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Peroxisome Proliferator-activated Receptor γ Induces a Phenotypic Switch from Activated to Quiescent Hepatic Stellate Cells

2004; Elsevier BV; Volume: 279; Issue: 12 Linguagem: Inglês

10.1074/jbc.m310284200

1083-351X

Saswati Hazra, Shigang Xiong, Jiaohong Wang, Richard A. Rippe, Vibhor Krishna, Krishna Chatterjee, Hidekazu Tsukamoto,

Liver Disease Diagnosis and Treatment

Depletion of peroxisome proliferator-activated receptor γ (PPARγ) accompanies myofibroblastic transdifferentiation of hepatic stellate cells (HSC), the primary cellular event underlying liver fibrogenesis. The treatment of activated HSC in vitro or in vivo with synthetic PPARγ ligands suppresses the fibrogenic activity of HSC. However, it is uncertain whether PPARγ is indeed a molecular target of this effect, because the ligands are also known to have receptor-independent actions. To test this question, the present study examined the effects of forced expression of PPARγ via an adenoviral vector on morphologic and biochemical features of culture-activated HSC. The vector-mediated expression of PPARγ itself is sufficient to reverse the morphology of activated HSC to the quiescent phenotype with retracted cytoplasm, prominent dendritic processes, reduced stress fibers, and accumulation of retinyl palmitate. These effects are abrogated by concomitant expression of a dominant negative mutant of PPARγ that prevents transactivation of but not binding to the PPAR response element. PPARγ expression also inhibits the activation markers such as the expression of α-smooth muscle actin, type I collagen, and transforming growth factor β1; DNA synthesis; and JunD binding to the activator protein-1 (AP-1) site and AP-1 promoter activity. Inhibited JunD activity by PPARγ is not due to reduced JunD expression or JNK activity or to a competition for p300. But it is due to a JunD-PPARγ interaction as demonstrated by co-immunoprecipitation and glutathione S-transferase pull-down analysis. Further, the use of deletion constructs reveals that the DNA binding region of PPARγ is the JunD interaction domain. In summary, our results demonstrate that the restoration of PPARγ reverses the activated HSC to the quiescent phenotype and suppresses AP-1 activity via a physical interaction between PPARγ and JunD. Depletion of peroxisome proliferator-activated receptor γ (PPARγ) accompanies myofibroblastic transdifferentiation of hepatic stellate cells (HSC), the primary cellular event underlying liver fibrogenesis. The treatment of activated HSC in vitro or in vivo with synthetic PPARγ ligands suppresses the fibrogenic activity of HSC. However, it is uncertain whether PPARγ is indeed a molecular target of this effect, because the ligands are also known to have receptor-independent actions. To test this question, the present study examined the effects of forced expression of PPARγ via an adenoviral vector on morphologic and biochemical features of culture-activated HSC. The vector-mediated expression of PPARγ itself is sufficient to reverse the morphology of activated HSC to the quiescent phenotype with retracted cytoplasm, prominent dendritic processes, reduced stress fibers, and accumulation of retinyl palmitate. These effects are abrogated by concomitant expression of a dominant negative mutant of PPARγ that prevents transactivation of but not binding to the PPAR response element. PPARγ expression also inhibits the activation markers such as the expression of α-smooth muscle actin, type I collagen, and transforming growth factor β1; DNA synthesis; and JunD binding to the activator protein-1 (AP-1) site and AP-1 promoter activity. Inhibited JunD activity by PPARγ is not due to reduced JunD expression or JNK activity or to a competition for