-regulation of its levels substantially attenuated BI-induced smooth muscle cell migration and proliferation, resulting in reduced neointima formation. Furthermore, depletion of p115 RhoGEF levels also abrogated MCP1- or BI-induced Rac1–NFATc1–cyclin D1–CDK6–PKN1–CDK4–PAK1 signaling, which, as we reported previously, is involved in vascular wall remodeling. Our findings also show that protein kinase N1 (PKN1) downstream of Rac1–cyclin D1/CDK6 and upstream of CDK4–PAK1 in the p115 RhoGEF–Rac1–NFATc1–cyclin D1–CDK6–PKN1–CDK4–PAK1 signaling axis is involved in the modulation of vascular wall remodeling. Of note, we also observed that CCR2–G_i/o–Fyn signaling mediates MCP1-induced p115 RhoGEF and Rac1 GTPase activation. These findings suggest that p115 RhoGEF is critical for MCP1-induced HASMC migration and proliferation in vitro and for injury-induced neointima formation in vivo by modulating Rac1–NFATc1–cyclin D1–CDK6–PKN1–CDK4–PAK1 signaling.

Introduction

Migration and proliferation of vascular smooth muscle cells (VSMCs)³ occur during vascular development, in response to vascular injury, and during atherogenesis (1, 2). Various molecules, including cytokines, chemokines, and reactive oxygen species, are produced at the site of vascular injury, and they have been demonstrated to be involved in vascular wall remodeling (3–5). Among these various molecules, monocyte chemotactic protein 1 (MCP1) is one of the prominent chemokine produced at the site of vascular injury that possesses both chemotactic and mitogenic activity toward VSMCs (4, 5). A large body of evidence suggests that MCP1 plays a crucial role in the recruitment of monocytes, memory T cells, and dendritic cells to the sites of tissue injury, suggesting its importance in cell migration (6, 7). On the other hand, its role in cell proliferation appears to be controversial, as a few studies showed that it negatively modulates VSMC replication (8), whereas other reports, including our own, reveal that MCP1 enhances both VSMC migration and proliferation in the development of vascular wall remodeling (5, 9–11).

The Rho family of proteins (Rho, Rac, and CDC42) plays an important role in the regulation of various cellular functions, including cell migration and proliferation (12). Many studies have also reported their role in various vascular pathologies both in animals and humans (13–15). The activation of these Rho family of proteins is mediated by Rho guanine exchange factors (RhoGEFs) that promote the release of GDP in exchange for GTP (16). Among the 70 RhoGEFs known, the LARG, PDZ RhoGEF, and p115 RhoGEF appear to be involved in the activation of Rho proteins in the modulation of vascular tone, particularly hypertension (17–19). Despite these reports, very little is known about the role of these RhoGEFs in vascular wall remodeling. The work from our laboratory shows that Rac1 plays a role in MCP1-induced VSMC migration, proliferation, and injury-induced neointimal development. The purpose of this study was to identify the RhoGEF that is involved in Rac1 activation and mediating MCP1-induced vascular wall remodeling. Here we report that p115 RhoGEF mediates MCP1 and injury-induced Rac1 activation, enhancing VSMC migration and proliferation and, thereby, vascular wall remodeling. We also found that p115 RhoGEF, via Rac1, modulates the NFATc1–cyclin D1/CDK6–PKN1–CDK4–PAK1 signaling axis in MCP1/injury-induced vascular wall remodeling.

Results

We have shown previously that MCP1 is produced at sites of vascular injury and is involved in vascular wall remodeling (5). In addition, we have reported that Rac1 mediates NFATc1–cyclinD1/CDK6–CDK4–PAK1 signaling in the regulation of MCP1-induced VSMC migration and proliferation (20). However, we do not know how Rac1 is activated by MCP1 or injury. Because we have shown previously that p115 RhoGEF plays a role in Rac1 activation in response to thrombin in mediating HASMC proliferation and migration (21), we studied the role of regulator of G protein signaling RhoGEFs. We found that MCP1 induces tyrosine phosphorylation of p115 RhoGEF but not PDZ RhoGEF or LARG in HASMCs, with a maximum effect at 10 min (Fig. 1A). In addition, down-regulation of p115 RhoGEF levels using its siRNA attenuated MCP1-induced HASMC migration and proliferation (Fig. 1, B–D).

