PPARα inhibits vascular smooth muscle cell proliferation underlying intimal hyperplasia by inducing the tumor suppressor p16^INK4a

Vascular SMC proliferation is a crucial event in occlusive cardiovascular diseases. PPARα 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 PPARα controls SMC cell-cycle progression at the G₁/S transition by targeting the cyclin-dependent kinase inhibitor and tumor suppressor p16^INK4a (p16), resulting in an inhibition of retinoblastoma protein phosphorylation. PPARα 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, PPARα 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 PPARα inhibits SMC proliferation through p16. Thus, the PPARα/p16 pathway may be a potential pharmacological target for the prevention of cardiovascular occlusive complications of atherosclerosis.

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).

PPARα 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, PPARα can also negatively interfere with proinflammatory signaling pathways by a mechanism termed “transrepression” (13). PPARα 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 PPARα (13). Moreover, fibrates (gemfibrozil, bezafibrate, ciprofibrate, and fenofibrate), synthetic drugs initially used to treat lipid disorders (14), act as PPARα 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). PPARα mediates their systemic metabolic effects by regulating genes encoding proteins involved in lipid and lipoprotein metabolism (20). Recently, the functions of PPARα 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, PPARα 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 PPARα in SMC cell-cycle control has not yet been explored. In the present study, we demonstrate a role for PPARα 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 PPARα on SMC proliferation both in vitro and in vivo.

PPAR α 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 PPARα activation on SMC growth, the influence of a novel generation subtype-specific PPARα agonist, GW7647, which presents a high affinity (EC50 = 6 nM) and specificity (∼200-fold selectivity over PPARγ 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.

Figure 1

PPARα inhibits SMC G₁/S progression. (A and B) G₁-arrested hSMCs (A) and PPARα^–/– or PPARα^+/+ Sv/129 mSMCs (B) resumed growth at T0 in normal GM. The indicated concentrations of GW7647, Wy14,643 (10 μM), or vehicle (Me₂SO) were added in the normal GM. Cells were harvested at the indicated times for measure of DNA content (expressed as arbitrary units). Values are the mean ± SEM of triplicate points from a single experiment (n = 3/time point), which was repeated with 3 different cell preparations of primary SMCs with similar results. *P ≤ 0.05; ***P ≤ 0.001 vs. vehicle-treated hSMCs (A). Values in B followed by different symbols are statistically significantly different from each other. (C) Western blot analysis of in-cell pRB phosphorylation was performed using an anti-pRB antibody recognizing all species of the Rb gene product on protein extracts (30 μg) from Ad-GFP or Ad-PPARα–infected hSMCs and treated with vehicle or GW7647 (600 nM) for 10 hours.

Table 1

PPAR α negatively controls SMC G₁/S cell-cycle progression

To further characterize the exact role of PPARα in SMC cell-cycle control, aortic mouse SMCs (mSMCs) were isolated from PPARα^–/– and PPARα^+/+ mice. Interestingly, PPARα^–/– 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) genetic backgrounds proliferated at a much higher rate than corresponding PPARα^+/+ mSMCs. Moreover, incubation of mSMCs with Wy14,643, a specific murine PPARα agonist, inhibited cell proliferation of PPARα^+/+, but not PPARα^–/– mSMCs (Figure 1B). Similarly to hSMCs, PPARα 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-PPARα (Ad-PPARα) (Figure 1C). Altogether, these results indicate that PPARα activation inhibits both human and mouse vascular SMC proliferation by interfering with G₁/S cell-cycle progression.

PPAR α induces transcription of the G₁/S transition regulator and tumor suppressor p16. Since PPARα 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 PPARα target gene. Indeed, in both mSMCs and hSMCs, p16 mRNA levels were increased upon PPARα agonist incubation (Figure 2A). The induction of p16 mRNA occurred in a PPARα-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-PPARα–infected hSMCs (Figure 2A, upper panel). Moreover, Wy14,643 treatment induced p16 expression in PPARα^+/+, but not in PPARα^–/– mSMCs (Figure 2A, lower panel). Interestingly, p16 mRNA levels were decreased by approximately two-thirds in PPARα^–/– compared with PPARα^+/+ mSMCs.