p300. But it is due to a JunD-PPARγ interaction as demonstrated by co-immunoprecipitation and glutathione S-transferase pull-down analysis. Further, the use of deletion constructs reveals that the DNA binding region of PPARγ is the JunD interaction domain. In summary, our results demonstrate that the restoration of PPARγ reverses the activated HSC to the quiescent phenotype and suppresses AP-1 activity via a physical interaction between PPARγ and JunD. Hepatic stellate cells (HSC) 1The abbreviations used are: HSC, hepatic stellate cell(s); PPARγ, peroxisome proliferator-activated receptor γ; PPRE, peroxisome proliferator-activated receptor response element; AP, activator protein, TGFβ and TGFβ1, transforming growth factor β and β1, respectively; RXRα, retinoid X receptor α; JNK, c-Jun N-terminal kinase; GST, glutathione S-transferase; MOI, multiplicity of infection; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal bovine serum; GFP, green fluorescent protein; CREB, cAMP-response element-binding protein; CBP, CREB-binding protein; HPLC, high pressure liquid chromatography. are vitamin A-storing pericytes in the subendothelial space of the liver. Upon injury to the liver, HSC become transdifferentiated into myofibroblastic cells to participate in wound healing (1Hautekeete M.L. Geerts A. Virchows Arch. 1997; 430: 195-207Crossref PubMed Scopus (267) Google Scholar). This transdifferentiation is characterized by reduced vitamin A content, increased cell proliferation and migration, enhanced matrix protein expression, and induced expression of α-smooth muscle actin (1Hautekeete M.L. Geerts A. Virchows Arch. 1997; 430: 195-207Crossref PubMed Scopus (267) Google Scholar). This response of HSC constitutes the normal, reparative homeostatic response of the liver to injury. However, dysregulation of HSC leads to excessive accumulation of extracellular matrices, resulting in liver fibrosis and cirrhosis. No curative medical treatments are available for cirrhosis except liver transplantation, and a precise understanding of transdifferentiation of HSC is the prerequisite for eventual identification of “dysregulation” and future developments of specific therapeutic modalities for the disease. To this end, much investigative effort has been made to characterize transcriptional regulation that underlies HSC transdifferentiation. Such examples include identification of Kruppel-like factor 6, a differentially expressed zinc finger protein in activated HSC in vitro and in vivo (2Kim Y. Ratziu V. Choi S.G. Lalazar A. Theiss G. Dang Q. Kim S.J. Friedman S.L. J. Biol. Chem. 1998; 273: 33750-33758Abstract Full Text Full Text PDF PubMed Scopus (230) Google Scholar). This transcription factor binds to the GC box sites of TGFβ1, TGFβ receptor type I and II (2Kim Y. Ratziu V. Choi S.G. Lalazar A. Theiss G. Dang Q. Kim S.J. Friedman S.L. J. Biol. Chem. 1998; 273: 33750-33758Abstract Full Text Full Text PDF PubMed Scopus (230) Google Scholar), urokinase-type plasminogen activator (3Kojima S. Hayashi S. Shimokado K. Suzuki Y. Shimada J. Crippa M.P. Friedman S.L. Blood. 2000; 95: 1309-1316PubMed Google Scholar), and α1(I) procollagen (2Kim Y. Ratziu V. Choi S.G. Lalazar A. Theiss G. Dang Q. Kim S.J. Friedman S.L. J. Biol. Chem. 1998; 273: 33750-33758Abstract Full Text Full Text PDF PubMed Scopus (230) Google Scholar) and induces transcription of these fibrogenic genes. The myofibroblastic phenotype seen in activated HSC is best characterized by induction of α-smooth muscle actin that is mediated by c-Myb binding to an E-box element in its promoter (4Buck M. Kim D.J. Houglum K. Hassanein T. Chojkier M. Am. J. Physiol. 