Figure 1.

p115 RhoGEF mediates MCP1-induced HASMC migration and proliferation. A, top panel, quiescent HASMCs were treated with MCP1 (50 ng/ml) for the indicated time periods or left untreated, and cell extracts were prepared. An equal amount of protein from the control and each treatment was immunoprecipitated (IP) with phosphotyrosine (PY20) antibodies, and the immunocomplexes were immunoblotted (IB) with anti-p115 RhoGEF, anti-PDZ RhoGEF, or anti-LARG antibodies. An equal amount of protein from the same cell extracts was analyzed by Western blotting for total levels using specific antibodies. Bottom panel, equal amounts of protein from the control and each treatment were also immunoprecipitated with anti-p115 RhoGEF antibodies. The immunocomplexes were immunoblotted with PY20 antibodies, and the blot was reprobed for p115 RhoGEF for normalization. B, HASMCs were transfected with control or p115 RhoGEF siRNA (100 nm) and quiesced. Cell extracts were prepared, an equal amount of protein from each condition was analyzed by Western blotting for p115 RhoGEF levels using specific antibody, and the blot was reprobed for β-tubulin to show the effect of siRNA on its target and off-target molecules. C and D, HASMCs that were transfected with control or p115 RhoGEF siRNA and quiesced were treated with MCP1 (50 ng/ml) for 24 h or left untreated, and cell migration and DNA synthesis were measured by wound healing and [³H]thymidine incorporation assays, respectively. The bar graphs represent the mean ± S.D. values of three independent experiments. *, p < 0.01 versus control or siControl; **, p < 0.01 versus MCP1 or siControl + MCP1.

To find out whether arterial injury also stimulates p115 RhoGEF, and if so, the potential role of p115 RhoGEF in vascular wall remodeling, we used a rat carotid artery restenosis model. BI induced tyrosine phosphorylation of p115 RhoGEF, with maximum effect 5 days after injury (Fig. 2A). Migration of VSMCs from media to intima is a pathological manifestation of restenosis after angioplasty (3). To understand the role of p115 RhoGEF in VSMC migration from media to intima in vivo, we used an siRNA approach. Pluronic gel (30%, v/v)–mediated delivery of p115 RhoGEF siRNA (10 μg) into the injured artery depleted p115 RhoGEF levels substantially 3 days after transfection (Fig. 2B). Depletion of p115 RhoGEF levels by its siRNA decreased VSMC migration, as measured by a decrease in the number of SMCα actin–positive cells on the luminal surface of the injured artery compared with siControl (Fig. 2B). The assumption is that the migrated VSMCs in the intimal region proliferate, forming neointima following angioplasty or vein grafting (2, 3). So, to study the role of p115 RhoGEF in injury-induced VSMC proliferation, the arteries were isolated 7 days after injury and fixed. Cross sections were made and co-stained for SMCα-actin and Ki67 (a cellular marker for proliferation). Down-regulation of p115 RhoGEF levels led to a significant decrease in SMCα–actin/Ki67–positive cells in the intimal region of the injured arteries compared with siControl (Fig. 2C). Consistent with these observations, depletion of p115 RhoGEF levels also reduced injury-induced neointima formation, as measured by morphometric analysis (Fig. 2D).

Figure 2.

Down-regulation of p115 RhoGEF levels inhibits BI-induced SMC migration, proliferation, and neointima formation. A, common carotid arteries from the control or the indicated time periods of post-BI rats were dissected out, and tissue extracts were prepared. The tissue extracts containing an equal amount of protein from each group of pooled arteries were immunoprecipitated (IP) with Tyr(P)-20 antibodies, and the immunocomplexes were immunoblotted (IB) for p115 RhoGEF. An equal amount of protein from each sample was analyzed by Western blotting for total p115 RhoGEF or β-tubulin levels using specific antibodies. B, left panel, arteries from 3 days post-BI rats that were administered 10 μg of control or p115 RhoGEF siRNA were isolated. Tissue extracts were prepared, equal amounts of protein from each were analyzed by Western blotting for p115 RhoGEF levels, and the blot was reprobed for β-tubulin to show the effect of siRNA on its target and off-target molecules. Center panel, all conditions were the same as in the left panel, except that arteries were opened longitudinally and stained for SMCα actin. Right panel, the bar graph represents the quantitative analysis of SMC migration from six animals (mean ± S.D.). C, arteries from 1 week post-BI rats that were administered 10 μg of control or p115 RhoGEF siRNA were isolated and fixed. Cryosections were made and co-immunostained for Ki67 and SMCα-actin using specific antibodies. The bar graph shows the quantitative analysis of neointimal SMC proliferation from six animals (mean ± S.D.). D, arteries from 2 weeks post-BI rats that were administered 10 μg of control or p115 RhoGEF siRNA were isolated and fixed. Cross-sections were made and stained with H&E, and the I/M ratios were calculated. Left panel, representative pictures of BI common carotid artery cross-sections that were stained with H&E. Right panel, the bar graph shows mean ± S.D. values of the I/M ratios of the injured common carotid arteries from six animals. *, p < 0.05 versus uninjured or siControl + BI.