Figure 2

PPARα activation increases p16 gene transcription. (A) Regulation of p16 mRNA levels in hSMCs and mSMCs. Quantitative RT-PCR analyses were performed on RNA from Ad-GFP or Ad-PPARα–infected hSMCs treated or not treated with GW7647 (600 nM, upper panel) and on PPARα^–/– and PPARα^+/+ Sv/129 mSMCs treated or not treated with Wy14,643 (6 μM, lower panel). Values are expressed relative to the controls (mRNA levels in PPARα^+/+ mSMCs or hSMCs at T0) set as 1. (B) Induction of human p16 promoter activity by ligand-activated PPARα. pGL2 luciferase reporter constructs driven by the indicated p16 promoter deletion fragments (0.1 μg) were cotransfected in HeLa cells with or without pSG5 PPARα expression plasmid (0.3 μg). Cells were subsequently treated with fenofibric acid (100 μM) for 10 hours. Results (mean ± SD) from 1 representative experiment (n = 3) out of 3 is shown. (C) In vitro binding of PPARα to the p16DR1. EMSAs were carried out with end-labeled consensus DR1 (DR1cons), wild-type p16DR1 (p16DR1wt), or mutated p16DR1 (p16DR1mt) probes in the presence of unprogrammed reticulocyte lysate, RXR, PPARα, or both RXR and PPARα as indicated. (D) In-cell occupation of the p16 gene promoter by PPARα. Soluble chromatin was prepared from hSMCs treated with or without fenofibric acid (250 μM). IP was performed with anti-Sp1 or anti-PPARα antibodies, and DNA was amplified using specific primer pairs. Bottom lane shows schematic diagram depicting the fragments of the p16 and β-actin genes that were amplified. The nucleotide positions are indicated relative to the ATG. In panels B and D, A, B, and C denote the 3 Sp1-binding sites identified by Myohanen et al. (26) (see Results), and the putative transcription start site of the p16 promoter is indicated by an arrow.

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 PPARα, reporter constructs driven by different p16 promoter fragments were cotransfected with or without the pSG5-PPARα expression plasmid, and cells were subsequently treated with the PPARα agonists fenofibric acid or GW7647. Cotransfection of PPARα followed by ligand activation significantly induced activity of the –3.0-kb p16 gene promoter, indicating that p16 is a direct PPARα target gene (Figure 2B). PPARα-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 PPARα. Finally, deletion to –343 bp completely abolished activation of the p16 promoter by fenofibric acid–activated PPARα (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 PPARα-dependent p16 promoter activation, with the DR1 element mediating the majority of the response.

To determine whether PPARα physically binds to the p16DR1 site, electrophoretic mobility shift assays (EMSAs) were performed using in vitro–produced PPARα and RXR proteins and double-stranded [³²P]-labeled oligonucleotides (Table 2 and Figure 2C). Whereas neither RXR nor PPARα alone bound to the p16DR1 probe, a clear shift was observed upon incubation with both RXR and PPARα. This complex was supershifted by the anti-PPARα antibody and prevented by an excess of unlabeled wild-type, but not mutated, p16DR1 probe. Moreover, the mutated p16DR1 probe did not bind RXR/PPARα (Figure 2C). These results demonstrate that the RXR/PPARα heterodimer specifically binds in vitro to the p16DR1 site located at –1023 pb. In contrast, no PPARα/DNA complex formation was detected in EMSAs performed with oligonucleotides overlapping the proximal Sp1A- and B-binding sites (data not shown).

Table 2

Oligonucleotides used in this study

To investigate the in vivo occupancy of these p16DR1 and Sp1-binding sites by activated PPARα, chromatin IP (ChIP) assays were performed on DNA from hSMCs treated or not treated with fenofibric acid using anti-Sp1 and anti-PPARα antibodies (Figure 2D). In accordance with the EMSA results, the region spanning the p16DR1 sequence was immunoprecipitated by the anti-PPARα antibody, an effect which was enhanced by fenofibric acid treatment. In addition, in agreement with results of Myöhänen 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 PPARα in a PPARα ligand–dependent manner (Figure 2D). In contrast, no PPARα 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 β-actin genomic fragment did not yield a significant signal, thus demonstrating the specificity of PPARα immunoprecipitation and PCR amplification. Taken together, these results indicate that PPARα 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 α activation, resulting in the inhibition of CDK4-mediated pRB phosphorylation, is required for the SMC growth-inhibitory activity of PPARα. Next, it was determined whether the upregulation of p16 gene expression by PPARα results in an increase of p16 protein abundance and in-cell activity. Western blot analysis of hSMC whole-cell protein extracts revealed that PPARα overexpression or GW7647 treatment enhanced p16 protein levels in hSMCs, and the greatest increase was observed when cells were both infected with Ad-PPARα and treated with GW7647 (Figure 3A).

