Platelet-Derived Growth Factor Promotes the Expression of Peroxisome Proliferator-Activated Receptor γ in Vascular Smooth Muscle Cells by a Phosphatidylinositol 3-Kinase/Akt Signaling Pathway
Vascular diseases such as atherosclerosis are characterized by abnormal accumulation of vascular smooth muscle cells (VSMCs) within the intimal lining. The intimal VSMCs exhibit an increased expression of peroxisome proliferator-activated receptor γ (PPARγ), and the administration of pharmacological PPARγ agonists attenuates vascular lesion formation. The factors that regulate PPARγ expression in the vasculature are poorly defined. Here we report that platelet-derived growth factor (PDGF) upregulates PPARγ by the phosphatidylinositol 3-kinase (PI3-kinase)/Akt signaling pathway. Using Northern-blotting and Western-blotting analyses, we observed that the levels of PPARγ mRNA and protein were increased by 2- to 3.5-fold in human aortic smooth muscle cells (HASMCs) treated with PDGF (20 ng/mL). This was abolished by preincubation of HASMCs with a PI3-kinase inhibitor (LY294002, 50 μmol/L), and partially inhibited by a MEK1 inhibitor (U0126, 10 μmol/L), but not affected by a p38 kinase inhibitor (SB202190, 10 μmol/L). In addition, overexpression of the dominant-negative p85 subunit of PI3-kinase or Akt proteins blocked the PDGF-induced PPARγ expression. Taken together, our results suggest that PDGF induces PPARγ expression in VSMCs by a PI3-kinase/Akt signaling pathway. The characterization of factors and signaling pathways that modulate PPARγ expression in VSMCs may have important implications for understanding the pathogenesis of vascular diseases.
- platelet-derived growth factor
- peroxisome proliferator-activated receptor γ
- phosphatidylinositol 3-kinase
- signaling pathway
- vascular smooth muscle cells
Peroxisome proliferator-activated receptors (PPARs), including α, γ, and β/δ, are a family of ligand-activated nuclear transcriptional factors1,2 that form heterodimers with retinoid X receptors (RXRα), bind to the PPAR responsive element (PPRE), and thereby regulate target gene expression. PPARγ is found predominantly in adipose tissue, where it plays a crucial role in adipocyte differentiation, fat storage, and glucose homeostasis.3 Recent studies have documented that PPARγ is also present in all of the following critical vascular cells: endothelial cells, vascular smooth muscle cells (VSMCs), and monocytes/macrophages.4,5 This finding suggested that PPARγ may play a critical role in vascular biology. Indeed, it has been reported that PPARγ is highly expressed in cells within the atherosclerotic plaque and the neointima after balloon injury.6 Thiazolidinediones, a class of antidiabetic drugs that are specific ligands of PPARγ, inhibit neointima formation after balloon injury7 and the development of atherosclerosis in LDL receptor-deficient mice.8 Studies in vitro also demonstrate that PPARγ agonists inhibit VSMC proliferation and migration.9,10 It is postulated that the increase in PPARγ expression noted in vascular lesion may function as an endogenous inhibitor of vascular disease. However, the factors that regulate PPARγ expression in vascular cells remain poorly defined.
Platelet-derived growth factor (PDGF) is a potent mitogen and chemoattractant that functions as an important mediator in the pathogenesis of vascular disease.11,12 In many pathological conditions, PDGF expression within vasculature is increased by a local production from endothelial cells and VSMCs and local secretion by platelets. Stimulation of the PDGF receptors on VSMCs can activate several signaling pathways, including those mediated by p38, MEK1/MAPK, and phosphatidylinositol 3-kinase (PI3-kinase), which transduce the signal into the nucleus and stimulate the proliferation and migration of VSMCs.13,14
In the present study, we tested the hypothesis that PDGF regulates PPARγ expression in human and rat VSMCs. Indeed, our findings indicate that PDGF upregulates PPARγ expression in VSMCs by a PI3-kinase/Akt signaling pathway.
