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(Circulation Research. 2002;91:e35.)
© 2002 American Heart Association, Inc.

UltraRapid Communication

PPAR Inhibits TGF-ß–Induced ß₅ Integrin Transcription in Vascular Smooth Muscle Cells by Interacting With Smad4

Ulrich Kintscher, Christopher Lyon, Shu Wakino, Dennis Bruemmer, Xu Feng, Stephan Goetze, Kristof Graf, Aristidis Moustakas, Bart Staels, Eckart Fleck, Willa A. Hsueh, Ronald E. Law

From the Department of Medicine (U.K., C.L., S.W., D.B., W.A.H., R.E.L.), Division of Endocrinology, Diabetes and Hypertension, University of California, Los Angeles, School of Medicine, Los Angeles, Calif; Institute of Pharmacology and Toxicology (U.K.), Charité Hospital, Humboldt-University Berlin, Berlin, Germany; the Department of Medicine/Cardiology (D.B., S.G., K.G., E.F.), German Heart Institute Berlin, Berlin, Germany; the Department of Pathology (X.F.), University of Alabama, Birmingham, Ala; Ludwig Institute for Cancer Research (A.M.), Uppsala, Sweden; the Department d’Atherosclerose (B.S.), UR545 INSERM, Institut Pasteur de Lille and Faculté de Pharmacie, Universitè de Lille II, Lille, France.

Correspondence to Ronald E. Law, PhD, UCLA School of Medicine, Division of Endocrinology, Diabetes and Hypertension, Warren Hall, Second Floor, Suite 24-130, 900 Veteran Ave, Box 957073, Los Angeles, CA 90095. E-mail rlaw{at}mednet.ucla.edu

	Abstract

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Integrins play an important role in vascular smooth muscle cell (VSMC) migration, a crucial event in the development of restenosis and atherosclerosis. Transforming growth factor-ß (TGF-ß) is highly expressed in restenotic and atherosclerotic lesions, and known to induce integrin expression. Peroxisome proliferator-activated receptor (PPAR), a member of the nuclear receptor superfamily, regulates gene expression in a variety of vascular cells. We investigated the effects of PPAR ligands on TGF-ß–induced ß₃ and ß₅ integrin expression and potential interaction between PPAR and TGF-ß signaling. PPAR ligands WY-14643 (100 µmol/L) and 5,8,11,14-eicosatetranoic acid (ETYA, 50 µmol/L) inhibited TGF-ß–induced ß₅ integrin protein expression by 72±6.8% and 73±7.1%, respectively (both P<0.05). TGF-ß–stimulated ß₃ integrin expression was not affected by PPAR ligands. Both PPAR ligands also suppressed TGF-ß–induced ß₅ integrin mRNA levels. PPAR ligands inhibited TGF-ß–inducible transcription of ß₅ integrin by an interaction with a TGF-ß response element between nucleotides -63 and -44, which contains a Sp1/Sp3 transcription factor binding site. Nuclear complexes binding to the TGF-ß response region contained Sp1/Sp3 and TGF-ß–regulated Smad 2, 3, and 4 transcription factors. TGF-ß–stimulated Sp1/Smad4 nuclear complex formation was inhibited by WY-14643 and ETYA with a parallel induction of PPAR/Smad4 interactions. However, in vitro pull-down experiments failed to demonstrate direct binding between PPAR/Smad4. Both PPAR ligands blocked PDGF-directed migration of TGF-ß–pretreated VSMCs, a process mediated, in part, by ß₅ integrins. The present study demonstrates that PPAR activators inhibit TGF-ß–induced ß₅ integrin transcription in VSMCs through a novel indirect interaction between ligand-activated PPAR and the TGF-ß–regulated Smad4 transcription factors. The full text of this article is available at http://www.circresaha.org.

Key Words: peroxisome proliferator-activated receptor • integrin • transforming growth factor-ß • vascular smooth muscle cell

	Introduction

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Migration and proliferation of vascular smooth muscle cells (VSMCs) are crucial events in the development of restenosis after vascular interventions and atherosclerosis.^1,2 Both require the interaction of cell surface integrin receptors with the surrounding extracellular matrix.^2,3 Integrins are a family of heterodimeric transmembrane glycoproteins consisting of noncovalently associated and ß chains.⁴ The integrin complexes _vß₃ and _vß₅ have been shown to be expressed on VSMCs and to regulate their migration through interactions with the extracellular matrix proteins vitronectin and osteopontin.^5,6 Furthermore, _vß₃ and _vß₅ are upregulated in atherosclerotic lesions and during neointima formation after vascular injury underscoring their important role in these processes.^7,8 Expression of integrins can be regulated by cytokines and growth factors present in atherosclerotic and neointimal lesions.⁹

Members of the transforming growth factor-ß (TGF-ß) family play a critical role in the regulation of cellular growth, differentiation, migration, and death.¹⁰ TGF-ß isoforms are highly expressed in human atherosclerotic lesions where they exert both pro- and antiatherosclerotic actions on vascular cells.^11,12 TGF-ß is also an important mediator of restenosis after vascular interventions.¹³ TGF-ß mediates its effects through receptor serine/threonine kinases at the cell surface and their substrates, the Smad proteins.¹⁰ On receptor activation, Smad2 and 3 are phosphorylated, resulting in complex formation with Smad4. These complexes translocate into the nucleus where they activate transcription of target genes.¹⁰ TGF-ß induces integrin expression and extracellular matrix production in a variety of cells including VSMCs.^14,15 More specifically, TGF-ß upregulates the VSMC expression of ß₃ integrin subunits and increases cell migration.¹⁶

Members of the peroxisome proliferator-activated receptor (PPAR) family are ligand-activated nuclear hormone receptors that function as transcriptional regulators of genes linked to lipid metabolism and glucose homeostasis.¹⁷ Three different isoforms have been identified: PPAR, PPAR, and PPAR.