2000; 278: G321-G328PubMed Google Scholar). The significance of this mode of regulation is supported by the demonstration of prevention of the myofibroblastic phenotypic switch by the treatment of HSC with antisense oligonucleotides for c-Myb (4Buck M. Kim D.J. Houglum K. Hassanein T. Chojkier M. Am. J. Physiol. 2000; 278: G321-G328PubMed Google Scholar). Sustained NF-κB activation confers activated HSC their proliferative and antiapoptotic status that may be important in progressive liver fibrogenesis (5Lang A. Schoonhoven R. Tuvia S. Brenner D.A. Rippe R.A. J. Hepatol. 2000; 33: 49-58Abstract Full Text Full Text PDF PubMed Scopus (149) Google Scholar). NF-κB may also mediate inflammatory responses by HSC via induction of chemokines and adhesion molecules (6Hellerbrand C. Jobin C. Licato L.L. Sartor R.B. Brenner D.A. Am. J. Physiol. 1998; 275: G269-G278Crossref PubMed Google Scholar, 7Knittel T. Dinter C. Kobold D. Neubauer K. Mehde M. Eichhorst S. Ramadori G. Am. J. Pathol. 1999; 154: 153-167Abstract Full Text Full Text PDF PubMed Scopus (117) Google Scholar). Increased activator protein-1 (AP-1) activity is essential for induction of matrix metalloproteinase (8Smart D.E. Vincent K.J. Arthur M.J. Eickelberg O. Castellazzi M. Mann J. Mann D.A. J. Biol. Chem. 2001; 276: 24414-24421Abstract Full Text Full Text PDF PubMed Scopus (89) Google Scholar), tissue inhibitor of matrix metalloproteinase-1, and interleukin-6 (9Vincent K.J. Jones E. Arthur M.J. Smart D.E. Trim J. Wright M.C. Mann D.A. Gut. 2001; 49: 713-719Crossref PubMed Scopus (28) Google Scholar) gene transcription in activated HSC, where JunD is shown to play a pivotal role (9Vincent K.J. Jones E. Arthur M.J. Smart D.E. Trim J. Wright M.C. Mann D.A. Gut. 2001; 49: 713-719Crossref PubMed Scopus (28) Google Scholar). A complexity in the understanding of HSC differentiation is underscored by different cellular phenotypes that HSC are shown to express. In addition to the myofibroblastic phenotype exhibited by activated HSC, they also express MyoD, the myogenic transcription factor specific for skeletal muscle (9Vincent K.J. Jones E. Arthur M.J. Smart D.E. Trim J. Wright M.C. Mann D.A. Gut. 2001; 49: 713-719Crossref PubMed Scopus (28) Google Scholar). Neuronal markers such as GFAP (10Neubauer K. Knittel T. Aurisch S. Fellmer P. Ramadori G. J. Hepatol. 1996; 24: 719-730Abstract Full Text PDF PubMed Scopus (161) Google Scholar), N-CAM (6Hellerbrand C. Jobin C. Licato L.L. Sartor R.B. Brenner D.A. Am. J. Physiol. 1998; 275: G269-G278Crossref PubMed Google Scholar), nestin (11Niki T. Pekny M. Hellemans K. Bleser P.D. Berg K.V. Vaeyens F. Quartier E. Schuit F. Geerts A. Hepatology. 1999; 29: 520-527Crossref PubMed Scopus (240) Google Scholar), and synaptophysin (12Cassiman D. van Pelt J. De Vos R. Van Lommel F. Desmet V. Yap S.H. Roskams T. Am. J. Pathol. 1999; 155: 1831-1839Abstract Full Text Full Text PDF PubMed Scopus (134) Google Scholar) are also expressed in HSC, suggesting the neural phenotype and that N-CAM and nestin are induced in activated HSC. Activated HSC express leptin (13Potter J.J. Womack L. Mezey E. Anania F.A. Biochem. Biophys. Res. Commun. 1998; 244: 178-182Crossref PubMed Scopus (189) Google Scholar), an adipocyte-specific gene, raising an intriguing possibility that HSC may also share the adipocytic phenotype. In fact, the quiescent HSC is laden with lipids including triglycerides, cholesterol, and phospholipids in addition to retinyl esters (14Yamada M. Blaner W.S. Soprano D.R. Dixon J.L. Kjeldbye H.M. Goodman D.S. Hepatology. 1987; 7: 1224-1229Crossref PubMed Scopus (93) Google Scholar). In support of this notion, peroxisome proliferator-activated receptor γ (PPARγ), one of the key transcription factors for adipocyte differentiation (15Spiegelman B.M. Flier J.S. Cell. 1996; 87: 377-389Abstract Full Text Full Text PDF PubMed Scopus (1157) Google Scholar), is expressed in the quiescent HSC (16Miyahara T. Schrum L. Rippe R. Xiong S. Yee Jr., H.F. Motomura K. Anania F.A. Willson T.M. Tsukamoto H. J. Biol. Chem. 2000; 275: 35715-35722Abstract Full Text Full Text PDF PubMed Scopus (434) Google Scholar, 17Galli A. Crabb D.W. Ceni E. Salzano R. Mello T. Svegliati-Baroni G. Ridolfi F. Trozzi L. Surrenti C. Casini A. Gastroenterology. 