Previous studies from our laboratory have shown that NFATc1 mediates VSMC migration and proliferation in response to various agonists, including MCP1 (20, 22, 23). In addition, we have shown that Rac1 mediates MCP1-induced NFATc1 activation (20). Based on this information as well as the fact that p115 RhoGEF mediates activation of the Rho family of small GTPases (24, 25), we tested the role of p115 RhoGEF in MCP1-induced Rac1 and NFATc1 activation. Consistent with our previous observations, MCP1 induced Rac1 activation in a time-dependent manner in HASMCs, and depletion of p115 RhoGEF levels abrogated this effect (Fig. 3, A and B). This result infers that MCP1 activates Rac1 via p115 RhoGEF. Depletion of p115 RhoGEF levels also decreased NFATc1 translocation from the cytoplasm to the nucleus, suggesting the involvement of p115 RhoGEF in MCP1-induced NFATc1 activation as well (Fig. 4A). We have also shown previously that NFATc1, via enhancing cyclin D1 expression, regulates CDK6/CDK4-mediated PAK1 activation and CDK6-mediated PKN1 activation (20, 22). We have also shown a role for both PAK1 and PKN1 in VSMC migration, proliferation, and injury-induced neointima formation (20, 22). To understand the mechanisms by which p115 RhoGEF regulates MCP1-induced HASMC migration and proliferation, we tested its role in MCP1-induced cyclinD1 expression, CDK6 and CDK4 activities, and PKN1 and PAK1 phosphorylation. Down-regulation of p115 RhoGEF levels blocked MCP1-induced cyclinD1 expression, CDK6 and CDK4 activities, as well as PKN1 and PAK1 phosphorylation (Fig. 4B). In line with these observations, adenovirus-mediated expression of a dominant negative mutant of Rac1 blocked MCP1-induced cyclin D1 expression, CDK6 and CDK4 activities, and PKN1 and PAK1 phosphorylation (Fig. 4C). Furthermore, down-regulation of NFATc1 or cyclin D1 levels using their siRNAs inhibited MCP1-induced CDK6 and CDK4 activities and PKN1 and PAK1 phosphorylation (Fig. 4D). These observations indicate that p115 RhoGEF, via enhancing Rac1-dependent NFATc1-mediated cyclin D1 expression, regulates MCP1-induced CDK6/CDK4 activities and PKN1 and PAK1 phosphorylation. We have shown previously that CDK6 acts upstream of CDK4 in mediating MCP1-induced PAK1 activation (20). In addition, we have reported that the MCP1-induced NFATc1–cyclin D1–CDK6 axis regulates PKN1 activation (22). Because both PKN1 and PAK1 are activated by MCP1-induced p115 RhoGEF–Rac1–NFATc1–cyclinD1 signaling, we further wanted to test whether both PKN1 and PAK1 are acting independently or regulating each other in MCP1-induced HASMC migration or proliferation. Down-regulation of CDK6, PKN1, or CDK4 by their respective siRNAs attenuated MCP1-induced CDK4 activity and PAK1 phosphorylation (Fig. 5A). On the other hand, depletion of CDK6, but not CDK4, blocked MCP1-induced PKN1 phosphorylation (Fig. 5A), suggesting that PAK1 is downstream of CDK6, PKN1, and CDK4 in mediating MCP1-induced VSMC migration and proliferation. On the other hand, down-regulation of CDK6 but not CDK4 and PAK1 levels using their siRNAs attenuated MCP1-induced PKN1 phosphorylation, indicating that PKN1 acts downstream of CDK6 and upstream of CDK4 and PAK1 in MCP1-induced VSMC migration and proliferation (Fig. 5B). To test whether BI-induced p115 RhoGEF modulates VSMC migration and proliferation via Rac1–NFATc1–cyclin D1–CDK6–PKN1–CDK4-PAK1 signaling, we further studied the role of p115 RhoGEF in injury-induced activation of this signaling axis. Depletion of p115 RhoGEF levels via its siRNA in the arteries not only blocked BI-induced Rac1 activation, cyclin D1 expression, and CDK6 and CDK4 activities but also inhibited PKN1 and PAK1 phosphorylation (Fig. 5C). These results infer that p115 RhoGEF mediates Rac1–NFATc1–cyclin D1–CDK6–PKN1–CDK4–PAK1 signaling activation in response to BI in vivo in arteries, leading to enhanced SMC migration from the media to the intima and their proliferation in intima leading to neointima formation.

Figure 3.

p115 RhoGEF mediates MCP1-induced Rac1 activation in HASMCs. A, quiescent HASMCs were treated with MCP1 (50 ng/ml) for the indicated time periods or left untreated, and cell extracts were prepared. Equal amounts of protein from the control and each treatment were analyzed by pulldown assay for Rac1 and RhoA activation using GST-PAK1 and GST-Rhotekin beads, respectively. Equal amounts of protein from the same cell extracts were analyzed by Western blotting for total Rac1 and RhoA levels using specific antibodies. B, HASMCs were transfected with control or p115 RhoGEF siRNA (100 nm), quiesced, and treated with MCP1 (50 ng/ml) for 2 h or left untreated. Cell extracts were prepared and analyzed by pulldown assay for Rac1 using GST-PAK1 beads as described in A. Equal amounts of protein from the same cell extracts were also analyzed by Western blotting for total Rac1 levels. The bar graphs represent the mean ± S.D. values of three independent experiments. *, p < 0.01 versus control or siControl; **, p < 0.01 versus MCP1 or siControl + MCP1.