PPAR is expressed in vascular cells including endothelial cells, monocyte/macrophages, and VSMCs.¹⁸ PPAR can be activated by hypolipidemic fibrates, eicosanoids, or polyunsaturated fatty acids.¹⁹ Activation of PPAR has been shown to inhibit proinflammatory processes in these cells, therefore, playing an important role in the development of atherosclerosis and restenosis after vascular injury.¹⁸ In VSMCs, PPAR ligands inhibited the expression of inflammatory genes such as interleukin 6,6-keto prostaglandin F₁ and cyclooxygenase-2.²⁰ Antiinflammatory actions of PPAR in VSMCs have been intensively studied.²¹ In contrast, the effects of PPAR ligands on other VSMC functions, including the regulation of genes involved in migration, are mainly unexplored.

PPARs are capable of both positive and negative regulation of gene expression in response to ligand binding.¹⁷ PPARs positively regulate gene expression by binding to PPAR response elements in target genes as a heterodimer with retinoid X receptors (RXRs) followed by the recruitment of coactivators and consequent transcription of the target genes.¹⁷ Negative regulation of gene expression by PPARs can be mediated by transrepression, a mechanism involving either competition for limiting amounts of essential coactivators utilized by many other transcription factors, or through direct physical interactions between PPARs and specific transcription factors.^22,23 Direct interactions with NF-B or AP-1 have been identified as a mechanism for inhibition of gene transcription by PPAR.²²

Given the importance of integrins in VSMC functions involved in atherosclerosis and restenosis, we investigated the effects of PPAR ligands on TGF-ß–induced ß₃ and ß₅ integrin expression in VSMCs. Furthermore, we identified potential novel interactions of ligand-activated PPAR with the TGF-ß–regulated transcription factor, Smad4.

	Materials and Methods

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Materials
Platelet-derived growth factor (PDGF), dimethylsulfoxide (DMSO), vitronectin, glutamine, and antibiotics were purchased from Sigma. Dulbeccos’s modified Eagle’s medium (DMEM), Opti-MEM I medium, Trizol Reagent, and LipofectAMINE 2000 were from Life Technologies. Fetal bovine serum (FBS) was purchased from Irvine Scientific. Hybond ECL nitrocellulose membrane, horseradish peroxidase-linked anti-rabbit and anti-mouse antibody, ECL Western blotting detection reagents, random prime labeling system (Redi prime II), rapid hybridization buffer, and nylon Hybond-N⁺ membranes were from Amersham Life Sciences. Dual-luciferase reporter assay and gel shift assay systems were from Promega. Cell fixation and staining was performed using the Quik-Diff stain set from DADE. TGF-ß₁ was from R&D Systems. WY-14643 and 5,8,11,14-Eicosatetraynoic acid (ETYA) were from BIOMOL. Antibodies were purchased from the following providers: ß₅ integrin (sc-5401), Sp1 (sc-420X, sc-59G), Sp3 (sc-644X), Smad2/3 (sc-6032X), Smad4 (sc-7154X), and PPAR (sc-9000X) were from Santa Cruz Biotechnology; ß₃ integrin (22440D) was from BD Pharmingen; ß₅ integrin (AB 1926) was from Chemicon International Inc; and normal rabbit IgG, normal goat IgG, and normal mouse IgG were from Zymed.

Cell Culture
Rat aortic smooth muscle cells (RASMCs) were prepared from thoracic aortae of 2 to 3 month old Harlan Sprague-Dawley rats by using the explant technique.²⁴ RASMCs were cultured as previously described.²⁵ WY-14643, ETYA, or vehicle DMSO were added to cells 30 minutes before the treatment with TGF-ß. For all data shown, each individual experiment represented in the n value was performed using independent preparations of RASMCs.

Western Blot Analysis and Immunoprecipitation
RASMC were harvested after stimulation with TGF-ß in the absence or presence of PPAR ligands. Western immunoblotting and immunoprecipitation experiments were performed as previously described.²⁶ Nuclear Extracts were prepared as described.²⁷

RNA Isolation and Northern Blot Analysis
After stimulation with TGF-ß±PPAR ligands, total RNA was isolated using Trizol reagent. RNA (20 µg) was electrophoresed through 1% agarose gels containing formaldehyde, transferred to charged nylon membranes, and hybridized with cDNA labeled with [-³²P] dCTP by random labeling. The hybridization signals of the specific mRNA of ß₅ integrin were normalized to those of CHOB, a constitutively expressed gene initially isolated as Chinese hamster ovary clone B that encodes a ribosomal protein, to correct for differences in loading or transfer. The ß₅ integrin cDNA was kindly provided by R.S. Ross (University of California, Los Angeles, Calif). All experiments were repeated at least 3 times with a different cell preparation. Band intensity was analyzed by densitometry.