2002; 122: 1924-1940Abstract Full Text Full Text PDF PubMed Scopus (397) Google Scholar, 18Marra F. Efsen E. Romanelli R.G. Caligiuri A. Pastacaldi S. Batignani G. Bonacchi A. Caporale R. Laffi G. Pinzani M. Gentilini P. Gastroenterology. 2000; 119: 466-478Abstract Full Text Full Text PDF PubMed Scopus (368) Google Scholar), and its expression and activity decrease in HSC activation in vitro (16Miyahara T. Schrum L. Rippe R. Xiong S. Yee Jr., H.F. Motomura K. Anania F.A. Willson T.M. Tsukamoto H. J. Biol. Chem. 2000; 275: 35715-35722Abstract Full Text Full Text PDF PubMed Scopus (434) Google Scholar, 17Galli A. Crabb D.W. Ceni E. Salzano R. Mello T. Svegliati-Baroni G. Ridolfi F. Trozzi L. Surrenti C. Casini A. Gastroenterology. 2002; 122: 1924-1940Abstract Full Text Full Text PDF PubMed Scopus (397) Google Scholar) and in vivo (16Miyahara T. Schrum L. Rippe R. Xiong S. Yee Jr., H.F. Motomura K. Anania F.A. Willson T.M. Tsukamoto H. J. Biol. Chem. 2000; 275: 35715-35722Abstract Full Text Full Text PDF PubMed Scopus (434) Google Scholar). Further, the treatment of culture-activated HSC with the natural or synthetic ligands for PPARγ suppresses many functional parameters of the cell activation, including cell proliferation (17Galli A. Crabb D.W. Ceni E. Salzano R. Mello T. Svegliati-Baroni G. Ridolfi F. Trozzi L. Surrenti C. Casini A. Gastroenterology. 2002; 122: 1924-1940Abstract Full Text Full Text PDF PubMed Scopus (397) Google Scholar), expression of collagen, TGFβ, α-smooth muscle actin, monocyte chemotactic protein-1 genes, and chemotaxis (16Miyahara T. Schrum L. Rippe R. Xiong S. Yee Jr., H.F. Motomura K. Anania F.A. Willson T.M. Tsukamoto H. J. Biol. Chem. 2000; 275: 35715-35722Abstract Full Text Full Text PDF PubMed Scopus (434) Google Scholar, 18Marra F. Efsen E. Romanelli R.G. Caligiuri A. Pastacaldi S. Batignani G. Bonacchi A. Caporale R. Laffi G. Pinzani M. Gentilini P. Gastroenterology. 2000; 119: 466-478Abstract Full Text Full Text PDF PubMed Scopus (368) Google Scholar). More importantly, the treatment of the animal models of liver fibrosis with the PPARγ ligands ameliorates not only induction of fibrosis but also progression of preexisting fibrosis (17Galli A. Crabb D.W. Ceni E. Salzano R. Mello T. Svegliati-Baroni G. Ridolfi F. Trozzi L. Surrenti C. Casini A. Gastroenterology. 2002; 122: 1924-1940Abstract Full Text Full Text PDF PubMed Scopus (397) Google Scholar). Thus, these findings support the hypothesis that the maintenance of the quiescent state of HSC requires PPARγ and depletion of this adipogenic transcription factor underlies activation of HSC that can be circumvented by the ligand treatment. However, the ligands for PPARγ are also known to have receptor-independent effects. Using the embryonic stem cells from PPARγ null mice, neither macrophage differentiation nor anti-inflammatory effects of synthetic PPARγ ligands are shown to be dependent on PPARγ (19Chawla A. Barak Y. Nagy L. Liao D. Tontonoz P. Evans R.M. Nat. Med. 2001; 7: 48-52Crossref PubMed Scopus (966) Google Scholar). Indeed, the PPARγ ligand 15-deoxyprostaglandin J2 suppresses NF-κB activation by directly inhibiting IκB kinase in a PPARγ-independent manner (20Straus D.S. Pascual G. Li M. Welch J.S. Ricote M. Hsiang C.H. Sengchanthalangsy L.L. Ghosh G. Glass C.K. