PPAR inhibits vascular smooth muscle cell proliferation underlying intimal hyperplasia by inducing the tumor suppressor p16INK4a
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《临床调查学报》
1INSERM U545, Departement d’AtHÉrosclerose, Institut Pasteur de Lille et Faculte de Pharmacie, Universite Lille II, Lille, France.
2CHU d’Amiens and Faculte de Medecine, Universite Jules Verne de Picardie, Amiens, France.
3Department of Pharmacological Sciences, University of Milan, Milan, Italy.
4Division of Molecular Genetics and Centre for Biomedical Genetics, The Netherlands Cancer Institute, Amsterdam, The Netherlands.
Abstract
Vascular SMC proliferation is a crucial event in occlusive cardiovascular diseases. PPAR is a nuclear receptor controlling lipid metabolism and inflammation, but its role in the regulation of SMC growth remains to be established. Here, we show that PPAR controls SMC cell-cycle progression at the G1/S transition by targeting the cyclin-dependent kinase inhibitor and tumor suppressor p16INK4a (p16), resulting in an inhibition of retinoblastoma protein phosphorylation. PPAR activates p16 gene transcription by both binding to a canonical PPAR-response element and interacting with the transcription factor Sp1 at specific proximal Sp1-binding sites of the p16 promoter. In a carotid arterial–injury mouse model, p16 deficiency results in an enhanced SMC proliferation underlying intimal hyperplasia. Moreover, PPAR activation inhibits SMC growth in vivo, and this effect requires p16 expression. These results identify an unexpected role for p16 in SMC cell-cycle control and demonstrate that PPAR inhibits SMC proliferation through p16. Thus, the PPAR/p16 pathway may be a potential pharmacological target for the prevention of cardiovascular occlusive complications of atherosclerosis.
Introduction
Activation of vascular SMC proliferation is a key event in the development of atherosclerosis and its complications (1). In response to vascular injury, SMCs migrate from the media into the intimal layer of the arterial wall, where they proliferate and synthesize extracellular matrix, resulting in the formation of intimal hyperplasia. In this process, vascular SMCs undergo phenotypic changes from a differentiated and contractile state to a dedifferentiated and synthetic state (2). (Dys)regulation of SMC growth occurs thus during atherosclerotic plaque formation as part of a local inflammatory response in association with accumulation of lipids and fibrous connective tissue in the vascular wall (3). SMC proliferation is also the primary pathophysiological mechanism underlying complications of procedures used to treat atherosclerotic diseases, such as restenosis, a secondary occlusion of the arterial wall following transluminal angioplasty or stent implantation, and vein bypass graft failure (1).
As a key early event in these phenotypic changes, SMCs, which are in a quiescent state (G0) in normal uninjured vessels, transit through the G1 phase of the cell cycle and enter into the S phase to undergo replication (4). Cell-cycle progression is under the control of cyclin-dependent kinases (CDKs), which phosphorylate different specific target proteins through the 4 stages of the cell cycle (5). Notably, phosphorylation of the retinoblastoma gene product (pRB) by the specific G1 CDKs represents the critical checkpoint of G1/S transition (6). When underphosphorylated, pRB sequesters the E2F family transcription factors, which regulate genes encoding proteins required for S phase DNA synthesis. Phosphorylation of pRB releases E2F that permits the induction of E2F-dependent genes and therefore the irreversible induction of the mitosis process, after which cells are refractory to extracellular growth-inhibition signals. Thus, increased pRB phosphorylation correlates with the induction of SMC proliferation in injured vessels (1, 6). As CDKs are constitutively present in each phase of the cell cycle, CDK-mediated pRB phosphorylation is respectively activated or inhibited through timely interactions with expressed cyclins and CDK inhibitors (CDKIs) (7). Notably, the CDKI p16 is a tumor suppressor transcriptionally regulated by pRB, which plays a key role in the control of pRB phosphorylation and G1/S cell-cycle progression (8-11).
PPAR is a nuclear receptor that, upon ligand activation, regulates transcription after dimerizing with the retinoid X receptor (RXR) and DNA-binding to PPAR-response elements (PPREs) within the regulatory regions of target genes (12). PPREs usually consist of a direct repeat of the hexanucleotide AGGTCA sequence separated by 1 or 2 nucleotides (DR1 or DR2) (12). Furthermore, PPAR can also negatively interfere with proinflammatory signaling pathways by a mechanism termed "transrepression" (13). PPAR is activated by natural ligands such as fatty acids and derivatives (13). Notably, natural eicosanoids derived from arachidonic acid via the lipoxygenase pathway as well as oxidized phospholipids activate PPAR (13). Moreover, fibrates (gemfibrozil, bezafibrate, ciprofibrate, and fenofibrate), synthetic drugs initially used to treat lipid disorders (14), act as PPAR ligands (15).
Fibrates have been shown to decrease morbidity and mortality of cardiovascular diseases (16) and impede the progression of coronary atherosclerotic lesions when administered to postinfarction or diabetic patients (17-19). PPAR mediates their systemic metabolic effects by regulating genes encoding proteins involved in lipid and lipoprotein metabolism (20). Recently, the functions of PPAR have been extended to a regulatory action on cholesterol homeostasis and vascular inflammation acting directly at the level of the vascular wall, which could confer additional protective effects in the prevention of atherogenesis (13). Importantly, PPAR is also highly expressed in SMCs isolated from vascular tissues and from atherosclerotic lesions (21-23), where it controls IL-1–induced secretion of IL-6 and production of prostaglandins by inhibiting IL-6 and COX2 gene transcription. However, although recent studies have indicated that fibrates may regulate SMC proliferation in vitro (21, 24), a role for PPAR in SMC cell-cycle control has not yet been explored. In the present study, we demonstrate a role for PPAR in the control of SMC G1/S transition. Moreover, we identify the CDKI p16 as an important controller of vascular SMC proliferation and show that p16 mediates the effects of PPAR on SMC proliferation both in vitro and in vivo.
Results
PPAR inhibits SMC proliferation by controlling cell-cycle progression into the S phase.
Previous studies have indicated that certain fibrates may regulate human SMC (hSMC) proliferation in vitro (21, 24). To further analyze the effect of PPAR activation on SMC growth, the influence of a novel generation subtype-specific PPAR agonist, GW7647, which presents a high affinity (EC50 = 6 nM) and specificity (200-fold selectivity over PPAR and PPAR&Dgr;) (25), was first tested on hSMCs resuming proliferation in normal growth medium (GM) after G0/G1-phase synchronization (Figure 1A). Whereas vehicle-treated cells displayed a growth rate comparable to that of untreated cells (not shown) with a doubling time of approximately 24 hours, GW7647 treatment inhibited proliferation in a dose-dependent manner. To examine which stage(s) of the cell-cycle is (are) regulated by GW7647, the cell-cycle distribution of hSMCs resuming growth in GM was analyzed by flow cytometry analysis after 48 hours of treatment with GW7647 or vehicle. As shown in Table 1, GW7647 treatment increased and decreased the proportion of hSMCs in the G0/G1 and S phases, respectively.
To further characterize the exact role of PPAR in SMC cell-cycle control, aortic mouse SMCs (mSMCs) were isolated from PPAR–/– and PPAR+/+ mice. Interestingly, PPAR–/– mSMCs isolated from both Sv/129 (Figure 1B) and C57BL/6J (Supplemental Figure 1; supplemental material available online with this article; doi:10.1172/JCI22756DS1 http://dx.doi.org/10.1172/JCI22756DS1) genetic backgrounds proliferated at a much higher rate than corresponding PPAR+/+ mSMCs. Moreover, incubation of mSMCs with Wy14,643, a specific murine PPAR agonist, inhibited cell proliferation of PPAR+/+, but not PPAR–/– mSMCs (Figure 1B). Similarly to hSMCs, PPAR activation of mSMCs (assessed after 48-hour treatment in GM) was associated with a higher cell percentage in G0/G1 and a concomitant decrease of cell percentage in the S phase (Table 1).
Concomitantly, phosphorylation of pRB, the major checkpoint of G1/S progression (5), was reduced in GW7647-treated hSMCs, an effect which was even more pronounced upon infection with adenovirus-PPAR (Ad-PPAR) (Figure 1C). Altogether, these results indicate that PPAR activation inhibits both human and mouse vascular SMC proliferation by interfering with G1/S cell-cycle progression.
PPAR induces transcription of the G1/S transition regulator and tumor suppressor p16.
Since PPAR is a nuclear receptor acting at the transcriptional level (12) and since the tumor suppressor p16 is a transcriptionally regulated CDKI controlling G1/S cell-cycle progression functionally related to in-cell pRB expression and phosphorylation levels (8-10), we hypothesised that p16 could be a PPAR target gene. Indeed, in both mSMCs and hSMCs, p16 mRNA levels were increased upon PPAR agonist incubation (Figure 2A). The induction of p16 mRNA occurred in a PPAR-dependent manner and was already evident within 4 hours of treatment (Figure 2A). In hSMCs, the increase of p16 mRNA levels upon GW7647 treatment was more pronounced in Ad-PPAR–infected hSMCs (Figure 2A, upper panel). Moreover, Wy14,643 treatment induced p16 expression in PPAR+/+, but not in PPAR–/– mSMCs (Figure 2A, lower panel). Interestingly, p16 mRNA levels were decreased by approximately two-thirds in PPAR–/– compared with PPAR+/+ mSMCs.