Figure 3

PPARα activation increases p16 protein levels and p16-CDK4 interaction and decreases CDK4-mediated pRB phosphorylation in hSMCs and mSMCs. (A–C) Protein extracts of hSMCs infected and treated for 12 hours (A) or 24 hours (B and C) with GW7647 (600 nM) were submitted to Western blot analysis either directly (A and B) or after IP with an anti-CDK4 antibody (C). Cellular p16 protein levels (A) and in-cell CDK4-mediated pRB phosphorylation (B) were measured using an antibody specific for p16 or an anti-pRB antibody recognizing a site specifically phosphorylated by cyclin D1/CDK4 (P∼Ser780-ppRB) (28), respectively. (C) Anti-CDK4 immunoprecipitates were assayed for kinase activity using the 769–921 pRB fragment as substrate (63) (top panel), or analyzed by Western blot using specific anti-p16 (middle panel) or anti-CDK4 antibodies (lower panel). (D) Protein extracts of PPARα^–/– or PPARα^+/+ C57BL/6J mSMCs treated for 24 hours with Wy14,643 (6 μM) were submitted to Western blot analysis using the anti-p16 and anti–P∼Ser780-ppRB antibodies.

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 PPARα controls p16 function, it was further determined whether the induction of p16 following PPARα activation affected p16/CDK4 interaction and CDK4 activity. To this end, in-cell pRB phosphorylation by CDK4, determined using an anti–P∼Ser780-ppRB antibody (28), and in vitro CDK4-associated activity using pRB as substrate were measured after PPARα 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 PPARα activation (Figure 1C), both analyses indicated that treatment with the PPARα ligand decreases pRB phosphorylation by CDK4 and that this effect is enhanced by PPARα overexpression. Moreover, the levels of p16 associated with CDK4 were increased in hSMCs infected with Ad-PPARα or treated with GW7647, and this interaction was further enhanced in Ad-PPARα–infected GW7647-treated hSMCs (Figure 3C, middle panel). Besides, in-cell p16 protein levels and CDK4-dependent pRB phosphorylation respectively increased and decreased in PPARα^+/+ but not in PPARα^–/– mSMCs following Wy14,643 treatment (Figure 3D).

Finally, the role of p16 in mediating the growth-inhibitory effects of PPARα 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 PPARα occurs via inhibition of the activity of G₁ CDKs, such as CDK4, due to an increased recruitment of p16.

Figure 4

p16 deficiency in mSMCs prevents the growth-inhibitory effects of fenofibrate. G₁-arrested p16^–/– or p16^+/+ C57BL/6J mSMCs resumed growth at T0 in normal GM containing or not containing Wy14,643 (10 μM). (A) Cells were harvested at the indicated times for measurement of DNA content (expressed as arbitrary units). Values are the mean ± SEM of triplicate points from a single experiment (n = 3/time point), which was repeated with 3 different cell preparations of primary mSMCs with similar results. (B) Cells were arrested after 24 hours for protein extraction followed by Western blot analysis using the anti–P∼Ser780-ppRB antibody.

PPAR α activation inhibits in vivo SMC proliferation upon carotid arterial injury by increasing p16 gene expression. The in vivo role of PPARα 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 PPARα^–/– mice,whereas it was drastically decreased in PPARα^+/+ mice treated with fenofibrate, whose action in mice requires PPARα expression (33, 34) (Figure 5, A and B). As previously reported (2, 21, 30), SMCs, identified by α-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 PPARα^+/+). 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 PPARα^–/– 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), PPARα 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).

