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J. Biol. Chem., Vol. 277, Issue 13, 11505-11512, March 29, 2002

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Peroxisome Proliferator-activated Receptor  Is Up-regulated during Vascular Lesion Formation and Promotes Post-confluent Cell Proliferation in Vascular Smooth Muscle Cells^*

Jifeng Zhang, Mingui Fu, Xiaojun Zhu, Yan Xiao, Yongshan Mou, Hui Zheng, Mukaila A. Akinbami, Qian Wang, and Yuqing E. Chen^§

From the Cardiovascular Research Institute, Morehouse School of Medicine, Atlanta, Georgia 30310

Received for publication, November 2, 2001, and in revised form, December 28, 2001

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Although peroxisome proliferator-activated receptor (PPAR)  is widely expressed in many tissues, the role of PPAR is poorly understood. In this study, we report that PPAR was up-regulated in vascular smooth muscle cells (VSMC) during vascular lesion formation. By using Northern blot analysis, we demonstrated that PPAR was increased by 3-4-fold in VSMC treated with platelet-derived growth factor (PDGF) (20 ng/ml). In addition, PDGF-induced PPAR mRNA expression neither needs de novo protein synthesis nor affects the stability of PPAR mRNA in VSMC. Preincubation of VSMC with phosphatidylinositol 3-kinase inhibitor (LY294002, 50 µmol/liter) or infection of VSMC with an adenovirus carrying the gene for a dominant negative form of Akt abrogated PDGF-induced PPAR mRNA expression, suggesting that phosphatidylinositol 3-kinase/Akt signaling pathway is involved in the regulation of PDGF-induced PPAR mRNA expression in VSMC. To explore the role of PPAR in VSMC, we generated rat vascular smooth muscle cells (A7r5) stably overexpressing PPAR and the control green fluorescent protein. Overexpression of PPAR in VSMC increased post-confluent cell proliferation by increasing the cyclin A and CDK2 as well as decreasing p57^kip2. Taken together, the results suggest that PPAR plays an important role in the pathology of diseases associated with VSMC proliferation, such as primary atherosclerosis and restenosis.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

The peroxisome proliferator-activated receptors (PPARs)¹ including , /, and  are members of the superfamily of nuclear receptors. The general structural features of the family include a central DNA-binding domain and a carboxyl-terminal domain that mediates ligand binding, dimerization, and transactivation functions. PPARs function as a heterodimer with retinoid X receptors, another member of this family, to bind to the PPAR-responsive element, a DR1 element, which is a direct repeat of two similar hexanucleotide (5'-AGGTCA-3') half-sites separated by one nucleotide on its target genes (1). In the presence of both PPAR- and retinoid X receptor-specific ligands, this type of interaction confers synergistic activation of target genes (1).

PPAR is highly expressed in the liver, muscle, kidney, and heart, where it stimulates the -oxidative degradation of fatty acids. PPAR is most abundantly expressed in fat cells, large intestine, and cells of the monocyte lineage (2). PPAR has been linked to adipocyte differentiation and insulin sensitivity. Both PPAR and PPAR are expressed in monocytes/macrophages, endothelial cells, and vascular smooth muscle cells (VSMC) in both medial and intimal layers. Activation of PPAR has been found to inhibit VSMC production of inflammatory factors (3), although activation of PPAR has been reported to decrease both VSMC proliferation (4) and matrix production after vascular injury (5). Therefore, PPAR and PPAR are emerging as important determinants of vascular function and structure (6-8).

Although PPAR (also known as PPAR and NUC-1) is widely expressed in many tissues, the physiological or pathophysiological roles of PPAR are unclear. With no connection to important clinical manifestations, along with the lack of marketed PPAR-specific ligands, the research to define PPAR function has been hampered for many years. However, PPAR has recently been linked to colon cancer proliferation (9-11), preadipocyte proliferation (12, 13), macrophage lipid accumulation (14), and embryo implantation (15). Interestingly, prostacyclin (PGI₂), which is the natural ligand of PPAR, is the characteristic prostanoid released by vascular endothelial and smooth muscle cells in response to stimulation by cytokines such as tumor necrosis factor  (16). Taken together, we hypothesize that PPAR plays an important role in VSMC proliferation.

VSMC proliferation is the key component of vasculoproliferative diseases including atherosclerosis, restenosis, and vein-graft failure (17, 18). Cytokines and growth factors such as tumor necrosis factor  and PDGF participate in these processes (19). In eukaryotic cells, the commitment to divide is made in the G₁ phase of the cell cycle in response to various stimuli, including growth factors. The D- and E-type cyclins in combination with cyclin-dependent kinases (CDKs) regulate passage through the G₁ phase (20, 21). Overexpression of cyclin D or E can shorten G₁ phase (22-24), suggesting that the cyclin family is critical for progression through G₁ phase.