Previously, a pRB-responsive site and proximal Sp1-binding sites (denoted A, B, and C) have been shown to be involved in the basal activity of the p16 promoter (11, 26). Here, the presence of a putative degenerated DR1 PPRE sequence (AGGAGACAGGACA) at position –1023 to –1011 (denoted the p16DR1 site) was also revealed by computer-assisted analysis. To investigate the putative function of these elements in mediating the response to ligand-activated PPAR, reporter constructs driven by different p16 promoter fragments were cotransfected with or without the pSG5-PPAR expression plasmid, and cells were subsequently treated with the PPAR agonists fenofibric acid or GW7647. Cotransfection of PPAR followed by ligand activation significantly induced activity of the –3.0-kb p16 gene promoter, indicating that p16 is a direct PPAR target gene (Figure 2B). PPAR-dependent p16 promoter activation was maintained upon 5' deletion to –1.0 kb. Further nucleotide deletions to –869 and –703 bp, as well as mutation of p16DR1, resulted in a significant, albeit incomplete reduction of p16 promoter activation by PPAR. Finally, deletion to –343 bp completely abolished activation of the p16 promoter by fenofibric acid–activated PPAR (Figure 2B). Identical results were obtained in cells treated with GW7647 (250 nM) (data not shown). These observations indicate that both p16DR1 and Sp1A–C elements are involved in PPAR-dependent p16 promoter activation, with the DR1 element mediating the majority of the response.
To determine whether PPAR physically binds to the p16DR1 site, electrophoretic mobility shift assays (EMSAs) were performed using in vitro–produced PPAR and RXR proteins and double-stranded [32P]-labeled oligonucleotides (Table 2 and Figure 2C). Whereas neither RXR nor PPAR alone bound to the p16DR1 probe, a clear shift was observed upon incubation with both RXR and PPAR. This complex was supershifted by the anti-PPAR antibody and prevented by an excess of unlabeled wild-type, but not mutated, p16DR1 probe. Moreover, the mutated p16DR1 probe did not bind RXR/PPAR (Figure 2C). These results demonstrate that the RXR/PPAR heterodimer specifically binds in vitro to the p16DR1 site located at –1023 pb. In contrast, no PPAR/DNA complex formation was detected in EMSAs performed with oligonucleotides overlapping the proximal Sp1A- and B-binding sites (data not shown).
To investigate the in vivo occupancy of these p16DR1 and Sp1-binding sites by activated PPAR, chromatin IP (ChIP) assays were performed on DNA from hSMCs treated or not treated with fenofibric acid using anti-Sp1 and anti-PPAR antibodies (Figure 2D). In accordance with the EMSA results, the region spanning the p16DR1 sequence was immunoprecipitated by the anti-PPAR antibody, an effect which was enhanced by fenofibric acid treatment. In addition, in agreement with results of Myhnen et al. (26), genomic p16 promoter DNA regions encompassing the Sp1-binding elements were immunoprecipitated by the anti-Sp1 antibody. Interestingly, similarly to with the DR1 element, the region encompassing the Sp1A and C sites also bound PPAR in a PPAR ligand–dependent manner (Figure 2D). In contrast, no PPAR binding was observed on the Sp1B site. As negative controls, PCR amplification using primers covering a region localized just downstream of the –1023DR1 site or a -actin genomic fragment did not yield a significant signal, thus demonstrating the specificity of PPAR immunoprecipitation and PCR amplification. Taken together, these results indicate that PPAR activates the p16 promoter through direct DNA binding to the p16DR1 site as well as tethering with Sp1 to the proximal Sp1A–C response element.
Induction of p16 following PPAR activation, resulting in the inhibition of CDK4-mediated pRB phosphorylation, is required for the SMC growth-inhibitory activity of PPAR.
Next, it was determined whether the upregulation of p16 gene expression by PPAR results in an increase of p16 protein abundance and in-cell activity. Western blot analysis of hSMC whole-cell protein extracts revealed that PPAR overexpression or GW7647 treatment enhanced p16 protein levels in hSMCs, and the greatest increase was observed when cells were both infected with Ad-PPAR and treated with GW7647 (Figure 3A).
Initially, p16 was identified by virtue of its ability to inhibit G1/S cell-cycle progression by binding to CDK4, thus preventing CDK4 association with D-type cyclins and further initiation of pRB phosphorylation by the cyclin D/CDK4 complexes (8, 9, 27). To verify that PPAR controls p16 function, it was further determined whether the induction of p16 following PPAR activation affected p16/CDK4 interaction and CDK4 activity. To this end, in-cell pRB phosphorylation by CDK4, determined using an anti–PSer780-ppRB antibody (28), and in vitro CDK4-associated activity using pRB as substrate were measured after PPAR overexpression and/or activation with GW7647 in hSMCs (Figure 3, B and C, upper panel). In fact, concurring with the decrease of the phosphorylated/nonphosphorylated pRB ratio following PPAR activation (Figure 1C), both analyses indicated that treatment with the PPAR ligand decreases pRB phosphorylation by CDK4 and that this effect is enhanced by PPAR overexpression. Moreover, the levels of p16 associated with CDK4 were increased in hSMCs infected with Ad-PPAR or treated with GW7647, and this interaction was further enhanced in Ad-PPAR–infected GW7647-treated hSMCs (Figure 3C, middle panel). Besides, in-cell p16 protein levels and CDK4-dependent pRB phosphorylation respectively increased and decreased in PPAR+/+ but not in PPAR–/– mSMCs following Wy14,643 treatment (Figure 3D).
Finally, the role of p16 in mediating the growth-inhibitory effects of PPAR activation was analyzed in vitro using SMCs isolated from p16–/– mice (29). Strikingly, primary mSMCs isolated from p16–/– mice proliferated at a much higher rate than p16+/+ mSMCs (Figure 4A). Moreover, p16–/– mSMCs did not respond to Wy14,643 treatment. Accordingly, the ratio of cell percentage in S compared with G1 phases (data not shown) as well as in-cell CDK4-mediated pRB phosphorylation (Figure 4B) were enhanced in p16–/– mSMCs and not affected by Wy14,643 treatment. Altogether, these data indicate that the G1 cell-cycle arrest induced by ligand-activated PPAR occurs via inhibition of the activity of G1 CDKs, such as CDK4, due to an increased recruitment of p16.
PPAR activation inhibits in vivo SMC proliferation upon carotid arterial injury by increasing p16 gene expression.
The in vivo role of PPAR in the control of SMC proliferation was next examined by using a mechanical carotid artery injury mouse model (30) in Sv/129 mice, a strain highly susceptible to intimal hyperplasia (31). The neointimal thickness in response to injury was determined by measuring the intima/media (I/M) area ratio throughout the entire injured segment starting from the junction between the internal and external left carotid branches. In agreement with previous reports (31, 32), injury induced a marked increase in intimal hyperplasia in wild-type mice as observed 3 weeks after injury (Figure 5, A and B). Interestingly, the I/M ratio was markedly enhanced in PPAR–/– mice,whereas it was drastically decreased in PPAR+/+ mice treated with fenofibrate, whose action in mice requires PPAR expression (33, 34) (Figure 5, A and B). As previously reported (2, 21, 30), SMCs, identified by -actin staining, were abundantly present in the media of uninjured carotids and constituted the major cell type underlying neointimal development in response to injury in this model (Figure 5, A–C, see PPAR+/+). Moreover, in agreement with the I/M ratio results, neointimal SMC and proliferating cell nuclear antigen–staining (PCNA-staining) were increased in carotid sections of PPAR–/– mice, and diminished in fenofibrate-treated mice compared with untreated wild-type mice (Figure 5, B and C). In agreement with the in vitro proliferation assays performed in mSMCs (Supplemental Figure 1), PPAR deficiency also provoked hyperplasia in C57BL/6J mice (Supplemental Figure 2), a strain reported to be more resistant to hyperplasia formation after carotid injury (31, 32).
Furthermore, p16 gene expression was measured in the uninjured carotid segment, which contains mainly SMCs in a quiescent state (1, 4) (Figure 5C), as well as in carotid segments at different times after injury (Figure 5D). Strikingly, p16 mRNA levels were induced after response to injury in PPAR+/+ but not in PPAR–/– mice, an effect that was further enhanced following fenofibrate treatment. A maximal PPAR-dependent induction of p16 mRNA levels was reached within 24 hours and remained elevated 3 weeks after injury. Thus, these data indicate that PPAR activation negatively controls neointima formation in response to injury, possibly via the induction of p16 gene expression in injured vessels.
Finally, the in vivo role of p16 in mediating the growth-inhibitory effects of PPAR activation were further analyzed using p16–/– mice (29). It is noteworthy that, similarly to what occurred in PPAR–/– mice (Figure 5 and Supplemental Figure 2), intimal hyperplasia was drastically increased in p16–/– mice (Figure 6). An extended neointima formation was observed over the entire lesion segment, which was accompanied by an increase in SMC abundance. Strikingly, fenofibrate treatment did not influence the I/M ratio in these p16–/– mice (Figure 6A). Therefore, p16 is required for the inhibitory effects of PPAR activation on vascular SMC growth underlying intimal hyperplasia development.