Figure 4.

p115 RhoGEF mediates MCP1-induced cyclin D1 expression, CDK4/6 activities, and PKN1 and PAK1 phosphorylation. A, HASMCs were transfected with control or p115 RhoGEF siRNA, quiesced, treated with and without MCP1 (50 ng/ml) for 30 min, fixed, permeabilized, and immunostained for NFATc1 using its specific antibody, followed by probing with Alexa Fluor 488-conjugated secondary antibody. The images were captured under a Zeiss Oberver.Z1 fluorescence microscope using an AxioCam MRm camera. B and D, HASMCs were transfected with control, p115 RhoGEF, NFATc1, or cyclin D1 siRNA (100 nm), quiesced, and treated with MCP1 (50 ng/ml) for 8 h or left untreated. Cell extracts were prepared and analyzed by Western blotting for cyclin D1, pPKN1, and pPAK1 levels using their specific antibodies or by immunocomplex kinase assay for CDK4/6 activities using truncated recombinant retinoblastoma protein and [γ-³²P]ATP as substrates. C, HASMCs were transduced with Ad-GFP or Ad-dnRac1 (10⁹ pfu), quiesced, and treated with MCP1 (50 ng/ml) for 8 h or left untreated. Cell extracts were prepared and analyzed for cyclin D1, pPKN1, and pPAK1 levels or CDK4/6 activities as described in B. The blots in B–D were reprobed for PKN1, PAK1, p115 RhoGEF, NFATc1, cyclin D1, or β-tubulin antibodies for normalization or to show the effect of a specific siRNA on its target and off-target molecules. The bar graphs represent the mean ± S.D. values of three independent experiments. *, p < 0.01 versus siControl; **, p < 0.01 versus siControl + MCP1.

Figure 5.

PKN1 mediates CDK4-dependent PAK1 phosphorylation. A and B, HASMCs were transfected with control, CDK6, PKN1, CDK4, or PAK1 siRNA (100 nm), quiesced, and treated with MCP1 (50 ng/ml) for 8 h or left untreated. Cell extracts were prepared and analyzed by Western blotting for pPKN1 and pPAK1 levels using their specific antibodies or by immunocomplex kinase assay for CDK4/6 activities. Recombinant retinoblastoma protein was used as a substrate for CDK4/6 activities. C, arteries from uninjured or 5 days post-BI rats that were administered 10 μg of control or p115 RhoGEF siRNA were isolated. Tissue extracts were prepared and analyzed by Western blotting for cyclin D1, pPKN1, and pPAK1 levels or by immunocomplex kinase assay for CDK6/4 activities. The tissue extracts were also analyzed by pulldown assays for Rac1 using GST-PAK1 beads. Equal amounts of protein from the same tissue extracts were analyzed by Western blotting for total Rac1 levels. The blots in A–C were reprobed for CDK6, CDK4, PKN1, PAK1, p115 RhoGEF, or β-tubulin levels for normalization or to show the effect of a specific siRNA on its target and off-target molecules.

We have reported previously that MCP1-induced HASMC proliferation requires CCR2-dependent, G_i/o-mediated Fyn activation (26). Based on these observations, we asked whether CCR2, G_i/o, and Fyn signaling plays a role in MCP1-induced p115 RhoGEF tyrosine phosphorylation. Inhibition of CCR2 by its antagonist CCR2A or G_i/o by pertussis toxin or siRNA-mediated depletion of Fyn blocked MCP1-induced p115 RhoGEF tyrosine phosphorylation and Rac1 activation (Fig. 6, A and B). These results suggest that activation of CCR2–G_i/o–Fyn signaling is required for MCP1-induced p115 RhoGEF tyrosine phosphorylation and Rac1 activation. Many reports have demonstrated that tyrosine phosphorylation of RhoGEFs, including LARG and p115 RhoGEF, is sufficient for activation of their GEF activity (27, 28). Therefore, to find out whether tyrosine phosphorylation of p115 RhoGEF is required for its GEF activity, we performed RhoGEF exchange assay using Rac1 and RhoA as substrates. MCP1 induced p115 RhoGEF exchange activity more robustly with Rac1 than RhoA, and inhibition of CCR2 or G_i/o or depletion of Fyn levels substantially reduced its activity (Fig. 6, C and D). These findings reveal a correlation between CCR2-, G_i/o -, and Fyn-mediated tyrosine phosphorylation of p115 RhoGEF and its GEF activity. To gain more evidence of how tyrosine phosphorylation of p115 RhoGEF could lead to activation of Rac1, we tested for its capacity to form a complex with Rho GTPases. It was exciting to find that p115 RhoGEF, while having no effect on RhoA, forms a complex with Rac1 in a time-dependent manner in response to MCP1 (Fig. 6E). In addition, blockade of its tyrosine phosphorylation by inhibition of CCR2 or G_i/o or depletion of Fyn levels attenuated MCP1-induced p115 RhoGEF association with Rac1 (Fig. 6F). These results suggest a correlation between tyrosine phosphorylation of p115 RhoGEF and activation of Rac1.