ß₅ Promoter Luciferase Reporter Constructs
Generation of progressive 5'- to 3'-deletion constructs of murine ß₅ integrin promoters has been described previously, and were kindly provided by S.L. Cheng and F.P. Ross (Washington University, St Louis, Mo).²⁸ A 1-kb fragment was isolated from the AccI-digested products of the ß₅ genomic 9-kb fragment. Deletion constructs of this 1-kb fragment were obtained by using the Exonuclease III/Mung Bean Nuclease kit from Stratagene. All the fragments were subcloned into pGL3-basic vector containing the luciferase reporter gene. The promoter constructs used in this study spanned from -875, -63, and -43 to +110 from the transcription start site.

Transient Transfection and Luciferase Assay
RASMCs were transfected using LipofectAMINE 2000 for promoter activity analysis. Cells were grown to 70% to 80% confluence in 6-well plates and placed in Opti-MEM I medium. After 16 hours incubation, cells were transfected with 0.1 µg promoter construct to be tested and 5 ng pRL-CMV, a renilla luciferase control reporter vector. After 24 hours starvation in Opti-MEM I medium, cells were treated with WY-14643 and ETYA before stimulation with TGF-ß. After 24 hours, cells were lysed in reporter lysis buffer followed by measurement of luciferase activity using a Dual-Luciferase Reporter Assay System. Firefly luciferase activities from the ß₅ reporter constructs were normalized with renilla luciferase activities from pRL-CMV. All experiments were repeated at least 3 times with a different cell preparation.

Electrophoretic Mobility Shift Assay
For electrophoretic mobility shift assays (EMSA), radioactive double-stranded oligonucleotide -66/-42, labeled with T4 polynucleotide kinase and [-³²P] ATP, was incubated with nuclear extracts (2 µg) for 20 minutes in gel shift binding buffer (20% glycerol, 5 mmol/L MgCl₂, 2.5 mmol/L EDTA, 2.5 mmol/L DTT, 250 mmol/L NaCl, 50 mmol/L Tris-HCl [pH 7.5], and 0.25 mg/mL poly [dI-dC]). Assays were terminated by addition of 1 µL 10x gel loading buffer (250 mmol/L Tris-HCl [pH 7.5], 0.2% bromophenol blue, and 40% glycerol) and analyzed by electrophoresis using a 4% nondenaturing acrylamide gel (40:1 acrylamide: bisacrylamide) in 0.5x TBE-buffer. Gels were dried, and autoradiography was performed. For competitive oligonucleotide or immunodepletion assays, 100-fold unlabeled double-stranded oligonucleotides and 2 µg antibodies were incubated with nuclear extracts for 30 minutes before the addition of radiolabeled probe. For supershift-experiments, 2 µg of antibodies were added after incubation of the radiolabeled probe with nuclear proteins.

GST (Glutathione S-Transferase) Pull-Down Assay
The pSG5-hPPAR plasmid was described previously.²⁰ The pSG5-hPPAR1 plasmid was obtained from A. Elbrecht (Merck Research Laboratories). Expression vectors were in vitro transcribed and their transcripts in vitro translated in the presence of [³⁵S] methionine to label their recombinant protein products. GST-Smad3 and GST-Smad4 expression vectors were as previously published, and were produced in E. coli and purified using glutathione-Sepharose beads according to the manufacturer’s protocol (Amersham Life Sciences).²⁹ Approximately 10 µg of purified GST-fusion protein were mixed with 15 µL of [³⁵S] methionine-labeled PPAR or PPAR at 4°C in 500 µL of binding buffer (20 mmol/L Tris-HCl [pH 7.4], 100 mmol/L KCl, 0.7 mmol/L EDTA, 0.05% NP-40, 0.5 mmol/L PMSF, 10 µg/mL leupeptin and pepstatin, 2 µg/mL aprotinin) for 2 hours in the absence or presence of WY-14643 or rosiglitazone. Samples were centrifuged and washed 4 times to remove unbound protein. The washed beads were resuspended in 50 µL of SDS-PAGE loading dye and boiled to elute bound protein. Bound protein samples were size-fractionated by SDS-PAGE, stained with Coomassie Brilliant Blue R250, and analyzed by autoradiography.

Migration
RASMC migration was studied as previously described.³⁰ Transwell chambers were coated with vitronectin (10 µg/mL). Cells were pretreated with TGF-ß for 12 hours. PPAR ligands were added 30 minutes before stimulation with TGF-ß. For antibody experiments, RASMCs were incubated for 30 minutes with the indicated anti-integrin antibodies before plating in migration chamber.

Statistics
Analysis of variance was performed for statistical analysis, and values of P<0.05 were considered to be statistically significant. Data are expressed as mean±SEM.

	Results

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PPAR Ligands Inhibit TGF-ß–Induced ß₅ Integrin Expression
Treatment of quiescent RASMCs with TGF-ß induced ß₃ and ß₅ integrin protein expression by 2.7±0.7-fold and 3.3±0.5-fold, respectively, reaching a maximum at 5 ng/mL after 8 hours (P<0.01 versus untreated RASMCs) (Figures 1A through 1C). Neither higher concentrations of TGF-ß or longer periods of growth factor treatment further increased ß₃ and ß₅ integrin levels.