Discussion
The cardiovascular protective effects of fibrates were initially mainly attributed to a systemic action of PPAR on atherogenic dyslipidemia, characterized by a lowering of triglyceride levels and an increased concentration of plasma HDL-cholesterol concentrations (13). However, a number of recent clinical studies, including the Bezafibrate Coronary Atherosclerosis Intervention Trial (BECAIT; ref. 19), the Lopid Coronary Angiography Trial (LOCAT; ref. 17), and the Diabetes Atherosclerosis Intervention Study (DAIS; ref. 17), have revealed that fibrates inhibit atheromatous plaque progression via effects possibly independent or in addition to their systemic action. These observations are in line with results from in vivo studies performed in rabbits, which indicated that fibrate treatment can inhibit atherosclerosis without affecting plasma lipid profiles (35, 36). Moreover, in the secondary prevention Veterans Administration–HDL-Cholesterol Intervention Trial (VA-HIT), a decreased incidence of cardiovascular events following fibrate treatment was observed, which could not be attributed only to the effects of drug treatment on HDL levels (37). Concurring with these clinical data, compelling mechanistic evidence has recently emerged that PPAR controls cholesterol efflux and the inflammatory response in vessel wall cells (13), highlighting a direct regulatory role of PPAR on vessel wall function. However, despite the crucial role of SMC proliferation in the development of occlusive vascular diseases, the function of PPAR in the control of vascular SMC cell-cycle progression was not yet established. In the present study, we show that PPAR activation inhibits G1/S cell-cycle progression of both human and mouse primary SMCs in vitro, an effect attributed to the induction of the tumor suppressor gene p16. Upregulation of p16 by PPAR agonists results in the sequestration of CDK4 and the consequent decrease of CDK4-mediated pRB phosphorylation, thus providing a mechanism for the SMC G1 cell-cycle arrest induced by fibrates. The in vivo role of PPAR in the control of SMC proliferation through the induction of p16 gene expression was further demonstrated by studying the response to mechanical carotid artery injury (30). This carotid injury model is characterized by neointima formation due to an excessive accumulation of SMCs and deposition of extracellular matrix in the intimal layer of the vessel wall, a response which has been proposed to appropriately reflect the "in-stent" restenosis observed in humans (30).
In the search for the molecular mechanisms involved in the p16-dependent growth-inhibitory effects of PPAR on SMCs, PPAR activation was found to upregulate p16 promoter activity, thus identifying for the first time a molecular mechanism via which PPAR directly interferes with cell-cycle progression. It is noteworthy that numerous common disease predisposition factors and pathways between cancer and vascular remodeling pathologies have been identified, pointing to similarities between the development of both diseases (38). It has been suggested that both vascular diseases and cancer similarly develop from a clonal proliferation of altered cells at the sites of local tissue injury, inflammation, and/or genomic alterations (38). Similar to tumor development, the induction of intimal hyperplasia in response to injury involves the conversion of SMCs from a medial nonproliferating differentiated phenotype into a proliferating phenotype associated with pRB hyperphosphorylation (1). However, the molecular mechanisms underlying the relationship between these diseases remain to be characterized. Interestingly, the CDKI and tumor suppressor p27Kip1 (p27), whose protein degradation is prevented by PPAR activators (39), has been shown to inhibit the development of vascular diseases when transferred using an adenoviral vector into balloon-injured porcine arteries (40) as well as through bone marrow–derived immune cells (41). In this study we now report an endogenous regulatory role for the CDKI p16 as SMC growth inhibitor, which furthermore directly mediates the response to fibrates. In fact, p16 expression is correlated to both pRB phosphorylation and expression levels and is required for the negative feedback of G1/S transition (11, 27). Several studies have established that loss of p16 function can be an early event in tumor progression (42, 43) and that it is associated with tumor development (29, 44, 45). Thus, p16 promoter activity is currently utilized as a prognosis factor for several cancers, such as acute lymphoblastic leukemia, lymphoma, lung carcinoma, and melanoma (45). Here, we identify, for what we believe is the first time, a role for this CDKI, so far studied in cancer pathology, in vascular SMC proliferation underlying occlusive cardiovascular diseases such as restenosis (1).
Strikingly, PPAR deficiency and p16 deficiency both acted as molecular switches inducing SMC proliferation in vitro and in vivo (Figures 1 and 4 and Supplemental Figures 1 and 2). Interestingly, the induction of p16 expression alone and in response to fenofibrate treatment was much higher in response to injury than in healthy carotid arteries. Moreover, no significant morphological differences were observed in healthy carotid arteries isolated from untreated PPAR+/+ compared with PPAR–/– or fenofibrate-treated mice. Among the genes of the INK4 family, the CDKI p16 is specifically induced in response to oncogenic stress when it acts as a negative retrocontrol of increased pRB phosphorylation (27). In parallel, an important role for PPAR in the response to stress has been demonstrated previously upon fasting (46) or long-term high-fat feeding (47) as well as in the response to inflammatory factors such as leukotriene B4 (48), lipopolysaccharide (LPS), or IL-1 (49). Thus, our results demonstrating that both PPAR and p16 proteins are activated and function locally at the vascular artery injury site further extend these observations of a functional role for PPAR and p16 primarily in the response to stress.
Recently, substantial progress has been made in the prevention of restenosis, notably with the development of stents coated with antimitotic drugs, such as rapamycin (50). However, a potential risk of late thrombus formation associated with incomplete reendothelialization after surgery with these coated stents exists (50). It is noteworthy that, in our mouse model of restenosis, reendothelialization was completed 3 weeks after injury for all studied groups (Supplemental Figure 3). Moreover, thiazolidinedione PPAR activators —which have previously been shown to control vascular SMC proliferation (39, 51) — were recently shown to reduce coronary in-stent restenosis in type 2 diabetic patients (52, 53). The present report indicates a role for the PPAR/p16 pathway in vascular SMC proliferation in vitro and in a mouse model of restenosis. Further clinical studies are thus warranted to assess whether pharmacological activation of PPAR could provide a novel therapeutic strategy to prevent restenosis in treatments using either systemic administration or coated stents.
Methods
Reagents.
Wy14,643 and fenofibric acid were purchased from Sigma-Aldrich and Laboratoires Fournier SA, respectively. GW7647 (25) was kindly provided by P. J. Brown from GlaxoSmithKline (Research Triangle Park, North Carolina, USA).
Reverse transcription and quantitative PCR.
RNA extraction, reverse transcription, and quantification of mRNA levels were performed using an MX 4000 apparatus (Stratagene) as previously described (54). 28S RNA was measured as internal control. For in vitro assays, values are expressed as means ± SD of a representative experiment performed in triplicate (for a total of 3 samples measured).
Plasmids and Ad generation.
The Sp1 expression plasmid cloned in pcDNA3 was a gift from D. Monte (Institut de Biologie de Lille, Lille, France); the p16 reporter pGL2 (Promega) constructs were provided by G. Peters (Imperial Cancer Research Fund Laboratories, London, United Kingdom) (11). Mutations of the –1023 p16DR1 were introduced using the QuikChange site-directed mutagenesis kit (Stratagene) and the oligonucleotide mutated p16DR1 (p16DR1mt) (Table 2). The nucleotide positions are indicated relative to the ATG site. Recombinant Ad-GFP and Ad-PPAR were obtained by homologous recombination in Escherichia coli (55) after insertion of the cDNAs into the pAdCMV2 vector (Qbiogene). Viral stocks were created and titrated as previously described (56). SMCs (0.5 x 106) were infected at an input multiplicity of 100 virus particles/cell by adding virus stocks directly to the culture medium. Efficiency of hSMC infection was between 75–85%, as assessed by FACS analysis with the Ad-GFP vector.
Cell culture assays.
Human cervical HeLa cells and primary human aortic SMCs (hSMCs) were obtained from ATCC and BioWhittaker Europe, respectively. HeLa cells were cultured in DMEM medium (Invitrogen Corp.) supplemented with 10% FCS (HyClone), and L-glutamine (2 mM) as well as the antibiotics penicillin (50 U/ml) and streptomycin (55 μg/ml) (Gibco; Invitrogen Corp.). hSMCs were cultured using the SMC Growth Medium 2 Bullet Kit , purchased from BioWhittaker Inc. Mouse primary aortic SMCs (mSMCs) were prepared from aortas of 8- to 12-week-old PPAR+/+ or PPAR–/– mice on Sv/129 or C57BL/6J genetic backgrounds or of p16+/+ or p16–/– mice on a C57BL/6J genetic background as described (57) and were studied between the third and twelfth passages. mSMCs were cultured in DMEM supplemented with 10% FCS and antibiotics. In all assays, cell synchronization in the G0/G1 phase was induced by serum starvation for at least 24 hours in medium containing 0.4% FCS. To reinduce cell proliferation, cells were reincubated in routine GM (time 0 or T0 of the experiments). At T0, mSMCs were treated with the indicated PPAR ligand or an identical volume of Me2SO (vehicle). hSMCs were infected overnight and subsequently treated with PPAR ligands or vehicle as indicated. Three independent preparations from different mice were studied for each primary SMC type and resulted in similar results.
For cell proliferation assays, cellular DNA content was determined using fluorochrome 3,5-diaminobenzoicacid free acid (DABA) as described previously (58). Flow cytometry analysis profiles were determined by propidium iodide staining of nuclear DNA as described (59).