Materials and Methods
Human aortic smooth muscle cells (HASMCs) were purchased from Clonetics and were cultured in smooth muscle cell growth medium-2 containing (in ng/mL) human basic fibroblast growth factor 2, human epidermal growth factor 0.5, and amphotericin-B 50; 5% FBS; 50 μg/mL gentamicin; and 5 μg/mL bovine insulin (all purchased from Clonetics). For all experiments, early-passage (passages 5 to 7) HASMCs were grown to 80% to 90% confluence and made quiescent by serum starvation (0.4% FBS) for at least 24 hours. Each inhibitor examined was added 30 minutes before the addition of human recombinant PDGF-BB (Sigma). Cells from the rat aortic smooth muscle cell line A7r5, obtained from American Type Culture Collection (ATCC), were cultured in DMEM containing 10% FBS (Life Technologies). The A7r5 cells stably overexpressing a constitutively active Akt construct (Akt/A7r5) and the control GFP (GFP/A7r5) were kindly provided by Dr Gary Gibbons (Cardiovascular Research Institute, Morehouse School of Medicine, Atlanta, Ga).
Northern Blot Analysis
Total RNA (20 μg) isolated from each condition 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 (Stratagene). 32P-labeled cDNA probes were generated by using a random primer labeling system (Gibco-BRL). Blots were prehybridized, hybridized, and washed once with 1× SSC at 65°C, and once with 0.1× SSC, 1.0% SDS (wt/vol), at 65°C for 20 minutes. The lane-loading differences were normalized using the GAPDH cDNA probe.
Analysis of PPARγ mRNA Half-Life
PPARγ mRNA half-life experiments were carried out using HASMCs. The cells were exposed to vehicle or PDGF (20 ng/mL) for the indicated periods before mRNA stability measurements. Transcription was inhibited by the addition of actinomycin D (5 μg/mL). Northern blotting analysis was performed as described above.
Western Blot Analysis
Total cell lysates (50 μg) were subjected to SDS-PAGE and electrotransferred to nitrocellulose membrane (Bio-Rad). After blocking in 20 mmol/L Tris-HCl (pH 7.6) containing 150 mmol/L NaCl, 0.1% Tween 20, and 5% (wt/vol) nonfat dry milk, blots were incubated with specific antibodies against PPARγ (Santa Cruz Biotechnology), PPARγ2 (Affinity Biotech), or p85 subunit of PI3-kinase (Upstate Biotechnology) for 1 hour at room temperature. The specificity of each antibody has been well documented by the manufacturers and confirmed by other published reports.6,15 The blots were incubated with horseradish peroxidase-conjugated secondary antibody (Santa Cruz Biotechnology), and immunoactivity was visualized using the enhanced chemiluminescence detection system (Amersham Pharmacia Biotech) per the manufacturer’s instructions.
Adenovirus Preparation and Infection
Adenovirus was prepared as previously described.16 For generation of recombinant adenovirus encoding dominant-negative p85 of PI3-kinase (named Ad-p85DN), a mutated form of p85 subunit I in which the inner SH2 domain is deleted17 was cloned into pCMVTrack (an adenovirus shuttle vector) and cotransformed with AdEasy (an adenovirus backbone) into Escherichia coli. The virus DNA from in vivo recombination was isolated, digested, and used for transfection of packing cell HEK293. A transfection mix was prepared by adding 4 μg of PacI linearized plasmid DNA and 20 μL of LipofectAMINE (Life Technologies) to 500 μL of OptiMEM (Life Technologies) according to the manufacturer’s instructions. The resulting virus was amplified for three to four cycles, and at this point, viral titers were high enough to use for gene-transfer experiments. The viruses were purified by CsCl gradient and final yields were generally 1011 to 1012 plaque-forming units (pfu)/mL. In this study, the VSMCs were infected with adenovirus vectors at ≈5 pfu/cell. The cells were subjected to experiments from 24 to 48 hours after infection.