View larger version (31K): [in this window] [in a new window]   Figure 1. PPAR ligands inhibit TGF-ß–induced ß5 integrin expression. A and B, Quiescent RASMCs were treated with TGF-ß (5 ng/mL) for 8 hours in the absence or presence of the PPAR ligands WY-14643 (50 to 250 µmol/L) and ETYA (10 to 100 µmol/L). Representative immunoblots from 3 separate experiments are shown with antibodies that recognize ß5 integrin protein. *Unspecific band. Densitometric analysis of ß5 integrin protein levels are shown as percent of TGF-ß+DMSO–treated RASMCs. C, Quiescent RASMCs were treated with TGF-ß (5 ng/mL) for 8 hours ±PPAR ligands WY-14643 (100 µmol/L) and ETYA (50 µmol/L). Representative immunoblot from 3 separate experiments is shown with an antibody that recognizes ß3 integrin protein. D, Quiescent RASMCs were treated with TGF-ß (5 ng/mL) for 6 hours ±PPAR ligands WY-14643 (100 µmol/L) and ETYA (50 µmol/L). Representative Northern blots from 3 separate experiments showing ß5 integrin mRNA expression (Top) and CHOB mRNA levels (Bottom). Densitometric analysis of ß5 integrin mRNA levels are shown as percent of TGF-ß+DMSO–treated RASMCs. Experiments were repeated 3 times and results are presented as mean±SEM. *P<0.05, **P<0.01 vs TGF-ß/DMSO–treated RASMCs.

Pretreatment with the PPAR ligands WY-14643 (50 to 250 µmol/L) and ETYA (10 to 100 µmol/L) suppressed TGF-ß–mediated induction of ß₅ integrin expression reaching maximal inhibition at 100 µmol/L for WY-14643 (72±6.8% inhibition versus TGF-ß+DMSO; P<0.05) and at 50 µmol/L for ETYA (73±7.1% inhibition versus TGF-ß+DMSO; P<0.05) (Figures 1A and 1B). Similar concentrations of these PPAR-ligands have previously been used, without toxicity, in other studies, demonstrating inhibitory biological effects of PPAR-ligands.^22,31,32 The PPAR-ligands rosiglitazone (10 µmol/L) and troglitazone (10 µmol/L) had no effect on TGF-ß–induced ß₅ integrin protein expression (data not shown). TGF-ß–induced ß₃ integrin expression was not affected by either PPAR ligand (Figure 1C). The binding partner of ß₃/ß₅ integrins, namely _v integrin, was neither stimulated by TGF-ß nor regulated by the PPAR-ligand WY-14643 (100 µmol/L) (data not shown).

TGF-ß (5 ng/mL) also induced ß₅ integrin mRNA by 2.9±0.9-fold reaching a maximum after 6 hours (P<0.01 versus untreated RASMC) (Figure 1D). Both PPAR ligands blocked TGF-ß–induced ß₅ integrin mRNA expression with a maximal inhibition of 78.5±6.7% for WY-14643 (100 µmol/L; P<0.05 versus TGF-ß+DMSO) and 84.6±7.3% for ETYA (50 µmol/L; P<0.05 versus TGF-ß+DMSO) (Figure 1D).

PPAR Ligands Inhibit TGF-ß–Induced ß₅ Integrin Transcription
To determine whether the regulation of TGF-ß–induced ß₅ integrin mRNA by PPAR ligands resulted from an inhibition of gene transcription, RASMCs were transfected with a ß₅ integrin promoter (-875 to +110) region/luciferase reporter gene construct. TGF-ß (5 ng/mL) activated transcription from this ß₅-promoter region by 2.8±0.1-fold (P<0.05 versus untreated RASMCs) (Figure 2A). TGF-ß induction of this ß₅ integrin promoter activity was potently reduced by both PPAR ligands (WY 100 µmol/L, 1.2±0.4-fold activation versus untreated RASMCs; ETYA 50 µmol/L, 1±0.3-fold activation versus untreated RASMC; both P<0.05 versus TGF-ß+DMSO) (Figure 2A). These data suggest that PPAR ligands exert their inhibitory effects on ß₅ integrin mRNA expression through a transcriptional mechanism.

View larger version (34K): [in this window] [in a new window]   Figure 2. PPAR ligands inhibit TGF-ß–induced ß5 integrin transcription. A, RASMCs were transfected with the indicated ß5 integrin promoter luciferase constructs before the addition of TGF-ß (5 ng/mL) ±WY-14643 (100 µmol/L) or ETYA (50 µmol/L). After 24 hours stimulation, luciferase activity was measured and normalized with activity of cotransfected renilla luciferase. Results are shown as x-fold induction over untreated RASMCs. B, RASMCs were transfected with the ß5 integrin promoter luciferase constructs (-875/+110) and a pGL3-bascic vector. Experiments were performed as described under A. Experiments were repeated 3 times and results are presented as mean±SEM. #P<0.05 vs untreated RASMCs; *P<0.05 vs TGF-ß/DMSO–treated RASMCs.