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 4844-4849Crossref PubMed Scopus (948) Google Scholar). Troglitazone also selectively induces early growth response-1 gene independently of PPARγ (21Baek S.J. Wilson L.C. Hsi L.C. Eling T.E. J. Biol. Chem. 2003; 278: 5845-5853Abstract Full Text Full Text PDF PubMed Scopus (175) Google Scholar). Mitogen-activated protein kinases such as c-Jun N-terminal kinase (JNK), p38, and extracellular signal-regulated kinase are activated by PPARγ ligands such as 15-deoxyprostaglandin J2 and ciglitazone in astrocytes and preadipocytes through the mechanisms that are independent of PPARγ but involving reactive oxygen species (22Lennon A.M. Ramauge M. Dessouroux A. Pierre M. J. Biol. Chem. 2002; 277: 29681-29685Abstract Full Text Full Text PDF PubMed Scopus (130) Google Scholar). Therefore, it is yet to be determined whether PPARγ directly inhibits HSC activation. In order to test this question, the present study was undertaken to express PPARγ via an adenoviral vector in culture-activated HSC and to determine its effects on the cell activation. Our results demonstrate that the restoration of PPARγ in activated HSC induces a reversal of the morphological features of HSC to the quiescent phenotype associated with inhibition of the known activation markers such as induced α-smooth muscle actin, collagen, and TGFβ expression; enhanced DNA synthesis; increased AP-1 binding; and promoter activity. Inhibition of AP-1 binding is due to PPARγ-mediated interference of JunD binding but not to suppression of JunD expression or JNK activity. Further, our results demonstrate a direct interaction of the PPARγ DNA binding domain with JunD that appears to underlie inhibited JunD binding to the AP-1 site. HSC Isolation and Culture—HSC were isolated from normal male Wister rats as previously described (23Tsukamoto H. Cheng S. Blaner W.S. Am. J. Physiol. 1996; 270: G581-G586PubMed Google Scholar) by the Non-Parenchymal Liver Cell Core of the USC-UCLA Research Center for Alcoholic Liver and Pancreatic Diseases. The use of animals for this study was approved by the Institutional Animal Care and Use Committee of the University of Southern California (protocol 9823). In brief, the liver was sequentially digested with Pronase and type IV collagenase to isolate nonparenchymal cells. These cells were subsequently subjected to arabinogalactan gradient ultracentrifugation to collect a pure fraction of HSC from the interface between the medium and a density of 1.035. The purity of isolated HSC was examined by phase-contrast microscopy, UV-excited fluorescence microscopy, and the viability by trypan blue exclusion. Isolated HSC from normal rats were cultured in a 100-mm plastic dish, 6- or 12-well plates with low glucose DMEM supplemented with 10% fetal bovine serum (FBS), 100 mg/ml streptomycin, 10,000 units/ml penicillin, and 25 μg/ml amphotericin B. The cultures were maintained for 7 days before the addition of viral vectors. In addition, a spontaneously immortalized HSC line established from experimental biliary liver fibrosis (24Xiong S. Yavrom S. Hazra S. Wu D. She H. Hepatology. 2001; 34 (abstr.): 520AGoogle Scholar) was maintained in culture in 6-well plates with low glucose DMEM supplemented with 10% FBS and used for a transient transfection experiment for AP-1 promoter activity. Construction of Viral Vectors and Transduction of the Viral Vector-mediated Genes—Full-length PPARγ1 cDNA was cloned from pCMX-PPARγ into the transfer vector, subsequently allowing homologous recombination with the pAdEasy-1 adenoviral plasmid containing GFP (Stratagene, La Jolla, CA). GFP alone was used as a control for PPARγ (Ad.GFP). An adenoviral vector expressing a dominant negative mutant form of PPARγ with a GFP tag (Ad.dn.PPARγ) was used. This dn.PPARγ carries the mutations at leucine and glutamic acid residues in the conserved AF-2 region of the C-terminal end. These mutations cause inhibition of recruitment