For transient transfection assays, HeLa cells in 24-well plates at an initial density of 1.5 x 104 per well were transfected with ExGen 500 (Euromedex) at a ratio of 4 μl ExGen 500/0.5 μg of DNA. The indicated pGL2 reporter constructs (100 ng) were cotransfected with the indicated expression vectors and the internal control Renilla luciferase reporter plasmid pRL-null (10 ng). The amount of plasmids in the transfection mixture was equalized to 0.5 μg by adding pcDNA3 vector. After 12 hours, cells were incubated for an additional 10 hours in fresh medium supplemented with fenofibric acid (100 μM) or Me2SO (vehicle). Luciferase activity was assessed with the Dual-Luciferase Reporter Assay System (Promega) as previously described (54). Transfections were carried out in triplicate, and each experiment was repeated at least 3 times.
Protein assays.
Cells were washed twice with ice-cold PBS and then collected in ice-cold RIPA buffer (1x PBS, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS, phosphatase and protease inhibitors) or kinase lysis buffer (50 mM Tris, pH 8.0, 0,1% IGEPAL; Sigma-Aldrich). Lysates were incubated for 30 minutes on ice, and cell debris was removed by centrifugation. Extracts were then aliquoted and stored at –80°C.
For coimmunoprecipitation assays, whole-cell lysates were preincubated with 1.0 μg nonimmune rabbit IgG and 20 μl of protein A-Sepharose beads (Amersham Biosciences) during 30 minutes at 4°C and centrifuged. Cleared lysates (1 mg) were immunoprecipitated for 1 hour at 4°C with the anti-CDK4 antibody (RB-016, Microm Microtech France) and then incubated overnight at 4°C with protein A-Sepharose. The immunoprecipitated proteins were washed with lysis buffer and resuspended in electrophoresis sample buffer.
Samples of immunoprecipitated or total proteins (20–30 μg) were analyzed by Western blot using the following primary antibodies: anti-CDK4 (RB-003; Microm Microtech France), anti-p16 (554079 for hSMCs, BD Biosciences — Pharmingen; and MS-887 for mSMCs, Microm Microtech France), anti-pRB (554136; BD Biosciences — Pharmingen), anti–PSer780–phosphorylated pRB (anti–PSer780-ppRB) (sc-12901; Santa Cruz Biotechnology Inc.) raised against a pRB peptide phosphorylated on Ser-780, and anti–-actin (sc-7210; Santa Cruz Biotechnology Inc.) for loading control. Each blot is representative of 3 independent experiments. We have verified that immunoprecipitation with nonspecific IgG followed by Western blot analyses using anti-p16 or anti-CDK4 antibodies did not yield any signal (data not shown).
In vitro CDK4 activity was measured as previously described by Draetta et al. (60) using CDK4-immunoprecipitated SMC extracts (1 mg), a peptide corresponding to the carboxy terminal domain of pRB (sc-4112; Santa Cruz Biotechnology Inc.) as substrate, as well as 1 mg/ml casein (Sigma-Aldrich) in the assay buffer. After 5 minutes of preincubation at 30°C, the reaction was initiated by the addition of 1 μM adenosine triphosphate (ATP) and 5 μCi [-32P]ATP and continued at 30°C for 1 hour. Fifteen microliters of electrophoresis buffer (3x) was added to stop the reaction, and the samples were resolved on a 10% SDS-PAGE gel.
EMSAs.
Human PPAR and mouse RXR proteins were synthesized in vitro, and EMSAs were performed with double-stranded oligonucleotides (Table 2) end-labeled with [-32P]ATP as previously described (54). Competition experiments were performed by adding a 300-M excess of the indicated unlabeled oligonucleotides to the binding reaction just before the addition of the labeled probes. For supershift experiments, 0.2 μg anti-PPAR antibody (sc-9000; Santa Cruz Biotechnology Inc.) was preincubated for 20 minutes on ice in the binding buffer before addition of DNA probes.
ChIP assays.
ChIP experiments were performed as previously described (54). At 2 hours and 30 minutes before harvesting, hSMCs were washed with serum-free DMEM medium containing 0.2% bovine serum albumin and incubated in this medium with either fenofibric acid (250 μM) or Me2SO (vehicle) as control. Cells were sonicated and lysates immunoprecipitated using the indicated antibodies. Extracted DNA was PCR amplified using specific primers for the p16 and the -actin gene promoters (see Table 2) (54). An equal volume of nonprecipitated (input) genomic DNA was amplified as positive control. One-fifteenth (input) or one-fifth (precipitated DNA) of PCR products were separated on an ethidium-bromide–stained 2% agarose gel.
Animals.
All animal experiments were conducted with the approval of the Pasteur Institute review board, Lille, France, following guidelines for animal care and use (61). Studies were performed on 8- to 10-week-old PPAR+/+ and PPAR–/– mice with Sv/129 or C57BL/6J genetic backgrounds (a kind gift of F. Gonzalez, National Cancer Institute, NIH, Bethesda, Maryland, USA; ref. 62) as well as p16–/– mice with a C57BL/6J genetic background (30). One group of p16+/+ or p16–/– mice was treated orally with fenofibrate at 0.05% in chow diet (i.e., 50 mg/kg body weight/day) for 1 week before and 3 weeks after the surgical procedure. For the surgical procedures, the mice were anesthetized by intraperitoneal injection with a solution of domitor (0.68 mg/kg) and ketamin (67 mg/kg) (Orion Pharma) and allowed to recover on cotton following subcutaneous injection of the antidote antisedan at 1.7 mg/kg (Orion Pharma). At 3 weeks after injury, mice were anesthetized and then sacrificed by overdose of domitor/ketamin.
Carotid arterial injury model.
Intraluminal mechanical injury of the carotid arteries was performed by rupture of the internal elastic lamina and endothelial layer removal according to the method described by Carmeliet et al. (30). Briefly, the left internal carotid artery was exposed by blunt-end dissection, tied off distally, and looped proximally on the external branch with 5.0 silk black suture (Harvard apparatus unit; Ealing SA). A transverse section was made in the proximal portion of the external carotid artery, and a straight wire guide (C-SF-15-15; Cook) was passed toward the aortic arch and withdrawn 3 times with a rotating motion. After removal of the wire, the distal portion of the external carotid artery was tied off, and the skin incision was closed.
Tissue harvest, histology, and immunohistochemistry.
Tissue harvesting, fixation, embedding, and sectioning were performed as described by Carmeliet et al. (30), with slight modifications. The mice serving for morphometric analyses were first perfusion fixed with 4% phosphate-buffered formalin (pH 7.0) or methanol at physiological pressure. The left injured and corresponding right vessel segments were dissected and post-fixed overnight in 4% formalin or methanol placed between Whatman filter paper and then transferred to cold PBS. The vessel segments were then dehydrated in graded ethanol baths, cleared in toluene, and embedded in paraffin. Sections (6 μm) were created throughout the whole approximate 350-μm dissected fragment from the junction. Distance from junction was measured as the length between the junction of the external and internal branches of the left carotid artery (reference section at 0 μM, which appears right after the beginning of the injury) and the section position.
Sections at 50-μm increments starting from the junction were stained with Masson’s trichrome reagent (Sigma-Aldrich) according to the manufacturer's protocol. The sections were immunohistochemically stained with a mouse monoclonal anti–-actin antibody (1:100 dilution; Sigma-Aldrich) for SMC labeling, followed by detection with biotinylated secondary antibodies and streptavidin-horseradish peroxidase. Detection of PCNA was performed using a ready-to-use HRP-conjugated anti-PCNA monoclonal mouse antibody (DakoCytomation). Immunostains were visualized using the DAB substrate–chromogen system (DakoCytomation) for -actin and PCNA. Nuclear counterstaining was performed with hematoxylin. Images were captured by use of a JVC 3-CCD video camera.
Morphometric analyses.
Masson’s-stained sections were analyzed blindly by 2 observers using the computer-assisted Quips Image analysis system (Leica Mikroskopie und Systeme GmbH). Quantitative measurements of the neointima (i.e., luminal part limited by the internal elastic lamina) and media (i.e., part delineated by external and internal elastic laminas) area were made on complete cross-sectioned carotid arteries stained with Masson’s trichrome. Neointimal thickness was expressed as the ratio of the area of the neointima to the area of the media.
Statistics.
For in vitro assays, statistical differences were analyzed by the unpaired Student’s t test. For morphometric analyses, statistical differences in I/M ratio of PPAR–/– or fenofibrate-treated versus untreated PPAR+/+ mice were determined by ANOVA followed by Wilcoxon’s signed-rank analysis. Statistical significance was set at a value of P 0.05.
Acknowledgments
F. Gizard was supported by consecutive SFA-Fournier and Astra-Zeneca 1-year fellowships. C. Amant was supported by a Lefoulond-Delalande fellowship. Grants were provided by the Fondation Leducq. The authors thank M. Canavesi for the preparation of mSMC lines, P.J. Brown for the kind gift of GW7647, D. Monte and G. Peters for the gifts of plasmids, and F. Gonzalez for the gift of PPAR–/– mice. We thank also the members of the B. Staels Laboratory for valuable discussions.
Florence Gizard and Carole Amant contributed equally to this work.
Footnotes
Nonstandard abbreviations used: Ad, adenovirus; ADT, adenosine triphosphate; CDK, cyclin-dependent kinase; CDKI, CDK inhibitor; ChIP, chromatin IP; EMSA, electrophoretic mobility shift assay; GM, growth medium; hSMC, human SMC; I/M, intima/media; mSMC, mouse SMC; p16, p16INK4a; PCNA, proliferating cell nuclear antigen; ppRB, phosphorylated pRB; PPRE, PPAR-response element; pRB, retinoblastoma gene product; RXR, retinoid X receptor.