Recently, a TGF-ß response element within the ß₅ integrin promoter (ß₅ TßRE) was identified between nucleotides -63 and -44.³³ In order to locate the region essential for the inhibitory actions of PPAR ligands on TGF-ß–induced ß₅ integrin promoter activity, RASMCs were transfected with ß₅ integrin promoter deletion constructs spanning from -63 to +110 and from -43 to +110. TGF-ß (5 ng/mL) induced ß₅ integrin promoter activity by 2.3±0.14-fold when the promoter construct was deleted to -63 (P<0.05 versus untreated RASMCs), whereas further deletion of the construct to -43 resulted in a complete loss of the inductive effects of TGF-ß, corroborating a major ß₅ TßRE between nucleotides -63 and -44 (Figure 2A). These data suggest that PPAR ligands inhibit TGF-ß–inducible transcription of ß₅ integrin by a potential interaction with the ß₅ TßRE between nucleotides -63 to -44.

Transient transfection of a pGL3-basic vector in RASMC revealed no regulation of luciferase activity compared with the ß₅ integrin promoter (-875 to +110) construct (Figure 2B).

PPAR Ligands Inhibit Binding of Nuclear Factors to the TGF-ß Response Element of the ß₅ Integrin Promoter
To confirm that the ß₅ TßRE between nucleotides -63 and -44 was indeed the target of PPAR ligands, EMSAs were performed with double-stranded radiolabeled (ß₅ TßRE) oligonucleotides corresponding to the -66/-42 region of the ß₅ integrin promoter. Treatment with TGF-ß (5 ng/mL) increased complex formation between nuclear proteins and the -66/-42 oligonucleotide by 1.8±0.1-fold reaching a maximum after 4-hour treatment (P<0.05 versus untreated RASMCs) (Figure 3A). Pretreatment with WY-14643 (100 µmol/L) and ETYA (50 µmol/L) in TGF-ß–treated cells decreased complex formation at the ß₅ TßRE region to basal levels (both P<0.05 versus TGF-ß+DMSO) (Figure 3A). These data demonstrate that PPAR ligands prevent the binding of transcription factors to the ß₅ TßRE.

View larger version (36K): [in this window] [in a new window]   Figure 3. PPAR ligands inhibit binding of nuclear factors to the TGF-ß response element of the ß5 integrin promoter. A, Quiescent RASMCs were treated with TGF-ß (5 ng/mL) for 4 hours ±WY-14643 (100 µmol/L) or ETYA (50 µmol/L). Nuclear extracts were isolated and incubated with radioactive end-labeled double stranded -66/-42 oligonucleotide. Complexes were separated on a 4% nondenaturing acrylamide gel. Cold -66/-42 oligonucleotide was added 30 minutes before the addition of the radioactive probe. Representative autoradiography from 3 separate experiments is shown. Densitometric quantitation is presented in the graph as x-fold induction over untreated RASMCs. Results are presented as mean±SEM. #P<0.05 vs untreated RASMC; *P<0.05, **P<0.01 vs TGF-ß/DMSO–treated RASMCs. B, Quiescent RASMCs were treated with TGF-ß (5 ng/mL) for 4 hours, and experiments were performed as described in A. Cold Sp-1– and AP2-consensus elements were added 30 minutes before addition of the radioactive probe. Representative autoradiography from 3 separate experiments is shown. C, Quiescent RASMCs were treated with TGF-ß (5 ng/mL) for 4 hours, and experiments were performed as described in A. Indicated antibodies were added after addition of the radioactive probe. Supershifted bands are indicated with a. Representative autoradiography from 3 separate experiments is shown.

TGF-ß Response Element of the ß₅ Integrin Promoter Binds Sp1/Sp3/Smad Proteins
Previous studies revealed the presence of a Sp1/Sp3 transcription factor binding site between nucleotides -53 and -48 in the ß₅ TßRE.³³ To identify transcription factors binding to the ß₅ TßRE region and to confirm the presence of a Sp1/Sp3 site within this region, a series of additional EMSA experiments was performed with labeled -66/-42 oligonucleotides. Consistent with the presence of a Sp1/Sp3 site, pretreatment of nuclear extracts with 100-fold excess unlabeled consensus Sp1 oligonucleotide decreased complex formation of nuclear extracts with the ß₅ TßRE probe (Figure 3B). Incubation with 100-fold consensus AP-2 oligonucleotide did not affect binding activity (Figure 3B). Treatment of nuclear extracts with anti-Sp1/Sp3 antibodies after addition of the radioactive probe induced a new, supershifted band, indicating the presence of these transcription factors in the complex binding to ß₅ TßRE (Figure 3C). Because Smad proteins are important mediators of TGF-ß signaling, nuclear extracts were incubated with anti-Smad 2, 3, and 4 antibodies after addition of labeled ß₅ TßRE oligonucleotide. Incubation with Smad 2, 3, and 4 antibodies also induced an additional supershifted band, demonstrating the presence of Smad transcription factors in nuclear complexes formed at the -66/-42 region (Figure 3C). Addition of anti-PPAR antibody showed no effect on the binding reaction, which indicates that the complexes does not contain PPAR (Figure 3C).

Ligand-Activated PPAR Inhibits Sp1/Smad4 Complex Formation by Interacting With Smad4
Immunoprecipitation experiments with nuclear extracts were performed to examine further potential mechanisms of PPAR’s inhibitory actions on Sp1/Sp3: Smad 2, 3, and 4 interactions at the ß₅ TßRE.