Figure 6.

CCR2, G_i/o, and Fyn mediate p115 RhoGEF tyrosine phosphorylation and activation. A, growth-arrested HASMCs were treated with MCP1 (50 ng/ml) in the presence or absence of CCR2A (25 nm) or Pertussis toxin (PT) (50 ng/ml) for 10 min or left untreated, and cell extracts were prepared. Equal amounts of protein from the control and each treatment were immunoprecipitated (IP) with phosphotyrosine antibodies, and the immunocomplexes were immunoblotted (IB) with anti-p115 RhoGEF antibodies or analyzed by pulldown assay for Rac1 activation using GST-PAK1 beads. B, HASMCs that were transfected with control or Fyn siRNA (100 nm) and quiesced were treated with MCP1 (50 ng/ml) for 10 min or left untreated. Cell extracts were prepared and analyzed for p115 RhoGEF tyrosine phosphorylation and Rac1 activation as described in A. C and D, all conditions were the same as in A and B, respectively, except that the cell extracts were analyzed for p115 RhoGEF activity using a RhoGEF exchange assay kit as described under “Materials and Methods.” In A and B, equal amounts of protein from the control and each treatment were also analyzed by Western blotting for p115 RhoGEF, Rac1, or Fyn levels for normalization or to show the effect of siRNA on its target and off-target molecules. E, equal amounts of protein from the control and the indicated treatments were immunoprecipitated with anti-p115 RhoGEF antibodies or mouse IgG. The immunocomplexes were analyzed for Rac1 or RhoA levels using their specific antibodies, and the blot was normalized for p115 RhoGEF levels. F, top panel, growth-arrested HASMCs were treated with MCP1 (50 ng/ml) in the presence and absence of CCR2A (25 nm) or Pertussis toxin (PT) (50 ng/ml) for 30 min or left untreated, and cell extracts were prepared. Equal amounts of protein from the control and each treatment were immunoprecipitated with anti-p115 RhoGEF antibodies or mouse IgG, and the immunocomplexes were analyzed for Rac1 levels as described in E. Center and bottom panels, cells were transfected with control or Fyn siRNA, growth-arrested, treated with MCP1 (50 ng/ml) or left untreated, and analyzed for Rac1 association with p115 RhoGEF as described in E. The same cell extracts were analyzed by Western blotting for Fyn levels, and the blot was normalized for β-tubulin levels using specific antibodies to show the effect of the siRNA on its target and off-target molecule levels.

Discussion

The important findings of this study are as follows: MCP1 activates p115 RhoGEF in mediating HASMC migration and proliferation in vitro, and BI also triggers p115 RhoGEF stimulation in the mediation of VSMC migration and proliferation, leading to neointima formation in vivo. We have reported previously that injury induces the expression of MCP1 in a rat carotid artery, mediating its inward remodeling (5). Based on these observations, it may be suggested that MCP1, via activation of p115 RhoGEF, mediates the migration and proliferation of VSMCs in vascular wall remodeling. Many studies have shown the importance of Rho proteins (namely, RhoA, Rac1, and CDC42) in the regulation of cell migration and proliferation (29–31). The activation of Rho proteins requires their conversion from an inactive GDP-bound state to an active GTP-bound state, and this transition is dependent on the activation of guanine nucleotide exchange factors, RhoGEFs (16, 32). In mammals, RhoGEFs comprise a large family, with ∼70 members regulating the activities of about 20 Rho proteins (18). The p115 RhoGEF belongs to a small subfamily of regulator of G protein signaling–containing RhoGEFs, which includes PDZ RhoGEF and LARG (33, 34). It was demonstrated that the p115 RhoGEF mediates angiotensin II-induced vascular tone and blood pressure (27, 35). In addition, this study shows that MCP1-induced HASMC migration and proliferation and injury-induced vascular wall remodeling require the activation of p115 RhoGEF. A large body of data suggests that p115 RhoGEF exhibits specificity for RhoA (25, 36). However, our findings indicate that the p115 RhoGEF, without any effect on RhoA, mediates MCP1-induced Rac1 activation. Rac1 plays a central role in stress fiber formation, and many reports have shown its requirement for both cell migration and proliferation (37–39). In line with these observations, we have reported previously that activation of Rac1 is required for MCP1-induced HASMC stress fiber formation, migration, and proliferation (20). In addition, we have shown that Rac1 plays a role in injury-induced vascular wall remodeling (21).