In this report, we document that PPAR is up-regulated in VSMC during vascular lesion formation, and we demonstrate that PDGF stimulation increases PPAR expression by 3-4-fold in VSMC. We also show that overexpression of PPAR in VSMC promotes the proliferation of confluent cells by increasing the cyclin A and CDK2 but decreased p57^kip2.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Materials-- Platelet-derived growth factor (PDGF) BB, actinomycin D, and cycloheximide were purchased from Sigma. LY294002, SB202190, and U0126 were obtained from Biomol Research Laboratories (Plymouth Meeting, PA). Cell culture medium and phosphate-buffered saline were purchased from Invitrogen.

Cell Culture and Stimulation-- The rat aortic smooth muscle cells (RASMC) were prepared as described previously (25). The RASMCs were confirmed by  smooth muscle actin immunochemical staining using anti--actin kit (Dako). Passage 6-10 RASMCs were used in DMEM/F-12 containing 10% FBS, 100 units/ml penicillin, 100 µg/ml streptomycin, and 200 mM L-glutamine. Human aortic smooth muscle cells were purchased from BioWhittaker (San Diego, CA). The cells were cultured in smooth muscle cell growth medium-2 containing 5% FBS, 2 ng/ml human basic fibroblast growth factor, 0.5 ng/ml human epidermal growth factor, 50 µg/ml gentamicin, 50 ng/ml amphotericin-B, and 5 µg/ml bovine insulin. For all experiments, early passages 5-7 of human aortic smooth muscle cells were grown to 80-90% confluence and made quiescent by serum starvation (0.4% FBS) for at least 24 h. The LY294002, SB202190, and U0126 inhibitors were added 30 min before the addition of human recombinant PDGF-BB (Sigma). The actinomycin D was added to the cells after PDGF-BB stimulation for 6 h. The cycloheximide was added to the cells at the same time with PDGF-BB. The rat vascular smooth muscle cell line A7r5 was purchased from ATCC (catalog number CRL-1444, Manassas, VA).

Balloon Injury and Immunohistochemical Staining-- Male Sprague-Dawley rat weighing 280-300 g were purchased from Taconic Farms (Germantown, NY). Balloon-catheter injury was induced when rats were under ketamine (90 mg/kg) and xylazine (5 mg/kg) anesthesia. The left common carotid artery wall was injured with an embolectomy balloon catheter (2F Fogarty, Edwards Life Sciences, Memphis, TN) to induce neointimal formation as described previously, and the right common carotid artery was served as control (26). Animals were killed with an overdose of pentobarbital (120 mg/kg) and subjected to whole body perfusion with 4% paraformaldehyde at 7, 14, and 28 days after balloon injury. The carotid arteries were removed, cut into cross-sectional segments, and embedded in paraffin. Sections 5 µm thick (n = 5 per animal) were immunohistologically stained with a polyclonal antibody (Santa Cruz Biotechnology, 1:500 dilution) against PPAR. The sections were counterstained with hematoxylin. The image was displayed in a high resolution monitor and digitized by a video frame grabber (PCVISION Plus, Imaging Technology) on an IBM-compatible computer.

RNA Isolation and Northern Blot Analysis-- Twenty µg of total RNA, isolated from each condition by using acid-guanidinium thiocyanate, was subjected to electrophoresis through 1% formaldehyde-agarose gels. After transferring to nylon membranes (Bio-Rad), the RNA was cross-linked to the membrane by an UV cross-linker (Bio-Rad). ³²P-Labeled cDNA probes were generated by using the random primer labeling system (Invitrogen). Blots were pre-hybridized, hybridized, and were washed once with 1× SSC at 65 °C for 30 min and once with 0.1× SSC, 1.0% SDS (w/v) at 65 °C for 15 min. The lane loading differences were normalized using the GAPDH.

Western Blot Analysis-- Fifty µg of total cell lysate isolated from each condition was subjected to SDS-PAGE and electrotransfered to nitrocellulose membrane (Bio-Rad). After blocking in 20 mM Tris-HCl, pH 7.6, containing 150 mM NaCl, 0.1% Tween 20, and 5% (w/v) non-fat dry milk, blots were incubated for 1 h at 4 °C with specific antibodies (Santa Cruz Biotechnology) against PPAR (sc-7197), cyclin A (sc-596), cyclin E (sc-481), Cdk2 (sc-163), p57^kip2 (sc-1040), or actin (sc-1616). The blots were then incubated with horseradish peroxidase-conjugated secondary antibody (Santa Cruz Biotechnology). Immunoactivity was visualized by the enhanced chemiluminescence detection system (ECL, Amersham Biosciences) according to the manufacturer's instructions.