Given a central role of Smad4 in TGF-ß–mediated ß₅ integrin regulation, we focused our studies on this transcription factor.³³ Treatment with TGF-ß (5 ng/mL) for 4 hours increased binding of nuclear Smad4 to nuclear Sp1, whereas binding of Smad4 to Sp3 was not affected (Figure 4A and 4B). Immunoprecipitation with nonspecific anti-goat and anti-rabbit IgGs did not yield any Smad4 bands. Sp1/Sp3 partner proteins for Smad4 are efficiently immunoprecipitated by their corresponding antibodies (Figures 4A and 4B). Pretreatment with WY-14643 (100 µmol/L) and ETYA (50 µmol/L) inhibited TGF-ß–induced Smad4/Sp1 binding in nuclear extracts, suggesting a potential interaction between ligand-activated PPAR and Smad4/Sp1 complexes (Figure 4A). In contrast, Smad4/Sp3 complexes were not affected by either of the PPAR-ligands (Figure 4B). Cytosolic and nuclear protein levels of Sp1/Sp3 or Smad4 were not affected by PPAR-ligands (data not shown).

View larger version (27K): [in this window] [in a new window]   Figure 4. Ligand-activated PPAR inhibits Sp1/Smad4 complex formation by interacting with Smad4. Quiescent RASMCs were treated with TGF-ß (5 ng/mL) for 4 hours ±WY-14643 (100 µmol/L) or ETYA (50 µmol/L). Nuclear extracts were isolated and immunoprecipitated with either goat anti-Sp1 (A) or rabbit anti-Sp3 (B) or rabbit anti-PPAR (C). Representative immunoblots from 3 separate experiments are shown with antibodies that recognize Smad4 and as positive controls with the immunoprecipitation antibodies. For negative controls, samples were immunoprecipitated with nonspecific anti-goat or anti-rabbit IgG.

Because direct interactions between PPAR and transcription factors have been described recently, we hypothesized an interaction between ligand-activated PPAR and either Smad4 or Sp1. Treatment with TGF-ß (5 ng/mL) alone did not induce binding of PPAR to Smad4 (Figure 4C). Pretreatment of TGF-ß–stimulated cells with WY-14643 (100 µmol/L) or ETYA (50 µmol/L) led to enhanced formation of complexes containing PPAR and Smad4 (Figure 4C), whereas PPAR/Sp1 binding was not affected (data not shown). To further clarify whether PPAR directly interacts with Smad4, GST-pull-down experiments with in vitro translated PPAR protein and GST-Smad4 fusion proteins were performed. A direct interaction between PPAR and Smad4 could not be detected with or without the PPAR ligand WY-14643 (250 µmol/L) (Figure 5A). As a positive control we used in vitro translated PPAR, which is known to directly interact with Smad3 (Figure 5B). ³⁴ This interaction is enhanced by treatment with the PPAR ligand rosiglitazone (10 µmol/L) (Figure 5B).

View larger version (24K): [in this window] [in a new window]   Figure 5. PPAR does not directly bind to Smad4. A, In vitro translated PPAR protein was incubated with a GST-Smad4 fusion protein in the absence and presence of the PPAR ligand WY-14643 (250 µmol/L). Input of PPAR protein is shown in the left lane. Right two lanes show the GST-Smad4/PPAR sample. B, In vitro translated PPAR protein was incubated with a GST-Smad3 fusion protein in the absence and presence of the PPAR ligand rosiglitazone (10 µmol/L). Input of PPAR protein is shown in the left lane. Right two lanes show the GST-Smad3/PPAR sample.

In combination, these data demonstrate that ligand-activated PPAR blocks TGF-ß–induced Smad4/Sp1 interactions. Although a direct PPAR/Smad4 association could not be detected, these two proteins may be a part of a multiprotein complex that is the target for the observed inhibitory effects of PPAR ligands on TGF-ß–induced binding of Sp1/Sp3/Smad2,3,4 complexes to the ß₅ TßRE.

PPAR Ligands Inhibit PDGF-Directed Migration of TGF-ß–Pretreated RASMCs
To elucidate whether the inhibitory effects of PPAR ligands on TGF-ß–induced ß₅ integrin expression translated into an inhibition of RASMC migration, we studied the migratory response of TGF-ß–treated and nontreated cells toward PDGF in the absence or presence of PPAR ligands.