We have demonstrated previously that NFATc1, via its role in cyclin D1 expression and CDK6 and CDk4 activation, mediates the phosphorylation and activation of PAK1, an effector of Rac1, in enhancing MCP1-induced HASMC migration and proliferation and injury-induced neointima formation (20). Furthermore, our studies have revealed that Rac1 acts upstream of NFATc1 in triggering cyclin D1 expression, CDK6 and CDK4 activities, and PAK1 phosphorylation in mediating MCP1-induced HASMC migration and proliferation and injury-induced neointima formation (20). In this study, we show that p115 RhoGEF acts upstream of Rac1 in mediating NFATc1 activation, cyclin D1 expression, CDK6 and CDK4 activities, and PAK1 phosphorylation in MCP1-induced HASMC migration and proliferation and injury-induced neointima formation. Many studies have shown that cyclin D1 and CDKs, besides their role in cell proliferation, mediate agonist-induced cell migration (20, 22, 40). In support of a role for CDKs in cell migration, previous studies from our laboratory as well as others have shown that CDK6 and CDK5 mediate PKN1 and PAK1 phosphorylation in VSMCs and neurons, respectively (22, 41). To establish connections between CDKs and Rho GTPase effectors, in this study we also show that MCP1-induced, CDK4-dependent PAK1 phosphorylation and activation in HASMCs require PKN1, as down-regulation of its levels using an siRNA approach attenuated MCP1-induced CDK4 and PAK1 activities. These findings also infer that CDK4 acts downstream of PKN1 in mediating PAK1 phosphorylation in MCP1-induced HASMC migration and proliferation. In addition, we observed that inhibition or down-regulation of p115 RhoGEF, Rac1, NAFTc1, Cyclin D1, or CDK6 in this sequential order attenuated their downstream molecules affecting PKN1 phosphorylation in response to MCP1, suggesting that PKN1 activation occurs downstream of CDK6 in MCP1-induced p115 RhoGEF–Rac1–NFATc1–cyclin D1–CDK6 signaling. These observations also reveal that MCP1-induced HASMC migration and proliferation require sequential activation of p115 RhoGEF, Rac1, NFATc1, cyclin D1/CDK6, PKN1, CDK4, and PAK1 signaling. PKN1 is a RhoA/Rac1 effector molecule and mediates cell migration by phosphorylating cytoskeletal proteins (42–44). The findings of this study suggest that one potential mechanism by which PKN1 might be involved in vascular wall remodeling could be via CDK4-dependent activation of PAK1. Furthermore, the finding that depletion of p115 RhoGEF levels attenuated BI-induced Rac1 activation, cyclinD1 expression, CDK6 and CDK4 activities, and PKN1 and PAK1 phosphorylation further supports our hypothesis that a signaling cascade that is activated in response to a potent chemoattractant protein, MCP1, in HASMCs in vitro also occurs in intact arteries in response to injury.

We have reported previously that CCR2-mediated G_i/o-dependent Fyn activation is required for MCP1-induced HASMC proliferation (26). In this regard, our findings reveal that CCR2, G_i/o, and Fyn signaling was also required for p115 RhoGEF tyrosine phosphorylation and perhaps for its activation by MCP1. It is interesting to note that the tyrosine phosphorylation of p115 RhoGEF also leads to its association with Rac1 while having no effect on RhoA in response to MCP1. Based on these observations, it is likely that tyrosine phosphorylation of p115 RhoGEF via CCR2, G_i/o, and Fyn is required for its association and activation of Rac1. Many reports have shown that tyrosine phosphorylation of RhoGEFs is essential for their GEF activities (27, 28). Furthermore, a role of Fyn in tyrosine phosphorylation and activation of Vav, another Rho family GEF, leading to Rac1 activation, has been reported (45, 46). In addition, previous studies from other laboratories have shown that CCR2 plays a role in the development of neointimal hyperplasia (47, 48). Similarly, a role of or G_i/o in the mediation of VSMC proliferation has been reported (49). Based on these observations and our findings, it may be suggested that CCR2–G_i/o–Fyn mediates p115 RhoGEF activation in the modulation of vascular wall remodeling. In summary, as depicted in Fig. 7, our results suggest that, downstream of CCR2, Gi/o, and Fyn, p115 RhoGEF, via Rac1-NFATc1-mediated cyclin D1 expression and CDK6–PKN1–CDK4–PAK1 signaling activation, enhances injury-induced vascular wall remodeling.

Figure 7.

Schematic of the potential mechanism(s) by which p115 RhoGEF could be activated by and involved in the mediation of MCP1-induced VSMC migration and proliferation and injury-induced neointima formation.