Quantitative RT-PCR-- The expression levels of PPAR mRNA in rat carotid arteries were quantitated by the quantitative real-time RT-PCR strategy (Roche LightCycler PCR system, Roche Molecular Biochemicals). Sham-operated and balloon-injured rat carotid arteries at 7, 14, and 28 days after surgery were harvested. The adventitia of artery was removed from the medial layer by gross forceps dissection. Pooled samples (n = 4) for each group were used for RNA preparation. Two primers, ratPup2 (5'-cagccataacgcacccttcatcatcc-3', nt 867-892) and ratPlow2 (5'-ggccaccagcagtccgtctttgttg-3', nt 1170 to 1146) corresponding to rat PPAR cDNA (GenBank^TM accession number NM_013141) were used for quantitative PCR. The PCR results were normalized by GAPDH.

Adenovirus Preparation and Infection-- An adenovirus carrying the gene for a dominant negative form of Akt (ad-AktDN) was obtained from Dr. Ogawa (27). The ad-AktDN was amplified as described previously (28). In this study, VSMCs were infected with adenovirus vectors at ~5 plaque-forming units/cell. The cells were subjected to experiments 24-48 h after infection.

Construction of PPAR-A7r5 Stable Cells-- The mouse PPAR cDNA was a gift from Dr. Grimaldi (29). Establishment of stable transfectants of A7r5 cells expressing PPAR/GFP and GFP alone was performed by using the retroviral bicistronic expression vector pMX-IRES-GFP as described previously (30). Briefly, the PPAR cDNA was inserted into the upstream of the encephalomyocarditis virus internal ribosomal entry sequence (IRES) which drives a GFP gene in the retroviral vector pMX. We infected 5 × 10⁵ A7r5 cells with ~2 × 10⁶ virus supernatant in the presence of 4 µg/ml of Polybrene for 4 h. Forty eight hours after the infection, cells were sorted by a FACS system (BD PharMingen) according to their GFP levels. A homogeneous population of PPAR-A7r5 stable cells isolated by FACS was used for this study. At the same time, we generated the control A7r5 cells expressing GFP.

Cell Number Determination and Cell Cycle Distribution-- To investigate the growth rate and cell cycle distribution between PPAR-A7r5 and GFP-A7r5 cells, the cells were plated in 6-well plates at a density of 1.6 × 10⁵ cells per well in DMEM containing 10% FBS. The medium was replaced every other day. The number of cells was determined by the Coulter counter (model ZM, Coulter Electronics Ltd., Inc., Hialeah, FL) at different time intervals after seeding and was averaged for four wells.

Flow cytometry was performed to analyze cell cycle distribution between PPAR-A7r5 and GFP-A7r5 cells. Briefly, the post-confluent cells were trypsinized, centrifuged at 1500 rpm for 5 min, washed with phosphate-buffered saline, and treated with 20 µg/ml RNase A and 0.2% Triton X-100 for 30 min at 37 °C. The cell DNA was then stained with 100 µg/ml propidium iodide for 30 min at 4 °C and covered with aluminum foil. Samples were analyzed for DNA content by using a standard method on a FACScan (BD PharMingen FACS System). DNA histogram analysis was performed using the CellQuest software (BD PharMingen).

Statistical Analysis-- Each experiment was repeated a minimum of three times. Statistical analyses were performed by analysis of variance and unpaired 2-tailed Student's t test. Data are presented as means ± S.E. The value for p < 0.05 was considered significant.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

PPAR Is Expressed in Vascular Smooth Muscle Cells-- To determine whether PPAR is expressed in VSMC, we examined the expression levels of PPAR in both human and rat aortic smooth muscle cells. By using both Northern blot and Western blot analyses, we demonstrated that PPAR is expressed in both human and rat VSMC (data not shown). These data document that PPAR is expressed in vascular smooth muscle cells as described previously (31).

PPAR Is Up-regulated during Vascular Lesion Formation after Balloon Injury of Rat Carotid Artery-- To explore the role of PPAR in VSMC, we compared the PPAR expression levels between normal and injured vessels using the balloon-injured carotid artery model. Immunohistochemical results showed that neointima formation in this model was associated with a significant increase in PPAR expression (Fig. 1B) compared with control (Fig. 1A).

View larger version (36K): [in this window] [in a new window]   Fig. 1.   The PPAR expression is increased in rat carotid artery after balloon injury. A representative section shows strong immunoreactive staining for PPAR in neointima at 14 days after balloon injury (B). The neointima is defined as the area between the vessel lumen and the internal elastic lamina (IEL). Faint staining for PPAR is observed in the media of both sham-operated (A) and balloon-injured (B) carotid arteries. Endothelial cells express PPAR as observed in the control artery. The sections were counterstained with hematoxylin. Magnification 600; bar, 10 µm. C, the level of PPAR expression was detected by using quantitative RT-PCR. Values normalized by GAPDH are expressed as means ± S.E. (n = 3; p < 0.01). In each experiment, the level of PPAR mRNA in sham-operated carotid artery was assigned an arbitrary value of 1.