Pretreatment of RASMCs with TGF-ß for 12 hours increased PDGF-directed migration on vitronectin (10 µg/mL) by 2.7±1.2-fold at 5 ng/mL TGF-ß compared with untreated RASMCs (P<0.05 versus PDGF alone) (Figure 6A). Incubation of RASMCs with WY-14643 (100 µmol/L) and ETYA (50 µmol/L) attenuated PDGF-directed migration without reaching statistical significance (WY-14643, 24.3±5.9% inhibition; ETYA, 25±13% inhibition versus PDGF alone) (Figure 6B). The inhibitory effects of PPAR ligands WY-14643 and ETYA became more pronounced when RASMCs were pretreated with TGF-ß (5 ng/mL) for 12 hours. Pretreatment with PPAR ligands blocked migration of these cells, reaching maximal inhibition at 100 µmol/L WY-14643 and at 50 µmol/L ETYA (WY-14643 100 µmol/L, 65.5±1.1% inhibition; ETYA 50 µmol/L, 71±1.8% inhibition; P<0.05 versus TGF-ß–treated RASMC+PDGF/DMSO) (Figure 6B). To study the involvement of ß₃ and ß₅ integrins in the antimigratory actions of PPAR ligands, RASMCs were treated with the PPAR ligands followed by an incubation with either anti–ß₃ integrin antibody or anti–ß₅ integrin antibody, and migration experiments were performed. Both PPAR ligands potently blocked PDGF-directed migration under conditions of ß₅ integrin–dependent migration (RASMCs incubated with anti-ß₃ antibody) (WY-14643 100 µmol/L, 71.9±1% inhibition; ETYA 50 µmol/L, 77.5±1.7% inhibition; both P<0.01) (Figure 6C). However, PPAR ligands had no significant inhibitory effect under conditions of ß₃ integrin–dependent migration (RASMCs incubated with anti-ß₅ antibody), implicating that PPAR-ligands antimigratory actions are predominantly mediated through ß₅ integrins (Figure 6C).

View larger version (28K): [in this window] [in a new window]   Figure 6. PPAR ligands inhibit PDGF-directed migration of TGF-ß–pretreated RASMCs. Migration of RASMCs was induced by addition of PDGF (20 ng/mL) to the lower compartment in a modified Boyden chamber assay. Migration filters were coated with vitronectin (10 µg/mL). A, Quiescent RASMCs were treated with TGF-ß (0.5 to 10 ng/mL) for 12 hours. Migration of cells is shown as x-fold induction over untreated RASMCs. Experiments were repeated 3 times and were done in duplicate. Data are expressed as mean±SEM. *P<0.05 vs untreated RASMCs without PDGF; #P<0.05 vs untreated RASMCs with PDGF. B, Quiescent RASMCs were treated with WY-14643 (50 to 250 µmol/L), ETYA (10 to 100 µmol/L), or DMSO±TGF-ß (5 ng/mL) for 12 hours. Migration of cells is shown as percent of untreated RASMCs. Experiments were repeated 3 times and were done in duplicate. Data are expressed as mean±SEM. *P<0.05 vs untreated RASMCs. C, Quiescent RASMCs were treated with WY-14643 (100 µmol/L), ETYA (50 µmol/L), or DMSO+TGF-ß (5 ng/mL) (5 ng/mL) for 12 hours. RASMCs were incubated with anti-integrin antibodies for 30 minutes before migration experiments. Migration of cells is shown as % of anti-integrin–treated RASMCs. Experiments were repeated 3 times and were done in duplicate. Data are expressed as mean±SEM. **P<0.05 vs anti-integrin–treated RASMCs.

Together these data suggest that the inhibition of TGF-ß–treated RASMC migration by PPAR ligands, may result, at least in part, from their effect to inhibit TGF-ß–induced ß₅ integrin expression.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
The present study demonstrates that PPAR activators inhibit TGF-ß–induced transcription of ß₅ integrin in RASMCs. We further show that a PPAR/Smad4 association, which is likely mediated by additional nuclear proteins, may prevent formation of Sp1/Sp3/Smad2,3,4 complexes on a TGF-ß response element and constitutes a potential mechanism for PPAR ligand-mediated repression of ß₅ integrin promoter activity.

PPAR is expressed at substantial levels in VSMCs where it has been shown to function as a negative regulator of proinflammatory processes.²⁰ Its role in regulating VSMC migration, however, is largely unexplored. Integrins are important modulators of VSMC migration and are known to be regulated by TGF-ß.^2,3 We show that two different PPAR ligands inhibit TGF-ß–induced ß₅ integrin expression in RASMCs. To our knowledge, this is the first report describing the regulation of integrin expression by ligands of the PPAR class. Other nuclear hormone receptors, including the glucocorticoid receptor and the vitamin D receptor, have also been shown to regulate integrin expression in a variety of cells.^35,36

Recently, a TGF-ß response element has been identified between nucleotides -63 and -44 in the ß₅ integrin promoter in a murine osteoblastic cell line.³³ Transfection experiments in RASMCs revealed that the inhibitory effects of PPAR ligands on TGF-ß–induced ß₅ integrin expression are mediated by blocking transcriptional activity of the TGF-ß response element (-63/-44) in the ß₅ integrin promoter. Several molecular mechanisms can be invoked to explain the inhibitory actions of PPAR on gene transcription. Inhibition of transcription by PPAR might occur by competing for and sequestering of limiting transcriptional coactivators, such as CBP or p300.¹⁷ Delerive and colleagues²² demonstrated that direct physical interactions between PPAR and the p65 NF-B subunit or c-Jun account, at least in part, for PPAR-mediated repression of NF-B/AP-1–regulated gene transcription. Our data suggest a more specific effect of ligand-activated PPAR on the TGF-ß–regulated transcription factor Smad4 as the mechanism for repressing TGF-ß–stimulated ß₅ integrin transcription.