of coactivators such as CBP and SRC-1 and ligand-dependent release of corepressors while promoting the basal recruitment of corepressors (25Gurnell M. Wentworth J.M. Agostini M. Adams M. Collingwood T.N. Provenzano C. Browne P.O. Rajanayagam O. Burris T.P. Schwabe J.W. Lazar M.A. Chatterjee V.K. J. Biol. Chem. 2000; 275: 5754-5759Abstract Full Text Full Text PDF PubMed Scopus (254) Google Scholar). All adenoviral vectors (Ad.GFP, Ad.PPARγ, and Ad.dn.PPARγ) were propagated using 293A cells in high glucose DMEM medium containing 5% FBS and then purified using cesium chloride gradient followed by dialysis for removing cesium chloride. The viral particle was titrated by the TCID50 (tissue culture infectious dose 50) method. Briefly, dilutions of viruses were incubated with 293-A cells in 96-well plates, and the presence or absence of cytopathic effect in each well was determined for titration. A multiplicity of infection (MOI) of 100 was used for the efficient tansduction of viral vector-mediated genes in activated HSC without any toxicity. HSC were cultured for 7 days. On day 7 of culture, cells were transduced with Ad.GFP, Ad.PPARγ, or Ad.dn.PPARγ or in combination using a total MOI of 100. The next day, medium was changed and cultured for another 4 days. DNA Synthesis—DNA synthesis was determined by the rate of [3H]thymidine incorporation into DNA. Activated HSC were cultured in 24-well plates (27,000 cells/well) and were transduced with Ad.GFP or Ad.PPARγ and cultured for an additional 5 days. On the 4th day after infection, [methyl-3H]thymidine (PerkinElmer Life Sciences) (1 μCi/ml) was added into each well and incubated overnight. After washing the cells, DNA was precipitated with 10% trichloroacetic acid at 4 °C. After several washes of the trichloroacetic acid precipitates, incorporation of [3H]thymidine into DNA was determined by counting the radioactivity of the precipitates using liquid scintillation counter. The count was standardized by the cell number. RNA Extraction and Real Time PCR—Total RNA was extracted from HSC transduced with Ad.GFP, Ad.PPAR plus Ad.GFP, or Ad.PPARγ plus Ad.dn.PPARγ using Trizol reagent (Invitrogen). Two micrograms of RNA were reverse-transcribed at 37 °C for 60 min with Moloney murine leukemia virus reverse transcriptase. For PCR analysis, the synthesized cDNA was amplified using primers for PPARγ, α1(I) procollagen, TGFβ1, and β-actin (16Miyahara T. Schrum L. Rippe R. Xiong S. Yee Jr., H.F. Motomura K. Anania F.A. Willson T.M. Tsukamoto H. J. Biol. Chem. 2000; 275: 35715-35722Abstract Full Text Full Text PDF PubMed Scopus (434) Google Scholar). For real time PCR, 2 ng of total RNA was used in a 20-μl reaction for reverse transcription for 50 min followed by 40 cycles of PCR to produce PCR products using the TaqMan Gold RT-PCR kit (Applied Biosystems, Foster city, CA). Probes were 5,6 carboxyl fluorescein amidite labeled at the 5′-end and black hole quencher-1 labeled at the 3′-end (Biosearch Technologies Inc., Novato, CA). Here, ABI 7700 SDS was used as a detection system. Each Ct value was first normalized to the respective glyceraldehyde-3-phosphate dehydrogenase Ct value of a sample and subsequently to a control sample. A difference in -fold was calculated from these Ct values. Electrophoretic Mobility Gel Shift Assay—HSC nuclear proteins (5–10 μg) or PPARγ, JunD, or RXRα in vitro translated using the coupled transcription/translation systems (Promega, Madison, WI) were incubated in a reaction mixture (20 mm HEPES, pH 7.6, 100 mm KCl, 0.2 mm EDTA, 2 mm dithiothreitol, 20% glycerol, 200 μg/ml poly(dI-dC)) on ice for 15 min and further incubated with 1–2 ng of α-32P-labeled doubled-stranded ARE-7 (the PPAR response element from the adipocyte fatty acid-binding protein gene) or AP-1 element (26Ohata M. Lin M. Satre M. Tsukamoto H. Am. J. Physiol. 