To define the expression level of PPAR in injured artery, we performed quantitative RT-PCR experiments using RNA samples isolated from sham-operated and balloon-injured rat carotid arteries. As shown in Fig. 1C, the level of PPAR mRNA expression in balloon-injured rat carotid arteries was ~3.1-fold higher than that in sham-operated rat carotid arteries at 14 days after balloon injury. In addition, the PPAR mRNA levels were increased 1.7- and 2.1-fold at 7 and 28 days after injury, respectively (data not shown). These observations indicate that PPAR may function as an important determinant of vascular lesion formation.

PDGF Induces PPAR Expression in a Time- and Dose-dependent Manner in VSMC-- It was very interesting to document that PPAR is up-regulated during vascular lesion formation. To investigate whether PDGF is responsible for the increase of PPAR expression in the neointima, RASMCs were treated with 20 ng/ml PDGF for 0, 0.5, 2, 6, 12, 24, 48, and 72 h. Northern blot analysis showed that the levels of PPAR mRNA increased at 2 h, reached a peak at 6 h, and remained above the control level at least for 24 h after PDGF stimulation (Fig. 2A).

View larger version (47K): [in this window] [in a new window]   Fig. 2.   PDGF induces PPAR expression in a time- and dose-dependent manner. Rat aortic smooth muscle cells were made quiescent by serum starvation (0.4% FCS) for 24 h and then treated with 20 ng/ml PDGF to study the effect of time as indicated (A) or with an increasing concentration of PDGF for 6 h to study effect of dose (B). The PPAR mRNA levels were analyzed by Northern blot analyses. The values were normalized by GAPDH, and the control was assigned an arbitrary value of 1. C, Western blot analysis of PPAR protein levels. The cells were treated with 20 ng/ml PDGF for different times in A. Fifty µg of the cell lysate was used for the analysis. Values were normalized by actin level. Three independent experiments showed similar results.

The dose response to PDGF-induced PPAR mRNA expression was documented at 6 h of PDGF stimulation. As shown in Fig. 2B, the expression of PPAR mRNA was up-regulated in a dose-dependent manner. A significant increase was observed at a PDGF concentration as low as 5 ng/ml, whereas maximal increases were obtained at a concentration of 10 ng/ml. These results reveal that PDGF can activate PPAR gene expression in VSMC.

The effect of PDGF on PPAR protein levels was also assessed by Western blot analysis. RASMCs were treated with 20 ng/ml PDGF for 0, 0.5, 2, 6, 12, 24, and 48 h. Two bands around 52-55 kDa were detected by anti-PPAR antibody. The upper band, which may be caused by PDGF-induced phosphorylation of PPAR, was increased as early as 0.5 h and reached a peak at 24 h. The lower band was first decreased at 0.5 and 2 h and then increased at 12 h and reached a peak at 24 h. These results demonstrate that PDGF induces PPAR protein in a time-dependent manner in RASMC. In parallel experiments, similar results were observed in human aortic vascular smooth muscle cells (data not shown) by both Northern blot and Western blot analyses.

PDGF-induced PPAR mRNA Expression Does Not Affect the Stability of PPAR mRNA in VSMC-- To evaluate whether PPAR mRNA stability contributes to PDGF-induced PPAR gene expression, we examined the half-life of PPAR mRNA in RASMC. Northern blot analyses were performed with the addition of actinomycin D (5 µg/ml) after 6 h of PDGF (20 ng/ml) or vehicle stimulation. In RASMC, the half-life of PPAR mRNA was ~3.4 h. There was no significant difference between PDGF-treated and -untreated cells (Fig. 3A).

View larger version (22K): [in this window] [in a new window]   Fig. 3.   A, decay of PPAR mRNA in the presence of actinomycin D. Rat aortic smooth muscle cells were incubated with or without 20 ng/ml PDGF for 6 h, and de novo PPAR transcripts were inhibited by addition of actinomycin D (5 µg/ml). Total RNA was isolated at 0, 2, and 4 h after administration of actinomycin D. A representative Northern blot (top) and the quantitative graph (bottom) of three experiments are shown. The relative values were normalized by GAPDH. B, effects of cycloheximide on PDGF-induced PPAR mRNA expression. Total RNA was isolated from the cells treated with or without PDGF (20 ng/ml) and cycloheximide (10 µg/ml) for 24 h. A representative Northern blot (top) and the quantitative graph (bottom) are shown. The relative values were normalized by GAPDH (n = 3, *, p < 0.01).

PDGF-induced PPAR mRNA Expression Is Not Involved in de Novo Protein Synthesis-- To examine whether de novo protein synthesis is required for PDGF-induced PPAR gene expression, we examined PPAR mRNA levels from RASMC treated with or without PDGF (20 ng/ml) and in the presence or absence of cycloheximide (10 µg/ml) for 24 h. As shown in Fig. 3B, the protein translation inhibitor, cycloheximide, did not alter PPAR mRNA levels after 24 h of PDGF stimulation, suggesting that PDGF-induced PPAR mRNA expression does not require de novo protein synthesis.