Previous reports have shown that Smad4 is required for TGF-ß–induced ß₅ integrin transcription.³³ On TGF-ß stimulation Smad4 binds to Smad 2 and 3, and translocates to the nucleus where it binds to Sp1 transcription factors resulting in increased binding of Sp1/Sp3/Smad2,3,4 complexes to a Sp1/Sp3 binding site within the TGF-ß response region of the ß₅ integrin promoter.³³ By gel shift experiments, we demonstrated that nuclear protein complexes binding to the TGF-ß response region in the ß₅ integrin promoter contain Sp1/Sp3 and Smad 2, 3, and 4 transcription factors. More importantly, we observed that PPAR ligands potently inhibited formation of Sp1/Smad4 complexes. Recently, Fu and colleagues³⁴ demonstrated that PPAR inhibits TGF-ß–induced gene expression by directly interacting with Smad3, suggesting that physical interplay between PPAR family members and Smad transcription factors may constitute a mechanism for PPAR-mediated inhibition of TGF-ß–regulated gene expression. It appears that Smad3 does not play a major role in regulating ß₅ integrin promoter activity, because its binding partner, ligand-activated PPAR, did not affect ß₅ integrin protein expression. In contrast, based on the central role for Smad4 in ß₅ integrin promoter regulation, sequestering of Smad4 by PPAR may prevent the formation of Sp1/Sp3/Smad2,3,4 complexes at -66/-42 of the ß₅ integrin promoter. Although we could not detect a direct protein-protein interaction between PPAR and Smad4 in pull-down experiments, PPAR ligands still induced nuclear PPAR/Smad4 association in coimmunoprecipitation experiments. This data are consistent with a study by Pouponnot and colleagues,³⁷ demonstrating an interaction of Smad4 with the coactivator p300 in coimmunoprecipitation experiments, whereas a direct protein-protein interaction was not detected in experiments with a GST-Smad4 fusion protein. There are two potential explanations for these data. Smad4 may have a weak affinity to its binding partners, which mitigates the detection of an interaction in in vitro assays. Alternatively, Smad4 interactions with its binding partners may be mediated by association with additional nuclear proteins present within a multiprotein complex. Additional studies will be required to identify other proteins present in a PPAR/Smad4 multisubunit complex.

Interestingly, treatment of RASMCs with PPAR ligands did not affect TGF-ß–induced ß₃ integrin expression. Little is known about ß₃ integrin regulation by TGF-ß. ß₃ integrin mRNA expression is upregulated by TGF-ß.¹⁶ Although the ß₃ integrin promoter contains binding sites for a number of transcription factors (Sp1, AP-1, STAT, and NF-B) known to be regulated by PPAR, the role of those elements in TGF-ß–regulated transcription from the ß₃ integrin promoter is still unknown.³⁸ Our findings predict, that Smad4 proteins are not involved in TGF-ß–stimulated ß₃ integrin transcription. Future studies are required to more fully elucidate transcriptional mechanisms involved in TGF-ß–mediated ß₃ integrin expression.

PPAR-mediated interference with Smad4 activity may be functionally significant. PPAR-ligands inhibited migration of TGF-ß–treated RASMCs, at least in part, by their inhibitory effects on TGF-ß–induced ß₅ integrin expression. These data demonstrate that the inhibitory actions of PPAR on ß₅ integrin expression translates into the blockade of an important integrin-mediated VSMC behavior. PPAR ligands did not affect PDGF-directed migration of untreated RASMCs. This finding suggests that PPAR ligands do not inhibit VSMC migration through a generalized effect on cell movement or chemotaxis. Instead, integrin upregulation by TGF-ß was shown to be the specific pathway targeted by PPAR. Antimigratory activity of PPAR ligands, therefore, may be limited to pathophysiological states characterized by elevated levels of TGF-ß. Marx and colleagues³¹ and our group³⁰ showed that activation of PPAR blocks VSMC migration, which was associated with a blockade of matrix metalloproteinase (MMP)-9 activity. Interestingly, Marx and colleagues did not observe any effects of PPAR ligands on VSMC MMP-activity.³¹ These data suggest that PPAR inhibits VSMC migration by targeting a matrix-degrading, invasive component of the migratory process, whereas PPAR ligands are affecting integrin-mediated cell movement.

Beside their important role in cell migration, integrins are also involved in the regulation of cell proliferation.³⁹ The migratory and proliferative response of VSMCs after vascular injury are major contributors to the development of intimal hyperplasia followed by restenosis of the injured vessel.^1,2 Blocking of integrin function with anti-integrin antibodies has been shown to reduce postinjury restenosis in animals and decreased ischemic long-term complications after vascular interventions in humans.^40,41 In addition, blocking TGF-ß expression or signaling by ribozyme oligonucleotides against TGF-ß, soluble TGF-ß type II receptor, or anti–TGF-ß antibodies has been shown to attenuate neointima formation after vascular injury.^13,42,43 Decreased intimal lesion formation in these studies resulted from an inhibition of TGF-ß–mediated VSMC migration and/or proliferation as well as matrix accumulation. Inhibition of TGF-ß–mediated ß₅ integrin expression by activation of PPAR may, therefore, have beneficial effects on the development of postinjury intimal hyperplasia.

	Acknowledgments

 
This study was supported by an NIH Grant to W.A.H. (HL-58328-03). U.K. and D.B. are supported by a research fellowship by the Gonda (Goldschmied) Diabetes Center, University of California, Los Angeles. S.W. is supported by a fellowship by the Mary K. Iacocca Foundation. D.B. is supported by a grant from MSD Sharp&Dohme.

Received July 12, 2001; revision received October 16, 2002; accepted October 29, 2002.

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