Materials and methods

Reagents

Pertussis toxin (P7208), pluronic F127 (P2443), and anti-SMCα-actin antibodies (A2547) were purchased from Sigma-Aldrich (St. Louis, MO). Recombinant human MCP1 (279-MC) was obtained from R&D Systems Inc. (Minneapolis, MN). Anti-CDK4 (SC-260), anti-CDK6 (SC-56362), anti-Fyn (SC-365913), anti-PKN1 (SC-136037), anti-p115 RhoGEF (SC-74565), and anti-β-tubulin (SC-9104) antibodies, the CCR2 antagonist (SC-202525), and truncated retinoblastoma (Rb) protein (SC-4112) were bought from Santa Cruz Biotechnology Inc., (Santa Cruz, CA). Anti-Ki67 (ab15580) and anti-pPAK1 (ab2477) antibodies were purchased from Abcam (Cambridge, MA). Anti-cyclin D1 antibody (RB-010-P) was obtained from NeoMarkers (Fremont, CA). Anti-NFATc1 antibody (MA3-024) was purchased from Affinity BioReagents (Golden, CO). Anti-PAK1 (2602) and anti-pPKN1 (2611) antibodies were obtained from Cell Signaling Technology (Beverly, MA). The ABC kit, DAB kit, and Vectashield mounting medium were bought from Vector Laboratories Inc. (Burlingame, CA). Hoechst 33342 (3570), Lipofectamine 2000 reagent, human Fyn siRNA (AM51331), rat p115 RhoGEF siRNA (Stealth siRNA RSS329430, NM_021694.3), and Prolong Gold antifade mounting medium (P36930) were obtained from Invitrogen. The RhoGEF Exchange Assay Biochem kit (BK100) was bought from Cytoskeleton (Denver, CO). [γ-³²P]ATP (specific activity 3000 Ci/mmol) was obtained from MP Biomedicals (Irvine, CA). Protein A-Sepharose (CL-4B) and protein G-Sepharose beads were purchased from GE Healthcare. Human cyclin D1 siRNA (ON-TARGETplus SMARTpool L-003210-00, NM-053056), human CDK4 siRNA (ON-TARGETplus SMART-pool L-003238-00, NM_000075), human CDK6 siRNA (ON-TARGETplus SMARTpool L-003240-00, NM_001259), human NFATc1 siRNA (ON-TARGETplus SMART-pool L-003605-00, NM_172390), human PAK1 siRNA (ON-TARGETplus SMARTpool L-003521-00, NM_002576), human PKN1 siRNA (ON-TARGETplus SMARTpool L-004175-00, NM_005585), and control nontargeting siRNA (D-001810-10) were bought from Dharmacon RNAi Technologies (Chicago, IL).

Adenoviral vectors

The construction of Ad-GFP and Ad-dnRac1 was described previously (50, 51).

Cell culture

HASMCs were purchased from Cascade Biologicals (Portland, OR) and subcultured in medium 231 containing smooth muscle cell growth supplements, gentamycin (10 μg/ml), and amphotericin (250 ng/ml). Cells were used between four and ten passages.

CDK4/6 assays

Cell or tissue extracts containing equal amounts of protein were analyzed for CDK4/6 activities using an immunocomplex kinase assay as described previously (20, 22). The ³²P-labeled Rb protein bands were visualized by autoradiography, and their intensities were quantified using National Institutes of Health ImageJ.

DNA synthesis

DNA synthesis was measured by [³H]thymidine incorporation as described previously and expressed as counts per minute per dish (52).

Cell migration

Cell migration was measured by wound healing assay (53). Wherever an adenovirus was used, cells were infected with the respective adenovirus at a multiplicity of infection of 40 and growth-arrested before being subjecting to a migration assay. Cell migration is expressed as percentage of wound closure (total area − area not occupied by cells / total area × 100).

Western blotting

Cell or tissue extracts containing equal amounts of protein were analyzed by Western blotting for the indicated protein using its specific antibody as described previously (22). The band intensities were quantified using National Institutes of Health ImageJ.

RhoGEF exchange assay

Equal amounts of protein from the control and the indicated treatments were immunoprecipitated with anti-p115 RhoGEF antibodies overnight at 4 °C. The immunocomplexes were pulled down with protein A/G-Sepharose beads, washed with lysis buffer (1% Nonidet P-40, 20 mm HEPES (pH 7.4), and 150 mm NaCl), eluted with 0.2 m glycine buffer (pH 2.5), neutralized with 1 m Tris buffer (pH 8.0), and assayed for p115 RhoGEF activity using the RhoGEF Exchange Assay Biochem Kit according to the instructions of the manufacturer. Briefly, 2 μm Rac 1 or RhoA was added to an exchange buffer containing 0.75 μm N-methylanthraniloyl (mant)-GTP and allowed to equilibrate. After equilibration, the reaction mix was aliquoted (100 μl/well) into a 96-well plate, and fluorescence measurements were taken approximately every 30 s with excitation and emission wavelengths of 360 nm and 440 nm, respectively. After five readings (150 s), 10 μl of sample, negative control (buffer), or positive control (hDbs protein) was added, and the relative fluorescence intensity was measured every 30 s for 30 min in a SpectraMax GeminiXPS spectrofluorometer at room temperature. Lysis buffer was used as a negative control. The specific exchange rate was calculated according to the formula of the manufacturer and expressed as relative fluorescence units (RFU).