Figure 5

PPARα activation inhibits neointima formation after mechanical carotid injury. Left carotid arteries were performed on PPARα^–/– (n = 6) or PPARα^+/+ Sv/129 mice treated (n = 5) or not (n = 8) with fenofibrate. (A) Topographic pattern of I/M area ratio 3 weeks after injury. Values are mean ± SEM of the I/M ratios measured on Masson’s trichrome–stained sections at the indicated distances from the junction between the external and internal branches of the left carotid artery (reference section at 0 μM). *P ≤ 0.05 vs. wild-type sections. (B) Representative carotid sections of uninjured right and injured left carotids at 50 μm from the junction stained with Masson’s trichrome. Scale bars: 100 μM. The internal elastic lamina delineating the neointima is indicated by a black arrow. (C) Representative sections of mouse carotid arteries immunostained for SMC (α-actin staining, brown, upper panel) or for proliferation marking (PCNA immunostaining to identify cells in S phase, brown, lower panel) at 150 μm and 100 μm from the junction, respectively. No intima was found in uninjured right carotids. Scale bars: 100 μM. The I/M ratio is equal to 0 in uninjured right carotid because intima of the healthy carotid artery is only composed of the monolayer endothelial cells. (D) Fenofibrate induces p16 mRNA levels in injured carotid arteries. Mouse carotid arteries were dissected at the indicated times after intraluminal mechanical injury of the left artery. p16 mRNA levels in the carotid segments are expressed as means ± SEM (n = 5) of fold change relative to the uninjured wild-type arteries, arbitrarily set to 1. *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001 vs. uninjured wild-type mice.

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 PPARα^+/+ but not in PPARα^–/– mice, an effect that was further enhanced following fenofibrate treatment. A maximal PPARα-dependent induction of p16 mRNA levels was reached within 24 hours and remained elevated 3 weeks after injury. Thus, these data indicate that PPARα 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 PPARα activation were further analyzed using p16^–/– mice (29). It is noteworthy that, similarly to what occurred in PPARα^–/– 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 PPARα activation on vascular SMC growth underlying intimal hyperplasia development.

Figure 6

p16 deficiency results in the induction of carotid artery hyperplasia that is not inhibited by fenofibrate. Left carotid arteries from p16^–/– C57BL/6J mice treated or not treated (n = 5/group) with fenofibrate were mechanically injured and analyzed after injury as described in Figure 5. (A) Topographic pattern of I/M area ratio. Values are mean ± SEM of the I/M ratios measured at the indicated distances from the junction on Masson’s trichrome–stained sections. *P ≤ 0.05, **P ≤ 0.01 vs. p16^+/+ mice. No significant difference was found between fenofibrate-treated and untreated p16^–/– mice. (B) Representative sections of mouse carotid arteries stained for morphometry or SMC. Carotid artery sections from untreated mice were stained with Masson’s trichrome or with an α-actin antibody at 50 and 25 μM from the junction, respectively. Scale bars: 300 μM. The internal elastic lamina is indicated by black arrows.

The cardiovascular protective effects of fibrates were initially mainly attributed to a systemic action of PPARα 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 PPARα controls cholesterol efflux and the inflammatory response in vessel wall cells (13), highlighting a direct regulatory role of PPARα on vessel wall function. However, despite the crucial role of SMC proliferation in the development of occlusive vascular diseases, the function of PPARα in the control of vascular SMC cell-cycle progression was not yet established. In the present study, we show that PPARα 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 PPARα 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 PPARα 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 PPARα on SMCs, PPARα activation was found to upregulate p16 promoter activity, thus identifying for the first time a molecular mechanism via which PPARα 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 p27^Kip1 (p27), whose protein degradation is prevented by PPARγ 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, PPARα deficiency and p16 deficiency both acted as molecular switches inducing SMC proliferation in vitro and in vivo (Figures 1 and 4–4 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 PPARα^+/+ compared with PPARα^–/– 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 PPARα 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 PPARα and p16 proteins are activated and function locally at the vascular artery injury site further extend these observations of a functional role for PPARα 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 PPARγ 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 PPARα/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 PPARα could provide a novel therapeutic strategy to prevent restenosis in treatments using either systemic administration or coated stents.