Conflict of interest: The authors have declared that no conflict of interest exists.
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2CHU d’Amiens and Faculte de Medecine, Universite Jules Verne de Picardie, Amiens, France.
3Department of Pharmacological Sciences, University of Milan, Milan, Italy.
4Division of Molecular Genetics and Centre for Biomedical Genetics, The Netherlands Cancer Institute, Amsterdam, The Netherlands.
Abstract
Vascular SMC proliferation is a crucial event in occlusive cardiovascular diseases. PPAR is a nuclear receptor controlling lipid metabolism and inflammation, but its role in the regulation of SMC growth remains to be established. Here, we show that PPAR controls SMC cell-cycle progression at the G1/S transition by targeting the cyclin-dependent kinase inhibitor and tumor suppressor p16INK4a (p16), resulting in an inhibition of retinoblastoma protein phosphorylation. PPAR activates p16 gene transcription by both binding to a canonical PPAR-response element and interacting with the transcription factor Sp1 at specific proximal Sp1-binding sites of the p16 promoter. In a carotid arterial–injury mouse model, p16 deficiency results in an enhanced SMC proliferation underlying intimal hyperplasia. Moreover, PPAR activation inhibits SMC growth in vivo, and this effect requires p16 expression. These results identify an unexpected role for p16 in SMC cell-cycle control and demonstrate that PPAR inhibits SMC proliferation through p16. Thus, the PPAR/p16 pathway may be a potential pharmacological target for the prevention of cardiovascular occlusive complications of atherosclerosis.
Introduction
Activation of vascular SMC proliferation is a key event in the development of atherosclerosis and its complications (1). In response to vascular injury, SMCs migrate from the media into the intimal layer of the arterial wall, where they proliferate and synthesize extracellular matrix, resulting in the formation of intimal hyperplasia. In this process, vascular SMCs undergo phenotypic changes from a differentiated and contractile state to a dedifferentiated and synthetic state (2). (Dys)regulation of SMC growth occurs thus during atherosclerotic plaque formation as part of a local inflammatory response in association with accumulation of lipids and fibrous connective tissue in the vascular wall (3). SMC proliferation is also the primary pathophysiological mechanism underlying complications of procedures used to treat atherosclerotic diseases, such as restenosis, a secondary occlusion of the arterial wall following transluminal angioplasty or stent implantation, and vein bypass graft failure (1).
As a key early event in these phenotypic changes, SMCs, which are in a quiescent state (G0) in normal uninjured vessels, transit through the G1 phase of the cell cycle and enter into the S phase to undergo replication (4). Cell-cycle progression is under the control of cyclin-dependent kinases (CDKs), which phosphorylate different specific target proteins through the 4 stages of the cell cycle (5). Notably, phosphorylation of the retinoblastoma gene product (pRB) by the specific G1 CDKs represents the critical checkpoint of G1/S transition (6). When underphosphorylated, pRB sequesters the E2F family transcription factors, which regulate genes encoding proteins required for S phase DNA synthesis. Phosphorylation of pRB releases E2F that permits the induction of E2F-dependent genes and therefore the irreversible induction of the mitosis process, after which cells are refractory to extracellular growth-inhibition signals. Thus, increased pRB phosphorylation correlates with the induction of SMC proliferation in injured vessels (1, 6). As CDKs are constitutively present in each phase of the cell cycle, CDK-mediated pRB phosphorylation is respectively activated or inhibited through timely interactions with expressed cyclins and CDK inhibitors (CDKIs) (7). Notably, the CDKI p16 is a tumor suppressor transcriptionally regulated by pRB, which plays a key role in the control of pRB phosphorylation and G1/S cell-cycle progression (8-11).
PPAR is a nuclear receptor that, upon ligand activation, regulates transcription after dimerizing with the retinoid X receptor (RXR) and DNA-binding to PPAR-response elements (PPREs) within the regulatory regions of target genes (12). PPREs usually consist of a direct repeat of the hexanucleotide AGGTCA sequence separated by 1 or 2 nucleotides (DR1 or DR2) (12). Furthermore, PPAR can also negatively interfere with proinflammatory signaling pathways by a mechanism termed "transrepression" (13). PPAR is activated by natural ligands such as fatty acids and derivatives (13). Notably, natural eicosanoids derived from arachidonic acid via the lipoxygenase pathway as well as oxidized phospholipids activate PPAR (13). Moreover, fibrates (gemfibrozil, bezafibrate, ciprofibrate, and fenofibrate), synthetic drugs initially used to treat lipid disorders (14), act as PPAR ligands (15).
Fibrates have been shown to decrease morbidity and mortality of cardiovascular diseases (16) and impede the progression of coronary atherosclerotic lesions when administered to postinfarction or diabetic patients (17-19). PPAR mediates their systemic metabolic effects by regulating genes encoding proteins involved in lipid and lipoprotein metabolism (20). Recently, the functions of PPAR have been extended to a regulatory action on cholesterol homeostasis and vascular inflammation acting directly at the level of the vascular wall, which could confer additional protective effects in the prevention of atherogenesis (13). Importantly, PPAR is also highly expressed in SMCs isolated from vascular tissues and from atherosclerotic lesions (21-23), where it controls IL-1–induced secretion of IL-6 and production of prostaglandins by inhibiting IL-6 and COX2 gene transcription. However, although recent studies have indicated that fibrates may regulate SMC proliferation in vitro (21, 24), a role for PPAR in SMC cell-cycle control has not yet been explored. In the present study, we demonstrate a role for PPAR in the control of SMC G1/S transition. Moreover, we identify the CDKI p16 as an important controller of vascular SMC proliferation and show that p16 mediates the effects of PPAR on SMC proliferation both in vitro and in vivo.
Results
PPAR inhibits SMC proliferation by controlling cell-cycle progression into the S phase.
Previous studies have indicated that certain fibrates may regulate human SMC (hSMC) proliferation in vitro (21, 24). To further analyze the effect of PPAR activation on SMC growth, the influence of a novel generation subtype-specific PPAR agonist, GW7647, which presents a high affinity (EC50 = 6 nM) and specificity (200-fold selectivity over PPAR and PPAR&Dgr;) (25), was first tested on hSMCs resuming proliferation in normal growth medium (GM) after G0/G1-phase synchronization (Figure 1A). Whereas vehicle-treated cells displayed a growth rate comparable to that of untreated cells (not shown) with a doubling time of approximately 24 hours, GW7647 treatment inhibited proliferation in a dose-dependent manner. To examine which stage(s) of the cell-cycle is (are) regulated by GW7647, the cell-cycle distribution of hSMCs resuming growth in GM was analyzed by flow cytometry analysis after 48 hours of treatment with GW7647 or vehicle. As shown in Table 1, GW7647 treatment increased and decreased the proportion of hSMCs in the G0/G1 and S phases, respectively.
To further characterize the exact role of PPAR in SMC cell-cycle control, aortic mouse SMCs (mSMCs) were isolated from PPAR–/– and PPAR+/+ mice. Interestingly, PPAR–/– mSMCs isolated from both Sv/129 (Figure 1B) and C57BL/6J (Supplemental Figure 1; supplemental material available online with this article; doi:10.1172/JCI22756DS1 http://dx.doi.org/10.1172/JCI22756DS1) genetic backgrounds proliferated at a much higher rate than corresponding PPAR+/+ mSMCs. Moreover, incubation of mSMCs with Wy14,643, a specific murine PPAR agonist, inhibited cell proliferation of PPAR+/+, but not PPAR–/– mSMCs (Figure 1B). Similarly to hSMCs, PPAR activation of mSMCs (assessed after 48-hour treatment in GM) was associated with a higher cell percentage in G0/G1 and a concomitant decrease of cell percentage in the S phase (Table 1).
Concomitantly, phosphorylation of pRB, the major checkpoint of G1/S progression (5), was reduced in GW7647-treated hSMCs, an effect which was even more pronounced upon infection with adenovirus-PPAR (Ad-PPAR) (Figure 1C). Altogether, these results indicate that PPAR activation inhibits both human and mouse vascular SMC proliferation by interfering with G1/S cell-cycle progression.
PPAR induces transcription of the G1/S transition regulator and tumor suppressor p16.
Since PPAR is a nuclear receptor acting at the transcriptional level (12) and since the tumor suppressor p16 is a transcriptionally regulated CDKI controlling G1/S cell-cycle progression functionally related to in-cell pRB expression and phosphorylation levels (8-10), we hypothesised that p16 could be a PPAR target gene. Indeed, in both mSMCs and hSMCs, p16 mRNA levels were increased upon PPAR agonist incubation (Figure 2A). The induction of p16 mRNA occurred in a PPAR-dependent manner and was already evident within 4 hours of treatment (Figure 2A). In hSMCs, the increase of p16 mRNA levels upon GW7647 treatment was more pronounced in Ad-PPAR–infected hSMCs (Figure 2A, upper panel). Moreover, Wy14,643 treatment induced p16 expression in PPAR+/+, but not in PPAR–/– mSMCs (Figure 2A, lower panel). Interestingly, p16 mRNA levels were decreased by approximately two-thirds in PPAR–/– compared with PPAR+/+ mSMCs.