PDGF Induces PPAR Expression by PI3-kinase/Akt Signaling Pathway in VSMC-- To investigate the signaling pathways mediating PDGF-induced PPAR expression, we initially focused on defining the roles of the PI3-kinase, MEK/ERK, and p38 mitogen-activated protein kinase. RASMCs were treated with 20 ng/ml PDGF for 6 h after pretreatment with LY294002, a PI3-kinase inhibitor (50 µM); SB202190, a p38 kinase inhibitor (25 µM); or U0126, a MEK inhibitor (10 µM) for 30 min. As shown in Fig. 4, LY294002 completely blocked the effect of PDGF (p < 0.01). Although inhibition of p38 mitogen-activated protein kinase significantly attenuated the effect of PDGF-induced PPAR expression by 83 ± 9.5% (p < 0.01), SB202190 alone reduced the basic level of PPAR expression in VSMC by 70 ± 8.9% (p < 0.01). However, inhibition of MEK increased PDGF-induced PPAR mRNA expression by 44 ± 8% (p < 0.01).

View larger version (26K): [in this window] [in a new window]   Fig. 4.   The PI3-kinase inhibitor blocks PDGF-induced PPAR expression in VSMC. Cells were treated with LY294002 (LY) (50 µmol/liter, a PI3-kinase inhibitor), SB202190 (SB) (25 µmol/liter, a p38 kinase inhibitor), or U0126 (U) (10 µmol/liter, a MEK inhibitor) for 30 min and then 20 ng/ml PDGF was added and incubated for 6 h. PPAR mRNA levels were determined by Northern blot analyses. A representative Northern blot (top) and the quantitative graph of three experiments are shown (bottom). Values were normalized by GAPDH (n = 3, *, p < 0.01).

To define further whether the PI3-kinase/Akt signaling pathway mediates PDGF-induced PPAR gene expression in VSMC, we selectively blocked this signaling pathway by using ad-AktDN (an adenovirus carrying the gene for a dominant negative form of Akt). Blockade of the PI3-kinase/Akt pathway effectively prevented PDGF-induced PPAR gene expression in RASMC (Fig. 5). However, PPAR expression was not affected by the control adenovirus (Ad-GFP) infection in RASMC (data not shown). Taken together, these results provided the first evidence that PDGF-induced PPAR gene expression is regulated by a PI3-kinase/Akt-dependent pathway.

View larger version (27K): [in this window] [in a new window]   Fig. 5.   Overexpression of a dominant negative protein kinase B (Akt) blocks the PDGF-induced PPAR gene expression in VSMC. The confluent rat aortic smooth muscle cells were infected with an adenovirus carrying the gene for a dominant negative Akt. The cells were made quiescent after 24 h of adenovirus infection. A representative Northern blot is shown on the top panel. The average values of PPAR mRNA were normalized by GAPDH (bottom panel).

To confirm further the involvement of Akt in PDGF-induced PPAR expression in VSMC, we examined the PPAR protein levels in the rat vascular smooth muscle cell lines (A7r5) that were stably transfected with a constitutively active Akt versus control cells stably transfected with a GFP construct. We found that PPAR in Akt-A7r5 was easily detected by Western blot analysis but was undetectable in the control GFP-A7r5 (data not shown). These results further confirmed that Akt is involved in the regulation of PPAR expression in VSMC.

Construction of PPAR-A7r5 Cells-- We were intrigued by the initial observation that the rat embryonic aorta A7r5 clonal VSMC line failed to express PPAR by either Northern blot or Western blot analyses. This serendipitous finding of PPAR expression between A7r5 cells and RASMC provided us with the opportunity to examine PPAR-induced alterations in VSMC gene expression. To reconstitute PPAR expression in the A7r5 cells, retroviral expression vectors were used (Fig. 6A). The transfected cells had an ~4-kb transcript that contains a chimeric PPAR/GFP mRNA (Fig. 6B) that was translated into two separate proteins, GFP and PPAR (Fig. 6C). A homogeneous population of PPAR-A7r5 stable cells or the control A7r5 cells expressing GFP isolated by FACS was used for this study. There was no difference in size or morphology between PPAR-A7r5 and GFP-A7r5 cells (data not shown). This in vitro cell model system enabled us to define the role of PPAR in VSMC.

View larger version (24K): [in this window] [in a new window]   Fig. 6.   Construction of PPAR stable transfectants in A7r5 cells. A, the schematic diagram of PPAR stable expression vector. The mouse PPAR coding sequence was inserted upstream of the GFP gene. Expression was under the control of the cytomegalovirus (CMV) promoter. The internal ribosomal entry sequence (IRES) from encephalomyocarditis virus allowed the independent translation of PPAR and GFP from the bicistronic mRNA. LTR, long terminal repeat. B, Northern blot analysis of PPAR in A7r5 stable cells. Lane 1, GFP-A7r5 stable cells. Lane 2, PPAR-A7r5 stable cells. The 28 S ribosome RNA is shown in the bottom panel. C, Western blot analysis of PPAR in A7r5 stable cells. Identical blot was reprobed for actin to demonstrate the equal loading.