View Supplemental data

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. Monté and G. Peters for the gifts of plasmids, and F. Gonzalez for the gift of PPARα^–/– mice. We thank also the members of the B. Staels Laboratory for valuable discussions.

Florence Gizard and Carole Amant contributed equally to this work.

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, p16^INK4a; 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.

1. Dzau, VJ, Braun-Dullaeus, RC, Sedding, DG. Vascular proliferation and atherosclerosis: new perspectives and therapeutic strategies. Nat. Med. 2002. 8:1249-1256.
   View this article via: PubMed Google Scholar
2. Hao, H, Gabbiani, G, Bochaton-Piallat, ML. Arterial smooth muscle cell heterogeneity: implications for atherosclerosis and restenosis development. Arterioscler. Thromb. Vasc. Biol. 2003. 23:1510-1520.
   View this article via: PubMed Google Scholar
3. Ross, R. Atherosclerosis–an inflammatory disease. N. Engl. J. Med. 1999. 340:115-126.
   View this article via: PubMed Google Scholar
4. Yoshida, T, Owens, GK. Molecular determinants of vascular smooth muscle cell diversity. Circ. Res. 2005. 96:280-291.
   View this article via: PubMed Google Scholar
5. Walworth, NC. Cell-cycle checkpoint kinases: checking in on the cell cycle. Curr. Opin. Cell Biol. 2000. 12:697-704.
   View this article via: PubMed Google Scholar
6. Harbour, JW, Dean, DC. Rb function in cell-cycle regulation and apoptosis. Nat. Cell Biol. 2000. 2:E65-E67.
   View this article via: PubMed Google Scholar
7. Ekholm, SV, Reed, SI. Regulation of G(1) cyclin-dependent kinases in the mammalian cell cycle. Curr. Opin. Cell Biol. 2000. 12:676-684.
   View this article via: PubMed Google Scholar
8. Serrano, M, Hannon, GJ, Beach, D. A new regulatory motif in cell-cycle control causing specific inhibition of cyclin D/CDK4. Nature. 1993. 366:704-707.
   View this article via: PubMed Google Scholar
9. Tam, SW, Shay, JW, Pagano, M. Differential expression and cell cycle regulation of the cyclin-dependent kinase 4 inhibitor p16Ink4. Cancer Res. 1994. 54:5816-5820.
   View this article via: PubMed Google Scholar
10. Parry, D, Bates, S, Mann, DJ, Peters, G. Lack of cyclin D-Cdk complexes in Rb-negative cells correlates with high levels of p16INK4/MTS1 tumour suppressor gene product. EMBO J. 1995. 14:503-511.
    View this article via: PubMed Google Scholar
11. Hara, E, et al. Regulation of p16CDKN2 expression and its implications for cell immortalization and senescence. Mol. Cell. Biol. 1996. 16:859-867.
    View this article via: PubMed Google Scholar
12. Michalik, L, Desvergne, B, Wahli, W. Peroxisome-proliferator-activated receptors and cancers: complex stories. Nat. Rev. Cancer. 2004. 4:61-70.
    View this article via: PubMed Google Scholar
13. Marx, N, Duez, H, Fruchart, JC, Staels, B. Peroxisome proliferator-activated receptors and atherogenesis: regulators of gene expression in vascular cells. Circ. Res. 2004. 94:1168-1178.
    View this article via: PubMed Google Scholar
14. Brown, WV. Potential use of fenofibrate and other fibric acid derivatives in the clinic. Am. J. Med. 1987. 83:85-89.
    View this article via: PubMed Google Scholar
15. Willson, TM, Brown, PJ, Sternbach, DD, Henke, BR. The PPARs: from orphan receptors to drug discovery. J. Med. Chem. 2000. 43:527-550.
    View this article via: PubMed Google Scholar
16. Robins, SJ, et al. Relation of gemfibrozil treatment and lipid levels with major coronary events: VA-HIT: a randomized controlled trial. JAMA. 2001. 285:1585-1591.
    View this article via: PubMed Google Scholar
17. Steiner, G. The Diabetes Atherosclerosis Intervention Study (DAIS): a study conducted in cooperation with the World Health Organization. The DAIS Project Group. Diabetologia. 1996. 39:1655-1661.