Rac1 and RhoA pulldown assay

Equal amounts of protein from the control and the indicated treatments were incubated with 50 μl of 50% (w/v) GST-PAK1 or GST-Rhotekin beads overnight at 4 °C. After incubation, the beads were collected by centrifugation, washed with lysis buffer, and heated in Laemmli sample buffer, and the released proteins were analyzed by Western blotting for Rac1 and RhoA, respectively, using their specific antibodies. Bound Rac1 and RhoA levels were expressed as active Rac1 or active RhoA.

BI

The experiments involving animals were performed according to protocols approved by the Institutional Animal Care and Use Committee of the University of Tennessee Health Science Center (Memphis, TN). BI was performed essentially as described previously (22). Non-targeting siRNA or p115 RhoGEF siRNA in combination with Lipofectamine 2000 was mixed in 30% (w/v) pluronic gel and applied around the injured artery as described by Subramanian et al. (54). 1 day, 3 days, 5 days, or 2 weeks after BI, the animals were sacrificed with an overdose of ketamine/xylazine (200 mg/kg), and carotid arteries were isolated. The arteries were cleaned, and tissue extracts were prepared or processed for morphometric analysis. For morphometric analysis, arteries were fixed in 10% formalin, embedded in OCT compound (Sakura Finetek USA Inc., Torrance, CA), and 5-μm-thick sections were cut at equally spaced intervals in the middle of the injured common carotid artery and stained with hematoxylin and eosin. The intimal (I) and medial (M) areas were measured using National Institutes of Health ImageJ, and the I/M ratios were calculated.

In vivo SMC migration assay

In vivo SMC migration was measured as described by Bendeck et al. (55). Briefly, 3 days after BI, the carotid arteries were fixed in vivo with 10% formalin at physiological pressure. The middle 1 cm of the denuded (injured) common carotid artery was cut and fixed again in cold acetone for 10 min. The artery was then opened longitudinally and pinned down onto an agar plate with the luminal surface facing up. The arteries were rinsed with PBS and placed in 0.3% H₂O₂ for 30 min to block endogenous peroxidase activity. After blocking in 5% goat serum in PBS for 30 min, the arteries were incubated with anti-SMCα actin antibody (1:300) for 1 h, followed by incubation with biotinylated goat anti-mouse IgG for 30 min. Peroxidase labeling was carried out using an ABC kit, and the signals were visualized with a DAB kit. Finally, the arteries were placed on glass slides with the luminal surface facing up, and coverslips were placed. As a negative control, samples of the same specimens without incubation with the primary antibody were used. The luminal surface of the artery was examined under a light microscope at ×200 magnification, and the number of SMCα actin-positive cells were counted. In vivo SMC migration was expressed as number of cells per 0.1 mm² of the luminal surface.

Double immunofluorescence staining

Soon after isolation, the carotid arteries were snap-frozen in OCT compound. Cryosections (10 μm thick) were made using a cryostat (model CM3050S, Leica, Wetzlar, Germany). After blocking in goat serum, the cryosections were incubated with rabbit anti-rat Ki67 antibodies and mouse anti-rat SMCα actin antibodies for 1 h. After washing in PBS, the sections were incubated with goat anti-rabbit secondary antibodies conjugated with Alexa Flour 568 and goat anti-mouse secondary antibodies conjugated with Alexa Flour 488. Negative controls were processed in exactly the same way, except that incubation with primary antibodies was eliminated. The slides were observed under a Zeiss Oberver.Z1 fluorescence microscope, and the images were captured using an AxioCam MRm camera. SMC proliferation was measured as number of SMCα actin and Ki67-positive cells per field.

Statistics

All experiments were repeated three times with similar results. Data are presented as the means ± S.D. The treatment effects were analyzed by Student's t test, and p <0.05 was considered statistically significant. In the case of Western blotting, immunohistochemistry, and CDK4/6 activities, one set of the representative data is shown.

Author contributions

N. K. S. performed cell migration, proliferation, Western blotting, balloon injury, histochemical staining, and immunofluorescence staining and wrote the initial draft of the manuscript. J. J. performed immunofluorescence staining, cell migration, and DNA synthesis. G. N. R. conceived the overall scope of the project, planned the experiments, interpreted the results, and contributed to writing of the manuscript.