1997; 272: G589-G596Crossref PubMed Google Scholar) for an additional 30 min. The reaction mixture was then resolved on a 6% nondenaturing polyacrylamide gel in 4× TBE (45 mm Tris, 45 mm boric acid, 1 mm EDTA). The gel was dried and subjected to autoradiography. For a supershift analysis, polyclonal antibodies against c-Fos, FosB, JunD, or PPARγ (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) were added and incubated for an additional 20 min after the 30-min incubation. Transient Transfection and Reporter Gene Assay—To determine whether PPARγ expressed by the adenoviral vector induces the PPAR response element (PPRE) promoter activity, HSC were transiently transfected with a PPRE-luciferase construct (tk-PPRE × 3-luciferase) using Targefect F-2 (Advanced Targeting Systems, San Diego, CA). To examine the effect of PPARγ expression on AP-1 activity, HSC transduced with Ad.PPARγ or Ad.GFP were transiently transfected with an AP-1 luciferase construct containing seven repeats of TGACTAA from the 12-O-tetradecanoylphorbol-13-acetate-responsive element (Stratagene, La Jolla, CA). We also examined the effects of CBP/p300 expression on the PPARγ-mediated effect on AP-1 by co-transfecting the Ad.GFP or Ad.PPARγ HSC line with a CBP/p300 expression plasmid (a kind gift from Dr. Stallcup, University of Southern California). For the determination of transfection efficiency, Renilla phRL-TK vector was used (Promega, Madison, WI). For transfection, 10-day cultures of HSC or HSC line in 6-well plates (70,000 cells/well; 3 days after infection with a viral vector) were incubated with 2 μg of each reporter construct, 0.02 μg of Renilla phRL-TK, and 2 μl of F-2 reagent in 1 ml of serum-free high glucose DMEM. Two h later, 1 ml of DMEM with 10% FBS was added to achieve the final FBS concentration of 5% for overnight incubation. On the next day, the medium was changed to DMEM with 10% FBS, and the cells were incubated for another 30 h. The cell lysate was collected for determination of both firefly and Renilla luciferase activities using the Dual-Luciferase reporter assay system (Promega). The results were normalized by Renilla luciferase activity. Morphological Analysis—To investigate the effects of PPARγ on the HSC morphology, 7-day cultured HSC were transduced with Ad.GFP, Ad.PPARγ plus Ad.GFP, or Ad.PPARγ plus Ad.dn.PPARγ using a total MOI of 100 of virus and cultured for an additional 5 days. The cells were examined under a fluorescent microscopy and photographed for documentation. The cells were also fixed in 3% paraformaldehyde in a phosphate buffer for 1 h and washed with a phosphate buffer. Stress fibers were subsequently stained with rhodamine-labeled phalloidin for 30 min and washed with a phosphate buffer, and differential interference contrast images were acquired to assess the effects of PPARγ expression. Retinyl Palmitate Measurement—To investigate the effect of PPARγ on the formation of intracellular lipid droplets, the cells were cultured with retinol (5 μm), palmitate (100 μm), or both for 48 h and stained with Oil Red O solution (in 60% isopropyl alcohol) for 20 min followed by counterstaining with hematoxylin. The stained slides were examined and photographed using an inverted light microscope. For chemical quantification of retinyl palmitate (a predominant vitamin A form stored in HSC), lipids were extracted with methanol and hexane from HSC treated with both retinol and palmitate (27Bhat P.V. Lacroix A. J. Chromatogr. 