    View this article via: PubMed Google Scholar
18. Frick, MH, et al. Prevention of the angiographic progression of coronary and vein-graft atherosclerosis by gemfibrozil after coronary bypass surgery in men with low levels of HDL cholesterol. Lopid Coronary Angiography Trial (LOCAT) Study Group. Circulation. 1997. 96:2137-2143.
    View this article via: PubMed Google Scholar
19. Ericsson, CG, et al. Angiographic assessment of effects of bezafibrate on progression of coronary artery disease in young male postinfarction patients. Lancet. 1996. 347:849-853.
    View this article via: PubMed Google Scholar
20. Staels, B, et al. Mechanism of action of fibrates on lipid and lipoprotein metabolism. Circulation. 1998. 98:2088-2093.
    View this article via: PubMed Google Scholar
21. Zahradka, P, et al. Activation of peroxisome proliferator-activated receptors alpha and gamma1 inhibits human smooth muscle cell proliferation. Mol. Cell. Biochem. 2003. 246:105-110.
    View this article via: PubMed Google Scholar
22. Staels, B, et al. Activation of human aortic smooth-muscle cells is inhibited by PPARalpha but not by PPARgamma activators. Nature. 1998. 393:790-793.
    View this article via: PubMed Google Scholar
23. Marx, N, Schonbeck, U, Lazar, MA, Libby, P, Plutzky, J. Peroxisome proliferator-activated receptor gamma activators inhibit gene expression and migration in human vascular smooth muscle cells. Circ. Res. 1998. 83:1097-1103.
    View this article via: PubMed Google Scholar
24. Nigro, J, Dilley, RJ, Little, PJ. Differential effects of gemfibrozil on migration, proliferation and proteoglycan production in human vascular smooth muscle cells. Atherosclerosis. 2002. 162:119-129.
    View this article via: PubMed Google Scholar
25. Brown, PJ, et al. Identification of a subtype selective human PPARalpha agonist through parallel-array synthesis. Bioorg. Med. Chem. Lett. 2001. 11:1225-1227.
    View this article via: PubMed Google Scholar
26. Myohanen, S, Baylin, SB. Sequence-specific DNA binding activity of RNA helicase A to the p16INK4a promoter. J. Biol. Chem. 2001. 276:1634-1642.
    View this article via: PubMed Google Scholar
27. Sherr, CJ, Roberts, JM. CDK inhibitors: positive and negative regulators of G1-phase progression. Genes Dev. 1999. 13:1501-1512.
    View this article via: PubMed Google Scholar
28. Kitagawa, M, et al. The consensus motif for phosphorylation by cyclin D1-Cdk4 is different from that for phosphorylation by cyclin A/E-Cdk2. EMBO J. 1996. 15:7060-7069.
    View this article via: PubMed Google Scholar
29. Krimpenfort, P, Quon, KC, Mooi, WJ, Loonstra, A, Berns, A. Loss of p16Ink4a confers susceptibility to metastatic melanoma in mice. Nature. 2001. 413:83-86.
    View this article via: PubMed Google Scholar
30. Carmeliet, P, et al. Urokinase but not tissue plasminogen activator mediates arterial neointima formation in mice. Circ. Res. 1997. 81:829-839.
    View this article via: PubMed Google Scholar
31. Wang, X, Paigen, B. Comparative genetics of atherosclerosis and restenosis: exploration with mouse models. Arterioscler. Thromb. Vasc. Biol. 2002. 22:884-886.
    View this article via: PubMed Google Scholar
32. Kuhel, DG, Zhu, B, Witte, DP, Hui, DY. Distinction in genetic determinants for injury-induced neointimal hyperplasia and diet-induced atherosclerosis in inbred mice. Arterioscler. Thromb. Vasc. Biol. 2002. 22:955-960.
    View this article via: PubMed Google Scholar
33. Gonzalez, FJ, Peters, JM, Cattley, RC. Mechanism of action of the nongenotoxic peroxisome proliferators: role of the peroxisome proliferator-activator receptor alpha. J. Natl. Cancer Inst. 1998. 90:1702-1709.
    View this article via: PubMed Google Scholar