Overexpression of PPAR in VSMC Promotes Post-confluent Cell Proliferation-- To investigate the effect of PPAR overexpression in VSMC, we first examined the rate of cell growth in both PPAR-A7r5 and GFP-A7r5 cells by determining cell numbers. Although the growth rate of PPAR-A7r5 cells was similar to that of GFP-A7r5 cells before confluence, the PPAR-A7r5 cells grew significantly faster than GFP-A7r5 cells after confluence (Fig. 7A). Furthermore, the cell cycle distribution in PPAR-A7r5 and GFP-A7r5 cells was determined by flow cytometry analysis when the cells were quiescent and post-confluent. The percentage of S phase cells in PPAR-A7r5 was ~3.2-fold more than that in GFP-A7r5 (Fig. 7, B-D). We also examined the rate of apoptosis induced by serum withdrawal in both PPAR-A7r5 and GFP-A7r5 cells by both nuclear morphology and FACS analysis. There was no significant difference between PPAR-A7r5 and GFP-A7r5 cells (data not shown). Taken together, these results suggest that PPAR in VSMC is involved in cell proliferation.

View larger version (20K): [in this window] [in a new window]   Fig. 7.   PPAR promotes the G1  S progression in VSMC. A, overexpression of PPAR in VSMC promotes cell proliferation after confluence. PPAR-A7r5 and control GFP-A7r5 cells were plated in 6-well plates at a density of 1.6 × 105 cells/well. They were grown in DMEM/F-12 (Invitrogen) medium supplemented with 10% of fetal bovine serum. At defined time intervals, they were trypsinized and counted in a Coulter counter (model ZM Coulter Electronics Ltd., FL). B and C show the representative DNA histograms for the quiescent and post-confluent VSMCs. The GFP-A7r5 or PPAR-A7r5 cells were seeded in 6-well plates at a density of 5 × 105 cells/well. The cells were grown to confluence in DMEM/F-12 (Invitrogen) medium supplemented with 10% FBS. To make the cell quiescence, growth medium was removed and replaced with Opti-MEM (Invitrogen) for 48 h. 1 × 106 cells were analyzed by flow cytometry. D, mean percentage values of cells in S phase.

To investigate further the molecular basis that PPAR promotes VSMC growth, we analyzed the cell cycle proteins in both PPAR-A7r5 and GFP-A7r5 cells by Western blot analysis. As shown in Fig. 8, the levels of cyclin A and CDK2 proteins in PPAR-A7r5 cells were significantly higher than that in GFP-A7r5 cells in all growth conditions, and the level of CDK2 inhibitory protein, p57^kip2, was significantly lower in PPAR-A7r5 cells than in GFP-A7r5 cells after confluence. Interestingly, the level of CDK2 inhibitory protein, p27^kip1, was significantly higher in PPAR-A7r5 cells than in GFP-A7r5 cells after confluence, but there was no change in cyclin E levels between the two cell lines.

View larger version (61K): [in this window] [in a new window]   Fig. 8.   Overexpression of PPAR in VSMC alters cell cycle regulatory proteins. PPAR-A7r5 and the control GFP-A7r5 cells were plated in the T25 flask at a density of 5 × 105 cells/flask. They were grown in DMEM/F-12 (Invitrogen) medium supplemented with 10% fetal bovine serum. At defined stages as indicated, the cells were harvested, and 50 µg of protein extract was immunoblotted with the indicated antibodies. The relative values were normalized by actin. Three independent experiments showed similar results.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

It is postulated that pathological changes in vessel structures are induced in part by transcription factors that govern cell growth, death, differentiation, inflammation, and matrix production. PPAR and PPAR are members of a family of ligand-activated nuclear transcriptional factors that are emerging as important determinants of vascular function and structure. Activation of PPAR has been found to inhibit VSMC production of inflammatory factors (3). Recent studies have shown that the expression of PPAR was up-regulated in intimal VSMC (4), and activation of PPAR has been found to decrease both VSMC proliferation (32) and matrix production after vascular injury (5). Although it has been well documented that PPAR and PPAR are important determinants of vascular function and structure, the role of PPAR in vasculature is poorly understood.

In the present studies, we document that PPAR is expressed in VSMC and up-regulated by 1.7-, 3.1-, and 2.1-fold in rat carotid artery at 7, 14, and 28 days after injury, respectively. Interestingly, Admas et al. (31) showed that PPAR expression was up-regulated by 2.6-fold and was highest 4 h after injury, compared with control level, returned to the base line by 24 h, and did not change for 1 week, suggesting there is a biphasic increase in PPAR after vessel injury. A detailed time course of PPAR expression after vessel injury is currently ongoing to determine this contrasting finding in the two studies. In addition, further studies are required to understand the mechanism of this interesting phenomenon.