Previously, a pRB-responsive site and proximal Sp1-binding sites (denoted A, B, and C) have been shown to be involved in the basal activity of the p16 promoter (11, 26). Here, the presence of a putative degenerated DR1 PPRE sequence (AGGAGACAGGACA) at position –1023 to –1011 (denoted the p16DR1 site) was also revealed by computer-assisted analysis. To investigate the putative function of these elements in mediating the response to ligand-activated PPAR, reporter constructs driven by different p16 promoter fragments were cotransfected with or without the pSG5-PPAR expression plasmid, and cells were subsequently treated with the PPAR agonists fenofibric acid or GW7647. Cotransfection of PPAR followed by ligand activation significantly induced activity of the –3.0-kb p16 gene promoter, indicating that p16 is a direct PPAR target gene (Figure 2B). PPAR-dependent p16 promoter activation was maintained upon 5' deletion to –1.0 kb. Further nucleotide deletions to –869 and –703 bp, as well as mutation of p16DR1, resulted in a significant, albeit incomplete reduction of p16 promoter activation by PPAR. Finally, deletion to –343 bp completely abolished activation of the p16 promoter by fenofibric acid–activated PPAR (Figure 2B). Identical results were obtained in cells treated with GW7647 (250 nM) (data not shown). These observations indicate that both p16DR1 and Sp1A–C elements are involved in PPAR-dependent p16 promoter activation, with the DR1 element mediating the majority of the response.
To determine whether PPAR physically binds to the p16DR1 site, electrophoretic mobility shift assays (EMSAs) were performed using in vitro–produced PPAR and RXR proteins and double-stranded [32P]-labeled oligonucleotides (Table 2 and Figure 2C). Whereas neither RXR nor PPAR alone bound to the p16DR1 probe, a clear shift was observed upon incubation with both RXR and PPAR. This complex was supershifted by the anti-PPAR antibody and prevented by an excess of unlabeled wild-type, but not mutated, p16DR1 probe. Moreover, the mutated p16DR1 probe did not bind RXR/PPAR (Figure 2C). These results demonstrate that the RXR/PPAR heterodimer specifically binds in vitro to the p16DR1 site located at –1023 pb. In contrast, no PPAR/DNA complex formation was detected in EMSAs performed with oligonucleotides overlapping the proximal Sp1A- and B-binding sites (data not shown).
To investigate the in vivo occupancy of these p16DR1 and Sp1-binding sites by activated PPAR, chromatin IP (ChIP) assays were performed on DNA from hSMCs treated or not treated with fenofibric acid using anti-Sp1 and anti-PPAR antibodies (Figure 2D). In accordance with the EMSA results, the region spanning the p16DR1 sequence was immunoprecipitated by the anti-PPAR antibody, an effect which was enhanced by fenofibric acid treatment. In addition, in agreement with results of Myhnen et al. (26), genomic p16 promoter DNA regions encompassing the Sp1-binding elements were immunoprecipitated by the anti-Sp1 antibody. Interestingly, similarly to with the DR1 element, the region encompassing the Sp1A and C sites also bound PPAR in a PPAR ligand–dependent manner (Figure 2D). In contrast, no PPAR binding was observed on the Sp1B site. As negative controls, PCR amplification using primers covering a region localized just downstream of the –1023DR1 site or a -actin genomic fragment did not yield a significant signal, thus demonstrating the specificity of PPAR immunoprecipitation and PCR amplification. Taken together, these results indicate that PPAR activates the p16 promoter through direct DNA binding to the p16DR1 site as well as tethering with Sp1 to the proximal Sp1A–C response element.
Induction of p16 following PPAR activation, resulting in the inhibition of CDK4-mediated pRB phosphorylation, is required for the SMC growth-inhibitory activity of PPAR.
Next, it was determined whether the upregulation of p16 gene expression by PPAR results in an increase of p16 protein abundance and in-cell activity. Western blot analysis of hSMC whole-cell protein extracts revealed that PPAR overexpression or GW7647 treatment enhanced p16 protein levels in hSMCs, and the greatest increase was observed when cells were both infected with Ad-PPAR and treated with GW7647 (Figure 3A).
Initially, p16 was identified by virtue of its ability to inhibit G1/S cell-cycle progression by binding to CDK4, thus preventing CDK4 association with D-type cyclins and further initiation of pRB phosphorylation by the cyclin D/CDK4 complexes (8, 9, 27). To verify that PPAR controls p16 function, it was further determined whether the induction of p16 following PPAR activation affected p16/CDK4 interaction and CDK4 activity. To this end, in-cell pRB phosphorylation by CDK4, determined using an anti–PSer780-ppRB antibody (28), and in vitro CDK4-associated activity using pRB as substrate were measured after PPAR overexpression and/or activation with GW7647 in hSMCs (Figure 3, B and C, upper panel). In fact, concurring with the decrease of the phosphorylated/nonphosphorylated pRB ratio following PPAR activation (Figure 1C), both analyses indicated that treatment with the PPAR ligand decreases pRB phosphorylation by CDK4 and that this effect is enhanced by PPAR overexpression. Moreover, the levels of p16 associated with CDK4 were increased in hSMCs infected with Ad-PPAR or treated with GW7647, and this interaction was further enhanced in Ad-PPAR–infected GW7647-treated hSMCs (Figure 3C, middle panel). Besides, in-cell p16 protein levels and CDK4-dependent pRB phosphorylation respectively increased and decreased in PPAR+/+ but not in PPAR–/– mSMCs following Wy14,643 treatment (Figure 3D).
Finally, the role of p16 in mediating the growth-inhibitory effects of PPAR activation was analyzed in vitro using SMCs isolated from p16–/– mice (29). Strikingly, primary mSMCs isolated from p16–/– mice proliferated at a much higher rate than p16+/+ mSMCs (Figure 4A). Moreover, p16–/– mSMCs did not respond to Wy14,643 treatment. Accordingly, the ratio of cell percentage in S compared with G1 phases (data not shown) as well as in-cell CDK4-mediated pRB phosphorylation (Figure 4B) were enhanced in p16–/– mSMCs and not affected by Wy14,643 treatment. Altogether, these data indicate that the G1 cell-cycle arrest induced by ligand-activated PPAR occurs via inhibition of the activity of G1 CDKs, such as CDK4, due to an increased recruitment of p16.
PPAR activation inhibits in vivo SMC proliferation upon carotid arterial injury by increasing p16 gene expression.
The in vivo role of PPAR in the control of SMC proliferation was next examined by using a mechanical carotid artery injury mouse model (30) in Sv/129 mice, a strain highly susceptible to intimal hyperplasia (31). The neointimal thickness in response to injury was determined by measuring the intima/media (I/M) area ratio throughout the entire injured segment starting from the junction between the internal and external left carotid branches. In agreement with previous reports (31, 32), injury induced a marked increase in intimal hyperplasia in wild-type mice as observed 3 weeks after injury (Figure 5, A and B). Interestingly, the I/M ratio was markedly enhanced in PPAR–/– mice,whereas it was drastically decreased in PPAR+/+ mice treated with fenofibrate, whose action in mice requires PPAR expression (33, 34) (Figure 5, A and B). As previously reported (2, 21, 30), SMCs, identified by -actin staining, were abundantly present in the media of uninjured carotids and constituted the major cell type underlying neointimal development in response to injury in this model (Figure 5, A–C, see PPAR+/+). Moreover, in agreement with the I/M ratio results, neointimal SMC and proliferating cell nuclear antigen–staining (PCNA-staining) were increased in carotid sections of PPAR–/– mice, and diminished in fenofibrate-treated mice compared with untreated wild-type mice (Figure 5, B and C). In agreement with the in vitro proliferation assays performed in mSMCs (Supplemental Figure 1), PPAR deficiency also provoked hyperplasia in C57BL/6J mice (Supplemental Figure 2), a strain reported to be more resistant to hyperplasia formation after carotid injury (31, 32).
Furthermore, p16 gene expression was measured in the uninjured carotid segment, which contains mainly SMCs in a quiescent state (1, 4) (Figure 5C), as well as in carotid segments at different times after injury (Figure 5D). Strikingly, p16 mRNA levels were induced after response to injury in PPAR+/+ but not in PPAR–/– mice, an effect that was further enhanced following fenofibrate treatment. A maximal PPAR-dependent induction of p16 mRNA levels was reached within 24 hours and remained elevated 3 weeks after injury. Thus, these data indicate that PPAR activation negatively controls neointima formation in response to injury, possibly via the induction of p16 gene expression in injured vessels.
Finally, the in vivo role of p16 in mediating the growth-inhibitory effects of PPAR activation were further analyzed using p16–/– mice (29). It is noteworthy that, similarly to what occurred in PPAR–/– mice (Figure 5 and Supplemental Figure 2), intimal hyperplasia was drastically increased in p16–/– mice (Figure 6). An extended neointima formation was observed over the entire lesion segment, which was accompanied by an increase in SMC abundance. Strikingly, fenofibrate treatment did not influence the I/M ratio in these p16–/– mice (Figure 6A). Therefore, p16 is required for the inhibitory effects of PPAR activation on vascular SMC growth underlying intimal hyperplasia development.