1983; 272: 269-278Crossref PubMed Scopus (53) Google Scholar). Retinyl palmitate in the hexane-extractable lipid phase was analyzed by reverse-phase high pressure liquid chromatography (HPLC) with the wavelength detector set at 325 nm. To prepare samples for HPLC analysis, solvent was evaporated from portions of lipid extracts in a 37 °C water bath using a gentle stream of N2. Samples were resolubilized in 1:1 methanol-chloroform solvent before HPLC. For data acquisition and peak area integration, the Winflow software package system (Inus Systems, Tampa, FL) was used. The data were standardized with the cell number. The HPLC analyses were performed at the Cell Biology Core of the USC Research Center for Liver Diseases. Western Blot Analysis of Cellular Proteins—HSC were cultured in a 100-mm dish for 7 days. On day 7, HSC were transduced with Ad.GFP or Ad.PPARγ using an MOI of 100 and cultured for additional 5 days. On day 12, cells were washed with PBS once and lysed with a lysis buffer (20 mm Tris-HCl, pH 7.6, 20 mm NaF, 20 β-glycerophosphate, 0.5 mm Na3VO4, 2.5 metabisulfite, 5 mm benzamidine, 1 mm EDTA, 0.5 mm EGTA, and 300 NaCl, with 10% glycerol and protease inhibitors and 1% Triton X-100). The lysates were stored at -80 °C until assayed. An equal amount of the whole cell protein (100 μg) was separated by SDS-PAGE and electroblotted to nitrocellulose filters. Proteins were detected by incubating the filter with monoclonal anti-α-smooth muscle actin (Sigma), rabbit polyclonal anti-type I collagen (Rockland Inc., Gilbertsville, PA), anti-PPARγ, or anti-JunD antibody (Santa Cruz Biotechnology) at a concentration of 0.2–2 μg/10 ml in TBS (100 mm Tris-HCl, 1.5 m NaCl, pH 7.4) with 5% nonfat milk followed by incubation with a horseradish peroxidase-conjugated secondary antibody (Santa Cruz Biotechnology) at 1 μg/10 ml. Proteins were detected by a chemiluminescent method using the PIERCE ECL kit (Amersham Biosciences). JNK Assay—To assay the activity of JNK, HSC transduced with Ad.GFP or Ad.PPARγ were lysed for collection of the whole cell proteins as described above. Each 100 μg of lysate was used to immunoprecipitate with 2 μg of anti-JNK1 antibody (Santa Cruz Biotechnology). Immunoprecipitates were then incubated with 2 μg of the N-terminal peptide of c-Jun (Santa Cruz Biotechnology) in 30 μl of kinase reaction buffer (20 mm Tris-HCl, pH 7.5, 20 mm MgCl2, 2 mm dithiothreitol) containing 20 μm ATP and [γ-32P]ATP (0.5 μCi) for 1 h at 30 °C. The protein mixture was resolved by 10% SDS-PAGE followed by a transfer onto a nitrocellulose membrane and exposure of the membrane to a PhosphorImager. Phosphate incorporated into c-Jun was visualized. Total JNK levels were determined by immunoblot analysis. PPARγ Immunoprecipitation and JunD Immunoblot Analysis—In order to assess the protein-protein interaction between PPARγ and JunD, in vitro translated PPARγ and JunD were incubated with an anti-PPARγ antibody (Santa Cruz Biotechnology) for 2 h. Protein A/G beads were added and incubated for another 24 h. Beads were washed three times and subjected to SDS-PAGE. Blots were probed with rabbit polyclonal anti-JunD antibody (2 μg/10 ml) in TBS containing 5% nonfat milk. Horseradish peroxidase-conjugated secondary antibody (1 μg/10 ml) was added and incubated followed by detection of JunD with the ECL reagent as described. Glutathione S-Transferase (GST) Pull-down Assay—Expression vectors for GST, GST-PPARγ (pGEX-4T2 and pGEX-4T2-PPARγ provided by Dr. Akira Sugawara, Tohoku University School of Medicine, Sendai, Japan), and GST fusion proteins of PPARγ deletion mutants (pGEX-4T1-PPARγ deletion mutants provided by Dr. Shigeaki Kato, University of Tokyo, Tokyo, Japan) were produced in Escherichia coli BL21. The glutathione beads were coated with the fusion proteins by incubating both together overnight at 4 °C. JunD protein was translated in vitro from pcDNA3-mJunD plasmid (kindly provided by Dr. Sunita K. Agarwal, National Institutes of Health, Bethesda, MD) in the presence of [35S]methionine using the TNT-T7-coupl