34. Kockx, M, et al. Fibrates suppress fibrinogen gene expression in rodents via activation of the peroxisome proliferator-activated receptor-alpha. Blood. 1999. 93:2991-2998.
    View this article via: PubMed Google Scholar
35. Saitoh, K, et al. Anti-atheromatous effects of fenofibrate, a hypolipidemic drug. I: Anti-atheromatous effects are independent of its hypolipidemic effect in cholesterol-fed rabbits [In Japanese]. Nippon Yakurigaku Zasshi. 1995. 106:41-50.
    View this article via: PubMed Google Scholar
36. Saitoh, K, Mori, T, Kasai, H, Nagayama, T, Ohbayashi, S. Anti-atheromatous effects of fenofibrate, a hypolipidemic drug. II: Anti-atheromatous effects are independent of its hypolipidemic effect in hereditary hyperlipidemic rabbits [In Japanese]. Nippon Yakurigaku Zasshi. 1995. 106:51-60.
    View this article via: PubMed Google Scholar
37. Rubins, HB, et al. Gemfibrozil for the secondary prevention of coronary heart disease in men with low levels of high-density lipoprotein cholesterol. Veterans Affairs High-Density Lipoprotein Cholesterol Intervention Trial Study Group. N. Engl. J. Med. 1999. 341:410-418.
    View this article via: PubMed Google Scholar
38. Ross, JS, Stagliano, NE, Donovan, MJ, Breitbart, RE, Ginsburg, GS. Atherosclerosis and cancer: common molecular pathways of disease development and progression. Ann. N. Y. Acad. Sci. 2001. 947:271-292; discussion 292–293.
    View this article via: PubMed Google Scholar
39. Wakino, S, et al. Peroxisome proliferator-activated receptor gamma ligands inhibit retinoblastoma phosphorylation and G1--> S transition in vascular smooth muscle cells. J. Biol. Chem. 2000. 275:22435-22441.
    View this article via: PubMed Google Scholar
40. Tanner, FC, et al. Differential effects of the cyclin-dependent kinase inhibitors p27(Kip1), p21(Cip1), and p16(Ink4) on vascular smooth muscle cell proliferation. Circulation. 2000. 101:2022-2025.
    View this article via: PubMed Google Scholar
41. Boehm, M, et al. Bone marrow-derived immune cells regulate vascular disease through a p27(Kip1)-dependent mechanism. J. Clin. Invest. 2004. 114:419-426. doi:10.1172/JCI200420176.
    View this article via: JCI PubMed Google Scholar
42. Belinsky, SA, et al. Aberrant methylation of p16(INK4a) is an early event in lung cancer and a potential biomarker for early diagnosis. Proc. Natl. Acad. Sci. U. S. A. 1998. 95:11891-11896.
    View this article via: PubMed Google Scholar
43. Nuovo, GJ, Plaia, TW, Belinsky, SA, Baylin, SB, Herman, JG. In situ detection of the hypermethylation-induced inactivation of the p16 gene as an early event in oncogenesis. Proc. Natl. Acad. Sci. U. S. A. 1999. 96:12754-12759.
    View this article via: PubMed Google Scholar
44. Wong, DJ, Foster, SA, Galloway, DA, Reid, BJ. Progressive region-specific de novo methylation of the p16 CpG island in primary human mammary epithelial cell strains during escape from M(0) growth arrest. Mol. Cell. Biol. 1999. 19:5642-5651.
    View this article via: PubMed Google Scholar
45. Sharpless, NE, DePinho, RA. The INK4A/ARF locus and its two gene products. Curr. Opin. Genet. Dev. 1999. 9:22-30.
    View this article via: PubMed Google Scholar
46. Kersten, S, et al. Peroxisome proliferator-activated receptor alpha mediates the adaptive response to fasting. J. Clin. Invest. 1999. 103:1489-1498.
    View this article via: JCI PubMed Google Scholar
47. Guerre-Millo, M, et al. PPAR-alpha-null mice are protected from high-fat diet-induced insulin resistance. Diabetes. 2001. 50:2809-2814.
    View this article via: PubMed Google Scholar
48. Devchand, PR, et al. The PPARalpha-leukotriene B4 pathway to inflammation control. Nature. 1996. 384:39-43.
    View this article via: PubMed Google Scholar