Vasculoproliferative disorders such as primary atherosclerosis, restenosis, and vein-graft failure are characterized by the accumulation of intimal smooth muscle cell proliferation, migration, and extracellular matrix deposition (17, 18). Cytokines and growth factors such as PDGF participate in these processes. We postulate that PDGF may up-regulate PPAR expression in VSMC. Indeed, we documented that PDGF induced PPAR expression in a time- and dose-dependent manner in VSMC. This further suggests that PPAR is involved in VSMC proliferation during vascular lesion formation.

PDGF is an important regulator that mediates the aberrant behavior of VSMC in the pathogenesis of vascular diseases. PDGF binding to its receptor on VSMC can activate several signaling pathways including p38-, MEK1/ERK-, and PI3-kinase-mediated pathways, which transduce the signals into nucleus and stimulate the proliferation and migration of VSMC (33, 34). In the current study, we demonstrated that PDGF-induced PPAR mRNA expression was most likely because of an induction of transcription rather than altering the stability of PPAR mRNA because the addition of PDGF failed to change the degradation rates of PPAR mRNA in VSMC. In addition, PDGF-induced PPAR mRNA expression did not require de novo protein synthesis because the addition of protein synthesis inhibitor in VSMC did not abrogate PDGF-induced PPAR expression. Taken together, the data suggest that there may be PDGF-response elements in the PPAR promoter.

The PPAR gene is composed of 9 exons spanning more than 85 kb on chromosome 6p21.2 (35). To date, little is known about PPAR transcriptional regulation. The only report on the transcriptional regulation of PPAR gene revealed that there are two putative -catenin/Tcf-4-binding sites located on the promoter (9). The up-regulation of PPAR mediated by -catenin/Tcf-4 was identified as one of the mechanisms involved in the initiation of colorectal tumors. Obviously, studying the transcriptional regulation of the PPAR gene such as systematic deletion mapping of PDGF-response elements in the PPAR promoter will not only help explain PPAR gene regulation but also provide new insights that will define the role of PPAR in vasculoproliferative disorders, diabetes, and cancer. Although this is beyond the scope of the present study, we have successfully cloned an ~5.5-kb human PPAR gene promoter, and studies are underway to determine the potential PDGF-response elements in the PPAR promoter.

We have shown that inhibition of PI3-kinase abrogates the effect of PDGF-induced PPAR expression in VSMC, although the pharmacological probe used was relatively selective and the results were verified using adenoviral vector with a dominant negative mutant Akt construct. The level of PPAR in Akt-A7r5 cells stably transfected with a constitutively active Akt construct was significantly increased.² Taken together, these results provide the first definitive evidence that PPAR gene expression is regulated by a PI3-kinase/Akt signaling pathway. However, the MEK1 inhibitor increased PDGF-induced PPAR mRNA expression. Further studies are required to clarify whether activation of the MEK1/ERK signaling pathway inhibits PDGF-induced PPAR expression and to determine precisely the relationship between PI3-kinase/Akt and MEK1/ERK signaling pathways in the regulation of PDGF-induced PPAR expression in VSMC.

Activation of MEK1/ERK signaling pathway could induced PPAR phosphorylation, resulting in a down-regulation of PPAR activity (36-38). In contrast, PPAR phosphorylation induced by MEK1/ERK pathway could enhance the PPAR activity (39). Interestingly, our data suggested that PDGF induced not only PPAR expression but also PPAR phosphorylation. More experiments are required to confirm PPAR phosphorylation and to elucidate its function.

Although we have demonstrated that PDGF-induced PPAR gene expression was mediated by the PI3-kinase/Akt-dependent pathway, it remained to be determined whether the PPAR target genes are activated following PDGF stimulation in VSMC. However, it is currently not practical to resolve this issue because the PPAR gene targets within VSMC have not been defined. To understand the role of PPAR in vasculature, it would be necessary to define globally the PPAR target genes in VSMC. Our laboratory is currently using a DNA microarray analysis to approach this challenge.

For many years, the lack of PPAR-specific ligands has hampered efforts to define PPAR function. It has been reported that carbaprostacyclin (cPGI), a stable analog of PGI₂, is a PPAR ligand (40). cPGI is a synthetic ligand that is structurally different from the endogenous PGI₂. And questions have been raised whether cPGI₂ itself can act as a bona fide ligand for PPAR. However, testing the ability of PGI₂ to activate PPAR in VSMC is difficult because of the inherent instability of this compound. In neutral or acidic buffers, PGI₂ is rapidly hydrolyzed (30-120 s) to 6-keto prostaglandin F₁ (10). To resolve this problem, there is a need to create an experimental model in which PGI₂ is functional as the endogenous ligand for PPAR. Because PGI₂ is the major prostanoid released by both endothelial cells (41) and VSMC (16) including A7r5 (42), we postulate that PPAR-A7r5 may be a useful cell model to test the PPAR function in VSMC. It is important to note that a recent intriguing report has identified the first high affinity PPAR ligand, GW501516, with an EC₅₀ = 1.2 ± 0.1 nM and >1,000-fold selective for PPAR over other subtypes (43). This will definitely spur new interest in the study of PPAR function.