To determine whether PI3-kinase is an essential signaling molecule mediating PDGF-induced PPARγ expression, we generated an adenovirus (Ad-p85DN) of the dominant-negative mutant 85-kDa subunit of PI3-kinase. To test the function of Ad-p85DN, we measured the PI3-kinase activity of this mutated p85 as previously described.15
Transient Transfection and Luciferase Assays
Transient transfection was performed with 1 μg of total DNA per well of six-well plates and LipofectAMINE (Gibco-BRL) according to the manufacturer’s instructions (DNA:LipofectAMINE ratio, 1:3). Pilot studies have documented a transfection efficiency of 15% to 25% in rat VSMCs (A7r5, ATCC) using this approach. Briefly, A7r5 cells achieving 70% confluence in six-well plates were cotransfected with 800 ng of an expression vector containing the reporter gene luciferase driven by a ≈3.0-kb PPARγ1 promoter18 (a gift from Dr J. Auwerx, IGBMC, C.U. de Strasbourg, France), and 200 ng of the expression vector containing the reporter β-gal driven by the cytomegalovirus promoter (Clontech) was used as the control for transfection efficiency. Twenty-four hours after transfection, the cells were washed twice with PBS and subsequently cultured for 24 hours in serum-free medium. After incubating with PDGF at 20 ng/mL for 6 hours, the cells were prepared for luciferase activity measurement using the reporter luciferase assay kit (Promega Co). The luciferase activity was measured by a luminometer (Victor II, Perkin Elmer) and normalized by β-gal activity.
Each experimental condition was tested in triplicate, and each experiment was repeated a minimum of three times. Statistical analyses were performed by ANOVA or unpaired two-tailed Student test. Data are presented as mean±SEM.
PDGF Induces PPARγ Expression in HASMCs
Although it was reported that PPARγ expression is upregulated in the neointima,6 little is known about the mechanism. To test whether PDGF regulates PPARγ gene expression in HASMCs, we examined the effect of PDGF on PPARγ gene expression in HASMCs using Northern blotting analysis for mRNA expression and Western blotting analysis for the protein level. In response to 20 ng/mL PDGF stimulation, the level of PPARγ mRNA was increased by ≈2.8-fold, and the PPARγ protein was increased ≈2.2-fold (Figures 1A and 1B).
To determine whether PDGF-induced PPARγ gene expression in VSMCs is at the transcriptional level, we transfected an expression vector containing the luciferase reporter gene driven by a ≈3.0-kb PPARγ1 promoter into rat VSMCs (A7r5, ATCC). PDGF stimulation for 24 hours increased the luciferase activity by ≈2.2-fold (Figure 1C). These results indicated that PDGF stimulates PPARγ gene transcription in VSMCs.
PPARγ has two isoforms, PPARγ1 and PPARγ2, generated by alternative promoters and differential splicing.18 PPARγ2 has 30 additional amino acids at the N-terminus and is reportedly expressed primarily in adipocytes. To date, whether PPARγ2 is expressed in VSMCs is controversal.6,9 To identify which PPARγ isoform was regulated by PDGF, we performed Western blotting analyses using an anti-PPARγ antibody (Santa Cruz) that cross-reacts with both PPARγ1 and PPARγ2 and an anti-PPARγ2 antibody (Affinity Biotech) that only reacts with PPARγ2. The protein from rat white fatty tissue was used as positive control. As shown in Figure 1D, there was only one band around 52 kDa detected by the anti-PPARγ antibody in PDGF-induced cell lysates, and no band was detected by the anti-PPARγ2 antibody in these samples. With the positive control, two bands (around 52 to 55 kDa) were detected in fat tissue by the anti-PPARγ antibody, whereas only one band (≈55 kDa) was detected by the anti-PPARγ2-selective antibody. In addition, PPARγ2 mRNA was not detected in HASMCs by reverse transcriptase-polymerase chain reaction assay using the human PPARγ2-specific primers as previously described19 (data not shown). These data suggested that only PPARγ1 was regulated by PDGF in VSMCs, and that PPARγ2 was undetectable in PDGF-treated or -untreated VSMCs.