Discussion
The cardiovascular protective effects of fibrates were initially mainly attributed to a systemic action of PPAR on atherogenic dyslipidemia, characterized by a lowering of triglyceride levels and an increased concentration of plasma HDL-cholesterol concentrations (13). However, a number of recent clinical studies, including the Bezafibrate Coronary Atherosclerosis Intervention Trial (BECAIT; ref. 19), the Lopid Coronary Angiography Trial (LOCAT; ref. 17), and the Diabetes Atherosclerosis Intervention Study (DAIS; ref. 17), have revealed that fibrates inhibit atheromatous plaque progression via effects possibly independent or in addition to their systemic action. These observations are in line with results from in vivo studies performed in rabbits, which indicated that fibrate treatment can inhibit atherosclerosis without affecting plasma lipid profiles (35, 36). Moreover, in the secondary prevention Veterans Administration–HDL-Cholesterol Intervention Trial (VA-HIT), a decreased incidence of cardiovascular events following fibrate treatment was observed, which could not be attributed only to the effects of drug treatment on HDL levels (37). Concurring with these clinical data, compelling mechanistic evidence has recently emerged that PPAR controls cholesterol efflux and the inflammatory response in vessel wall cells (13), highlighting a direct regulatory role of PPAR on vessel wall function. However, despite the crucial role of SMC proliferation in the development of occlusive vascular diseases, the function of PPAR in the control of vascular SMC cell-cycle progression was not yet established. In the present study, we show that PPAR activation inhibits G1/S cell-cycle progression of both human and mouse primary SMCs in vitro, an effect attributed to the induction of the tumor suppressor gene p16. Upregulation of p16 by PPAR agonists results in the sequestration of CDK4 and the consequent decrease of CDK4-mediated pRB phosphorylation, thus providing a mechanism for the SMC G1 cell-cycle arrest induced by fibrates. The in vivo role of PPAR in the control of SMC proliferation through the induction of p16 gene expression was further demonstrated by studying the response to mechanical carotid artery injury (30). This carotid injury model is characterized by neointima formation due to an excessive accumulation of SMCs and deposition of extracellular matrix in the intimal layer of the vessel wall, a response which has been proposed to appropriately reflect the "in-stent" restenosis observed in humans (30).
In the search for the molecular mechanisms involved in the p16-dependent growth-inhibitory effects of PPAR on SMCs, PPAR activation was found to upregulate p16 promoter activity, thus identifying for the first time a molecular mechanism via which PPAR directly interferes with cell-cycle progression. It is noteworthy that numerous common disease predisposition factors and pathways between cancer and vascular remodeling pathologies have been identified, pointing to similarities between the development of both diseases (38). It has been suggested that both vascular diseases and cancer similarly develop from a clonal proliferation of altered cells at the sites of local tissue injury, inflammation, and/or genomic alterations (38). Similar to tumor development, the induction of intimal hyperplasia in response to injury involves the conversion of SMCs from a medial nonproliferating differentiated phenotype into a proliferating phenotype associated with pRB hyperphosphorylation (1). However, the molecular mechanisms underlying the relationship between these diseases remain to be characterized. Interestingly, the CDKI and tumor suppressor p27Kip1 (p27), whose protein degradation is prevented by PPAR activators (39), has been shown to inhibit the development of vascular diseases when transferred using an adenoviral vector into balloon-injured porcine arteries (40) as well as through bone marrow–derived immune cells (41). In this study we now report an endogenous regulatory role for the CDKI p16 as SMC growth inhibitor, which furthermore directly mediates the response to fibrates. In fact, p16 expression is correlated to both pRB phosphorylation and expression levels and is required for the negative feedback of G1/S transition (11, 27). Several studies have established that loss of p16 function can be an early event in tumor progression (42, 43) and that it is associated with tumor development (29, 44, 45). Thus, p16 promoter activity is currently utilized as a prognosis factor for several cancers, such as acute lymphoblastic leukemia, lymphoma, lung carcinoma, and melanoma (45). Here, we identify, for what we believe is the first time, a role for this CDKI, so far studied in cancer pathology, in vascular SMC proliferation underlying occlusive cardiovascular diseases such as restenosis (1).
Strikingly, PPAR deficiency and p16 deficiency both acted as molecular switches inducing SMC proliferation in vitro and in vivo (Figures 1 and 4 and Supplemental Figures 1 and 2). Interestingly, the induction of p16 expression alone and in response to fenofibrate treatment was much higher in response to injury than in healthy carotid arteries. Moreover, no significant morphological differences were observed in healthy carotid arteries isolated from untreated PPAR+/+ compared with PPAR–/– or fenofibrate-treated mice. Among the genes of the INK4 family, the CDKI p16 is specifically induced in response to oncogenic stress when it acts as a negative retrocontrol of increased pRB phosphorylation (27). In parallel, an important role for PPAR in the response to stress has been demonstrated previously upon fasting (46) or long-term high-fat feeding (47) as well as in the response to inflammatory factors such as leukotriene B4 (48), lipopolysaccharide (LPS), or IL-1 (49). Thus, our results demonstrating that both PPAR and p16 proteins are activated and function locally at the vascular artery injury site further extend these observations of a functional role for PPAR and p16 primarily in the response to stress.
Recently, substantial progress has been made in the prevention of restenosis, notably with the development of stents coated with antimitotic drugs, such as rapamycin (50). However, a potential risk of late thrombus formation associated with incomplete reendothelialization after surgery with these coated stents exists (50). It is noteworthy that, in our mouse model of restenosis, reendothelialization was completed 3 weeks after injury for all studied groups (Supplemental Figure 3). Moreover, thiazolidinedione PPAR activators —which have previously been shown to control vascular SMC proliferation (39, 51) — were recently shown to reduce coronary in-stent restenosis in type 2 diabetic patients (52, 53). The present report indicates a role for the PPAR/p16 pathway in vascular SMC proliferation in vitro and in a mouse model of restenosis. Further clinical studies are thus warranted to assess whether pharmacological activation of PPAR could provide a novel therapeutic strategy to prevent restenosis in treatments using either systemic administration or coated stents.
Methods
Reagents.
Wy14,643 and fenofibric acid were purchased from Sigma-Aldrich and Laboratoires Fournier SA, respectively. GW7647 (25) was kindly provided by P. J. Brown from GlaxoSmithKline (Research Triangle Park, North Carolina, USA).
Reverse transcription and quantitative PCR.
RNA extraction, reverse transcription, and quantification of mRNA levels were performed using an MX 4000 apparatus (Stratagene) as previously described (54). 28S RNA was measured as internal control. For in vitro assays, values are expressed as means ± SD of a representative experiment performed in triplicate (for a total of 3 samples measured).
Plasmids and Ad generation.
The Sp1 expression plasmid cloned in pcDNA3 was a gift from D. Monte (Institut de Biologie de Lille, Lille, France); the p16 reporter pGL2 (Promega) constructs were provided by G. Peters (Imperial Cancer Research Fund Laboratories, London, United Kingdom) (11). Mutations of the –1023 p16DR1 were introduced using the QuikChange site-directed mutagenesis kit (Stratagene) and the oligonucleotide mutated p16DR1 (p16DR1mt) (Table 2). The nucleotide positions are indicated relative to the ATG site. Recombinant Ad-GFP and Ad-PPAR were obtained by homologous recombination in Escherichia coli (55) after insertion of the cDNAs into the pAdCMV2 vector (Qbiogene). Viral stocks were created and titrated as previously described (56). SMCs (0.5 x 106) were infected at an input multiplicity of 100 virus particles/cell by adding virus stocks directly to the culture medium. Efficiency of hSMC infection was between 75–85%, as assessed by FACS analysis with the Ad-GFP vector.
Cell culture assays.
Human cervical HeLa cells and primary human aortic SMCs (hSMCs) were obtained from ATCC and BioWhittaker Europe, respectively. HeLa cells were cultured in DMEM medium (Invitrogen Corp.) supplemented with 10% FCS (HyClone), and L-glutamine (2 mM) as well as the antibiotics penicillin (50 U/ml) and streptomycin (55 μg/ml) (Gibco; Invitrogen Corp.). hSMCs were cultured using the SMC Growth Medium 2 Bullet Kit , purchased from BioWhittaker Inc. Mouse primary aortic SMCs (mSMCs) were prepared from aortas of 8- to 12-week-old PPAR+/+ or PPAR–/– mice on Sv/129 or C57BL/6J genetic backgrounds or of p16+/+ or p16–/– mice on a C57BL/6J genetic background as described (57) and were studied between the third and twelfth passages. mSMCs were cultured in DMEM supplemented with 10% FCS and antibiotics. In all assays, cell synchronization in the G0/G1 phase was induced by serum starvation for at least 24 hours in medium containing 0.4% FCS. To reinduce cell proliferation, cells were reincubated in routine GM (time 0 or T0 of the experiments). At T0, mSMCs were treated with the indicated PPAR ligand or an identical volume of Me2SO (vehicle). hSMCs were infected overnight and subsequently treated with PPAR ligands or vehicle as indicated. Three independent preparations from different mice were studied for each primary SMC type and resulted in similar results.
For cell proliferation assays, cellular DNA content was determined using fluorochrome 3,5-diaminobenzoicacid free acid (DABA) as described previously (58). Flow cytometry analysis profiles were determined by propidium iodide staining of nuclear DNA as described (59).
For transient transfection assays, HeLa cells in 24-well plates at an initial density of 1.5 x 104 per well were transfected with ExGen 500 (Euromedex) at a ratio of 4 μl ExGen 500/0.5 μg of DNA. The indicated pGL2 reporter constructs (100 ng) were cotransfected with the indicated expression vectors and the internal control Renilla luciferase reporter plasmid pRL-null (10 ng). The amount of plasmids in the transfection mixture was equalized to 0.5 μg by adding pcDNA3 vector. After 12 hours, cells were incubated for an additional 10 hours in fresh medium supplemented with fenofibric acid (100 μM) or Me2SO (vehicle). Luciferase activity was assessed with the Dual-Luciferase Reporter Assay System (Promega) as previously described (54). Transfections were carried out in triplicate, and each experiment was repeated at least 3 times.