Recent reports (10, 11) showed that PPAR promotes colon cancer proliferation and preadipocyte proliferation (12, 13). In the present study, we demonstrate that PDGF induces PPAR expression, and PPAR is up-regulated in neointima during vascular lesion formation. We hypothesize that overexpression of PPAR in VSMC is a sufficient condition to increase cell proliferation. Our results showed significantly faster growth rate in PPAR-A7r5 cells than the control GFP-A7r5 cells after confluence and no differences in growth rate between PPAR-A7r5 and GFP-A7r5 before cell confluence. This observation is consistent with a recent report (13) that PPAR promotes post-confluent cell proliferation in 3T3 fibroblasts. In addition, our data suggest that A7r5 can generate enough endogenous PPAR activators with comparable affinity to cPGI₂ because the addition of cPGI₂ into PPAR-A7r5 cells failed to alter the growth rate.²

VSMC proliferation is the major component of vasculoproliferative disorders. Vascular injury results in the release of growth factors and cytokines that stimulate quiescent, G₀/G₁-arrested VSMC to enter the cell cycle. Cell cycle progression is dependent on the expression and activation of specific enzymes, termed cyclin-dependent kinases (Cdks), which form complexes with their regulatory subunits, the cyclins. Activation of cyclin D + CDK4, cyclin D + CDK6, and cyclin E/A + CDK2 during G₁ phase results in G₁  S transition. Moreover, cyclin-dependent kinase inhibitors (p57^kip2, p27^kip1, and p21^cip1) are major negative regulators of the cell cycle by binding to and inhibiting the activation of CDK-cyclin complexes. In this study, we demonstrated that cyclin A and CDK2 in PPAR-A7r5 were significantly higher than that in GFP-A7r5. Consistent with this change, the level of p57^kip2 was significantly lower in PPAR-A7r5 than that in GFP-A7r5 cells after confluence. These changes could result in post-confluent VSMC proliferation by PPAR overexpression. Interestingly, the level of CDK2 inhibitory protein, p27^kip1, was significantly higher in PPAR-A7r5 than in GFP-A7r5 cells after confluence. Our results are consistent with recent studies (44, 45) that suggested that p21^cip1 and p27^kip1 of CDK function as positive regulators during the G₁ phase and as assembly factors to promote formation of cyclin-CDK holoenzyme complexes.

In summary, we report that PPAR is expressed in VSMC and up-regulated in neointima during vascular lesion formation. In addition, we demonstrate that PDGF-induced PPAR gene expression is mediated by PI3-kinase/Akt-dependent pathway. Overexpression of PPAR in VSMC promotes the post-confluent cell proliferation. Taken together, our results suggest that PPAR plays an important role in the modulation of vasculoproliferative disorders such as primary atherosclerosis, restenosis, and vein-graft failure.

	ACKNOWLEDGEMENTS

We thank Dr. Ogawa and Dr. Grimaldi for providing constructs for this study.

	FOOTNOTES

^* This work was supported in part by Starting Grant HL03676-02 from Morehouse Cardiovascular Research Institute, National Institutes of Health Grant NIHGMS S06GM08248, and the American Heart Association (YEC).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Both authors contributed equally to this work.

^§ To whom correspondence should be addressed: Cardiovascular Research Institute, Morehouse School of Medicine, 720 Westview Dr. S. W., Atlanta, GA 30310. Tel.: 404-752-1821; Fax: 404-752-1042; E-mail: echen@msm.edu.

Published, JBC Papers in Press, January 23, 2002, DOI 10.1074/jbc.M110580200

² J. Zhang, M. Fu, X. Zhu, Y. E. Chen, unpublished data.

	ABBREVIATIONS

The abbreviations used are: PPAR, peroxisome proliferator-activated receptor; PI3-kinase, phosphatidylinositol 3-kinase; VSMCs, vascular smooth muscle cells; PDGF, platelet-derived growth factor; PGI, prostacyclin I; cPGI, carbaprostacyclin I; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal bovine serum; RASMCs, rat aortic smooth muscle cells; RT, reverse transcriptase; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; nt, nucleotide; IRES, internal ribosomal entry sequence; FACS, fluorescence-activated cell sorter; GFP, green fluorescent protein; CDK, cyclin-dependent kinase; ERK, extracellular signal-regulated kinase; MEK, mitogen-activated protein kinase/ERK kinase.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

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