Time Course and Dose-Dependent Effect of PDGF on PPARγ Gene Expression in VSMCs
HASMCs were treated with 20 ng/mL of PDGF for 0, 0.5, 2, 6, 12, 24, 48, and 72 hours, and then the levels of PPARγ mRNA in the cells were determined by Northern blotting analyses. As shown in Figure 2A, the levels of PPARγ mRNAs induced by PDGF were dramatically increased at 2 hours, but declined to some extent at 6 hours. At 12 hours, the mRNA levels rose again to achieve the second peak at 24 hours and a sustained level for at least 72 hours.
The dose-response effect of PDGF-induced PPARγ expression was documented at 2 hours of PDGF stimulation. The expression of PPARγ mRNA was upregulated in a dose-dependent manner, with significant increases observed at a concentration as low as 5 ng/mL. Maximal increases were obtained at a PDGF concentration of 10 ng/mL (Figure 2B). These results revealed that PDGF activates PPARγ gene expression in VSMCs.
PDGF Stimulation Does Not Affect the PPARγ mRNA Stability in VSMCs
To evaluate whether PPARγ mRNA stability contributes to PDGF-induced PPARγ gene expression, we examined the half-life of PPARγ mRNA in HASMCs. Northern blotting analyses were performed with addition of actinomycin D (5 μg/mL) after 2 hours of PDGF (20 ng/mL) stimulation. In HASMCs, the half-life of PPARγ mRNA was ≈4 hours. There was no significant difference between PDGF-treated and -untreated cells (Figure 3).
PDGF Induces PPARγ Expression by PI3-Kinase/Akt Signaling Pathway in VSMCs
To investigate the signaling pathways mediating PDGF-induced PPARγ expression, we initially focused on defining the roles of PI3-kinase, MEK/ERK, and p38 MAPK. HASMCs were treated with 20 ng/mL PDGF for 2 hours after pretreatment with LY294002 (50 μmol/L, a PI3-kinase inhibitor), SB202190 (25 μmol/L, a p38 kinase inhibitor), or U0126 (10 μmol/L, a MEK inhibitor). As shown in Figure 4, inhibition of PI3-kinase completely blocked the effect of PDGF (P<0.01), whereas inhibition of MEK partially reduced the effect of PDGF by 65±8.5% (P<0.05). However, inhibition of p38 MAPK had no significant effects on PDGF-induced PPARγ mRNA expression.
To further define whether the PI3-kinase signaling pathway mediates PDGF-induced PPARγ gene expression in VSMCs, we generated an adenoviral vector containing a dominant-negative mutant p85 (Ad-p85DN) in which the inner SH2 domain of p85 subunit is deleted. It has been documented that this mutant of p85 subunit cannot activate the p110 subunit of PI3-kinase. Using Western blotting analyses, we confirmed that this p85 dominant-negative protein was overexpressed in HASMCs that were infected with Ad-p85DN (Figure 5D). Additionally, Ad-p85DN blocked the PI3-kinase activity induced by insulin-like growth factor-1 (IGF-1) in HASMCs using thin-layer chromatography (Figure 5E). In contrast, Ad-GFP did not affect PI3-kinase activity induced by IGF-1. These results indicated that Ad-p85DN could function as a dominant-negative inhibitor of the PI3-kinase pathway.
To test the hypothesis that the stimulatory effect of PDGF is mediated by the PI3-kinase/Akt pathway, we selectively blocked this signaling pathway by using Ad-p85DN or ad-AktDN (a dominant-negative adenovirus of Akt from W.O.20). Blockade of the PI3-kinase/Akt pathway using this approach effectively prevented PDGF-induced PPARγ gene expression in HASMCs (Figure 5). However, PPARγ expression was not affected by the control adenovirus (Ad-GFP) infection in HASMCs (data not shown). Taken together, these results provided the first evidence that PPARγ gene expression is regulated by a PI3-kinase/Akt-dependent pathway.