49. Gervois, P, et al. Global suppression of IL-6-induced acute phase response gene expression after chronic in vivo treatment with the peroxisome proliferator-activated receptor-alpha activator fenofibrate. J. Biol. Chem. 2004. 279:16154-16160.
    View this article via: PubMed Google Scholar
50. Froeschl, M, Olsen, S, Ma, X, O’Brien, ER. Current understanding of in-stent restenosis and the potential benefit of drug eluting stents. Curr. Drug Targets Cardiovasc. Haematol. Disord. 2004. 4:103-117.
    View this article via: PubMed Google Scholar
51. Law, RE, et al. Troglitazone inhibits vascular smooth muscle cell growth and intimal hyperplasia. J. Clin. Invest. 1996. 98:1897-1905.
    View this article via: JCI PubMed Google Scholar
52. Takagi, T, et al. Pioglitazone reduces neointimal tissue proliferation after coronary stent implantation in patients with type 2 diabetes mellitus: an intravascular ultrasound scanning study. Am. Heart J. 2003. 146:E5.
    View this article via: PubMed Google Scholar
53. Choi, D, et al. Preventative effects of rosiglitazone on restenosis after coronary stent implantation in patients with type 2 diabetes. Diabetes Care. 2004. 27:2654-2660.
    View this article via: PubMed Google Scholar
54. Barbier, O, et al. FXR induces the UGT2B4 enzyme in hepatocytes: a potential mechanism of negative feedback control of FXR activity. Gastroenterology. 2003. 124:1926-1940.
    View this article via: PubMed Google Scholar
55. Chartier, C, et al. Efficient generation of recombinant adenovirus vectors by homologous recombination in Escherichia coli. J. Virol. 1996. 70:4805-4810.
    View this article via: PubMed Google Scholar
56. Sardet, C, et al. E2F-4 and E2F-5, two members of the E2F family, are expressed in the early phases of the cell cycle. Proc. Natl. Acad. Sci. U. S. A. 1995. 92:2403-2407.
    View this article via: PubMed Google Scholar
57. Corsini, A, et al. Effect of the new calcium antagonist lercanidipine and its enantiomers on the migration and proliferation of arterial myocytes. J. Cardiovasc. Pharmacol. 1996. 28:687-694.
    View this article via: PubMed Google Scholar
58. Fiszer-Szafarz, B, Szafarz, D, Guevara de Murillo, A. A general, fast, and sensitive micromethod for DNA determination application to rat and mouse liver, rat hepatoma, human leukocytes, chicken fibroblasts, and yeast cells. Anal. Biochem. 1981. 110:165-170.
    View this article via: PubMed Google Scholar
59. Kherrouche, Z, De Launoit, Y, Monte, D. Human E2F6 is alternatively spliced to generate multiple protein isoforms. Biochem. Biophys. Res. Commun. 2004. 317:749-760.
    View this article via: PubMed Google Scholar
60. Draetta, G, Brizuela, L, Potashkin, J, Beach, D. Identification of p34 and p13, human homologs of the cell cycle regulators of fission yeast encoded by cdc2+ and suc1+. Cell. 1987. 50:319-325.
    View this article via: PubMed Google Scholar
61. Vu-Dac, N, et al. Transcriptional regulation of apolipoprotein A-I gene expression by the nuclear receptor RORalpha. J. Biol. Chem. 1997. 272:22401-22404.
    View this article via: PubMed Google Scholar
62. Lee, SS, et al. Targeted disruption of the alpha isoform of the peroxisome proliferator-activated receptor gene in mice results in abolishment of the pleiotropic effects of peroxisome proliferators. Mol. Cell. Biol. 1995. 15:3012-3022.
    View this article via: PubMed Google Scholar
63. Zarkowska, T, Mittnacht, S. Differential phosphorylation of the retinoblastoma protein by G1/S cyclin-dependent kinases. J. Biol. Chem. 1997. 272:12738-12746.
    View this article via: PubMed Google Scholar
64. Gizard, F, et al. The transcriptional regulating protein of 132 kDa (TReP-132) differentially influences steroidogenic pathways in human adrenal NCI-H295 cells. J. Mol. Endocrinol. 2004. 32:557-569.
    View this article via: PubMed Google Scholar