Protein assays.
Cells were washed twice with ice-cold PBS and then collected in ice-cold RIPA buffer (1x PBS, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS, phosphatase and protease inhibitors) or kinase lysis buffer (50 mM Tris, pH 8.0, 0,1% IGEPAL; Sigma-Aldrich). Lysates were incubated for 30 minutes on ice, and cell debris was removed by centrifugation. Extracts were then aliquoted and stored at –80°C.
For coimmunoprecipitation assays, whole-cell lysates were preincubated with 1.0 μg nonimmune rabbit IgG and 20 μl of protein A-Sepharose beads (Amersham Biosciences) during 30 minutes at 4°C and centrifuged. Cleared lysates (1 mg) were immunoprecipitated for 1 hour at 4°C with the anti-CDK4 antibody (RB-016, Microm Microtech France) and then incubated overnight at 4°C with protein A-Sepharose. The immunoprecipitated proteins were washed with lysis buffer and resuspended in electrophoresis sample buffer.
Samples of immunoprecipitated or total proteins (20–30 μg) were analyzed by Western blot using the following primary antibodies: anti-CDK4 (RB-003; Microm Microtech France), anti-p16 (554079 for hSMCs, BD Biosciences — Pharmingen; and MS-887 for mSMCs, Microm Microtech France), anti-pRB (554136; BD Biosciences — Pharmingen), anti–PSer780–phosphorylated pRB (anti–PSer780-ppRB) (sc-12901; Santa Cruz Biotechnology Inc.) raised against a pRB peptide phosphorylated on Ser-780, and anti–-actin (sc-7210; Santa Cruz Biotechnology Inc.) for loading control. Each blot is representative of 3 independent experiments. We have verified that immunoprecipitation with nonspecific IgG followed by Western blot analyses using anti-p16 or anti-CDK4 antibodies did not yield any signal (data not shown).
In vitro CDK4 activity was measured as previously described by Draetta et al. (60) using CDK4-immunoprecipitated SMC extracts (1 mg), a peptide corresponding to the carboxy terminal domain of pRB (sc-4112; Santa Cruz Biotechnology Inc.) as substrate, as well as 1 mg/ml casein (Sigma-Aldrich) in the assay buffer. After 5 minutes of preincubation at 30°C, the reaction was initiated by the addition of 1 μM adenosine triphosphate (ATP) and 5 μCi [-32P]ATP and continued at 30°C for 1 hour. Fifteen microliters of electrophoresis buffer (3x) was added to stop the reaction, and the samples were resolved on a 10% SDS-PAGE gel.
EMSAs.
Human PPAR and mouse RXR proteins were synthesized in vitro, and EMSAs were performed with double-stranded oligonucleotides (Table 2) end-labeled with [-32P]ATP as previously described (54). Competition experiments were performed by adding a 300-M excess of the indicated unlabeled oligonucleotides to the binding reaction just before the addition of the labeled probes. For supershift experiments, 0.2 μg anti-PPAR antibody (sc-9000; Santa Cruz Biotechnology Inc.) was preincubated for 20 minutes on ice in the binding buffer before addition of DNA probes.
ChIP assays.
ChIP experiments were performed as previously described (54). At 2 hours and 30 minutes before harvesting, hSMCs were washed with serum-free DMEM medium containing 0.2% bovine serum albumin and incubated in this medium with either fenofibric acid (250 μM) or Me2SO (vehicle) as control. Cells were sonicated and lysates immunoprecipitated using the indicated antibodies. Extracted DNA was PCR amplified using specific primers for the p16 and the -actin gene promoters (see Table 2) (54). An equal volume of nonprecipitated (input) genomic DNA was amplified as positive control. One-fifteenth (input) or one-fifth (precipitated DNA) of PCR products were separated on an ethidium-bromide–stained 2% agarose gel.
Animals.
All animal experiments were conducted with the approval of the Pasteur Institute review board, Lille, France, following guidelines for animal care and use (61). Studies were performed on 8- to 10-week-old PPAR+/+ and PPAR–/– mice with Sv/129 or C57BL/6J genetic backgrounds (a kind gift of F. Gonzalez, National Cancer Institute, NIH, Bethesda, Maryland, USA; ref. 62) as well as p16–/– mice with a C57BL/6J genetic background (30). One group of p16+/+ or p16–/– mice was treated orally with fenofibrate at 0.05% in chow diet (i.e., 50 mg/kg body weight/day) for 1 week before and 3 weeks after the surgical procedure. For the surgical procedures, the mice were anesthetized by intraperitoneal injection with a solution of domitor (0.68 mg/kg) and ketamin (67 mg/kg) (Orion Pharma) and allowed to recover on cotton following subcutaneous injection of the antidote antisedan at 1.7 mg/kg (Orion Pharma). At 3 weeks after injury, mice were anesthetized and then sacrificed by overdose of domitor/ketamin.
Carotid arterial injury model.
Intraluminal mechanical injury of the carotid arteries was performed by rupture of the internal elastic lamina and endothelial layer removal according to the method described by Carmeliet et al. (30). Briefly, the left internal carotid artery was exposed by blunt-end dissection, tied off distally, and looped proximally on the external branch with 5.0 silk black suture (Harvard apparatus unit; Ealing SA). A transverse section was made in the proximal portion of the external carotid artery, and a straight wire guide (C-SF-15-15; Cook) was passed toward the aortic arch and withdrawn 3 times with a rotating motion. After removal of the wire, the distal portion of the external carotid artery was tied off, and the skin incision was closed.
Tissue harvest, histology, and immunohistochemistry.
Tissue harvesting, fixation, embedding, and sectioning were performed as described by Carmeliet et al. (30), with slight modifications. The mice serving for morphometric analyses were first perfusion fixed with 4% phosphate-buffered formalin (pH 7.0) or methanol at physiological pressure. The left injured and corresponding right vessel segments were dissected and post-fixed overnight in 4% formalin or methanol placed between Whatman filter paper and then transferred to cold PBS. The vessel segments were then dehydrated in graded ethanol baths, cleared in toluene, and embedded in paraffin. Sections (6 μm) were created throughout the whole approximate 350-μm dissected fragment from the junction. Distance from junction was measured as the length between the junction of the external and internal branches of the left carotid artery (reference section at 0 μM, which appears right after the beginning of the injury) and the section position.
Sections at 50-μm increments starting from the junction were stained with Masson’s trichrome reagent (Sigma-Aldrich) according to the manufacturer's protocol. The sections were immunohistochemically stained with a mouse monoclonal anti–-actin antibody (1:100 dilution; Sigma-Aldrich) for SMC labeling, followed by detection with biotinylated secondary antibodies and streptavidin-horseradish peroxidase. Detection of PCNA was performed using a ready-to-use HRP-conjugated anti-PCNA monoclonal mouse antibody (DakoCytomation). Immunostains were visualized using the DAB substrate–chromogen system (DakoCytomation) for -actin and PCNA. Nuclear counterstaining was performed with hematoxylin. Images were captured by use of a JVC 3-CCD video camera.
Morphometric analyses.
Masson’s-stained sections were analyzed blindly by 2 observers using the computer-assisted Quips Image analysis system (Leica Mikroskopie und Systeme GmbH). Quantitative measurements of the neointima (i.e., luminal part limited by the internal elastic lamina) and media (i.e., part delineated by external and internal elastic laminas) area were made on complete cross-sectioned carotid arteries stained with Masson’s trichrome. Neointimal thickness was expressed as the ratio of the area of the neointima to the area of the media.
Statistics.
For in vitro assays, statistical differences were analyzed by the unpaired Student’s t test. For morphometric analyses, statistical differences in I/M ratio of PPAR–/– or fenofibrate-treated versus untreated PPAR+/+ mice were determined by ANOVA followed by Wilcoxon’s signed-rank analysis. Statistical significance was set at a value of P 0.05.
Acknowledgments
F. Gizard was supported by consecutive SFA-Fournier and Astra-Zeneca 1-year fellowships. C. Amant was supported by a Lefoulond-Delalande fellowship. Grants were provided by the Fondation Leducq. The authors thank M. Canavesi for the preparation of mSMC lines, P.J. Brown for the kind gift of GW7647, D. Monte and G. Peters for the gifts of plasmids, and F. Gonzalez for the gift of PPAR–/– mice. We thank also the members of the B. Staels Laboratory for valuable discussions.
Florence Gizard and Carole Amant contributed equally to this work.
Footnotes
Nonstandard abbreviations used: Ad, adenovirus; ADT, adenosine triphosphate; CDK, cyclin-dependent kinase; CDKI, CDK inhibitor; ChIP, chromatin IP; EMSA, electrophoretic mobility shift assay; GM, growth medium; hSMC, human SMC; I/M, intima/media; mSMC, mouse SMC; p16, p16INK4a; PCNA, proliferating cell nuclear antigen; ppRB, phosphorylated pRB; PPRE, PPAR-response element; pRB, retinoblastoma gene product; RXR, retinoid X receptor.
Conflict of interest: The authors have declared that no conflict of interest exists.
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