To further confirm the involvement of Akt in PDGF-induced PPARγ expression in VSMCs, we examined the PPARγ protein levels in the rat VSMC lines (A7r5) that were stably transfected with a constitutively active Akt versus control cells stably transfected with a GFP construct. The level of the PPARγ in Akt/A7r5 is ≈2.3-fold higher than that in the control GFP/A7r5 VSMCs (Figure 6). These results further confirmed that Akt is involved in the regulation of PPARγ expression in VSMCs.
Peroxisome proliferator-activated receptors are a family of nuclear receptors comprising three subtypes of designated isoforms, PPARα, PPARγ, and PPARδ (also termed PPARβ, FAAR, or NUC-1) with different tissue distributions.16 PPARγ has been documented in the regulation of adipocyte differentiation, lipid metabolism, and insulin action.3 Several groups have reported that PPARγ is present in rat and human VSMCs,9,10 and is highly expressed within the atherosclerotic plaque and the neointima after balloon injury.6 However, little is known about the regulation of PPARγ expression in VSMCs. In this study, we demonstrated that PDGF induced PPARγ expression in VSMCs by a PI3-kinase/Akt-dependent signaling pathway.
PDGF-induced PPARγ mRNA expression is most likely due to an induction of transcription rather than altering the stability of PPARγ mRNA, given that the addition of PDGF failed to change the degradation rates of PPARγ mRNA in HASMCs. It is noteworthy that there are two peaks (at 2 and 24 hours) of PDGF-induced PPARγ gene expression. What is the mechanism of this biphasic induction? Recently, it was reported that angiotensin II induced cytoplasmic-to-nuclear translocation of the nuclear factor (NF)-κB subunits with parallel changes in DNA binding activity in a biphasic manner, which was comparable with the biphasic induction of interleukin-6 stimulated by angiotensin II.21 Because the PPARγ1 promoter contains NF-κB sites (data not shown), we hypothesize that NF-κB may be involved in mediating the PDGF-induced PPARγ expression in VSMCs. Although systematic deletion mapping of PDGF response elements in the PPARγ promoter is necessary to understand this mechanism, that is beyond the scope of the present study.
PDGF is an important regulator that mediates the aberrant behavior of VSMCs in the pathogenesis of vascular diseases.12 PDGF binding to its receptor on VSMC can activate several signal pathways including p38-, MEK1/ERK-, and PI3-kinase-mediated signal pathways, which transduce the signals into the nucleus and stimulate the proliferation and migration of VSMCs.13,14 Our experiments have shown that the inhibition of PI3-kinase completely blocked the effect of PDGF-induced PPARγ expression in VSMCs. The MEK1 inhibitor partially reduced the effect of PDGF on PPARγ mRNA, whereas p38 kinase inhibitors had no effect on this action. Although those pharmacological probes are relatively selective, we also verified these results by using adenoviral vectors with dominant-negative mutant constructs. The concordance between the data using pharmacological probes and the dominant-negative constructs provides the first definitive evidence that PPARγ gene expression is regulated by a PI3-kinase-dependent pathway.
It was well known that Akt is a proximal downstream effector of the PI3-kinase pathway.22 Therefore, we examined the involvement of Akt in PDGF-induced PPARγ expression. Our results showed that infection with adenovirus containing dominant-negative mutant of Akt also completely inhibited the increase of PPARγ expression induced by PDGF in HASMCs. Furthermore, as an additional test of our hypothesis, we confirmed that Akt activation is a sufficient condition for the upregulation of PPARγ gene expression. In stably transfected VSMCs with a constitutive upregulation of Akt activity, we observed that the level of PPARγ in Akt/A7r5 was ≈2.3-fold higher than the control VSMC line, GFP/A7r5. Taken together, these results demonstrated that Akt is a key mediator of PDGF-induced PPARγ expression in VSMCs.
Although we demonstrated that PDGF-induced PPARγ gene expression is mediated by PI3-kinase-dependent pathway, it would be quite interesting to determine whether the PPARγ target genes are activated after PDGF stimulation in VSMCs. We examined two well-characterized PPARγ target genes, CD36 identified in macrophages23 and uncoupling protein-2 (UCP-2)24 identified in adipocytes, because the PPARγ target genes are not well characterized in VSMCs. Using Western blotting analyses, we found that the level of UCP-2 was increased ≈2.6-fold after 48 hours of PDGF stimulation in HASMCs (data not shown). However, CD36 was undetectable by both reverse transcriptase-polymerase chain reaction and Western blotting analyses in HASMCs. These results suggested that PPARγ might be the mediator of PDGF-induced UCP-2 expression. To further understand the role of PPARγ in vasculature, it is necessary to globally define the PPARγ target genes in VSMCs. We are currently using a DNA microarray analysis to approach this challenge.
To date, several putative targets of PI3-kinase/Akt pathway have been proposed, among them Bad,25 caspase-9,26 and the transcription factor Forkhead.27 Interestingly, a recent report demonstrated that NF-κB is also the target of Akt,28 suggesting that NF-κB may be the downstream mediator of Akt in PDGF-induced PPARγ expression. To further study the mechanism of PDGF-induced PPARγ gene expression in VSMCs, we recently cloned the ≈5.4-kb human PPARγ1 gene promoter (data not shown). Computer sequence analysis revealed that there were several consensus-binding motifs, including the NF-κB, GATA1, C/EBP, and AP1 sites in this 5.4-kb PPARγ1 gene promoter. To our knowledge, there are no published studies that have analyzed these functional elements in the PPARγ1 gene promoter. Therefore, cloning this 5.4-kb PPARγ gene promoter will provide a unique resource for us to study the molecular mechanisms that govern the upregulation of PPARγ in VSMC during neointima formation.
PPARγ exists in at least two isoforms, PPARγ1 and PPARγ2, and PPARγ2 has an NH2-terminal extension of 30 amino acids. PPARγ2 is expressed selectively in adipose tissue, whereas PPARγ1 is present in many tissues. Previous reports have identified that PPARγ mRNA and protein exist in rat and human aortic VSMCs. However, the expression pattern of PPARγ isoforms was not described. In accord with a recent report,6 our findings indicate that only PPARγ1 is expressed in cultured human aortic VSMCs. Furthermore, we found that PDGF activates the PPARγ1 promoter activity as shown in Figure 1C, but not PPARγ2 promoter (data not shown).
It is well documented that PDGF is one of the key pathological factors in vascular diseases. Several recent reports suggest that PPARγ may function as a protective factor in vascular diseases such as atherosclerosis.29,30 Therefore, PDGF-induced PPARγ expression might provide a feedback mechanism by which PPARγ inhibits PDGF-induced VSMC proliferation and migration. The characterization of factors and signaling pathways that modulate PPARγ expression in vasculature may have important implications for the pathogenesis of vascular diseases.
Although the regulation of PPARγ gene expression was the focus in this study, we realized that the phosphorylation of PPARγ by mitogen-activated protein kinase at the N-terminal domain of PPARγ inhibits the transcriptional activity in adipocytes.31,32 In the present study, we have not detected the phosphorylated band of PPARγ in HASMCs. However, it will provide a better understanding of the role of PPARγ in vasculature to define whether other growth factors or cytokines can induce PPARγ expression, and whether phosphorylation of PPARγ is involved in its transcriptional activation in VSMCs.
This work was partially supported by a starting grant from Morehouse Cardiovascular Research Institute (Enhancement of Cardiovascular and Related Research Areas, NIH/NHLBI 5 UH1 HL03676-02) and an institutional grant (NIH/NIHGMS S06GM08248), as well as by the American Heart Association (to Y.E.C. and J.D.). We thank Dr Gary Gibbons for the useful discussions of this project and the critical review of this manuscript.
Original received May 7, 2001; revision received August 30, 2001; accepted September 21, 2001.
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