Ligands of peroxisome proliferator-activated receptor γ modulate profibrogenic and proinflammatory actions in hepatic stellate cells

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Ligands of Peroxisome Proliferator–Activated Receptor ␥ Modulate Profibrogenic and Proinflammatory Actions in Hepatic Stellate Cells FABIO MARRA,* EVA EFSEN,* ROBERTO G. ROMANELLI,* ALESSANDRA CALIGIURI,* SABRINA PASTACALDI,* GIACOMO BATIGNANI,‡ ANDREA BONACCHI,* ROBERTO CAPORALE,§ GIACOMO LAFFI,* MASSIMO PINZANI,* and PAOLO GENTILINI* *Dipartimento di Medicina Interna and ‡Dipartimento di Fisiopatologia Clinica, Universita` di Firenze, Florence, Italy; and §Divisione di Ematologia, Azienda Ospedaliera Careggi, Florence, Italy

Background & Aims: Proliferation and migration of hepatic stellate cells (HSCs) and expression of chemokines are involved in the pathogenesis of liver inflammation and fibrogenesis. Peroxisome proliferator–activated receptor (PPAR)-␥ is a receptor transcription factor that controls growth and differentiation in different tissues. We explored the effects of PPAR-␥ agonists on the biological actions of cultured human HSCs. Methods: HSCs were isolated from normal human liver tissue and used in their myofibroblast-like phenotype or immediately after isolation. Activation of PPAR-␥ was induced with 15-deoxy-⌬12,14-prostaglandin J2 or with troglitazone. Results: PPAR-␥ agonists dose-dependently inhibited HSC proliferation and chemotaxis induced by platelet-derived growth factor. This effect was independent of changes in postreceptor signaling or expression of c-fos and c-myc and was associated with inhibition of cell cycle progression beyond the G1 phase. Activation of PPAR-␥ also resulted in a complete inhibition of the expression of monocyte chemotactic protein 1 at the gene and protein levels. Comparison of quiescent and culture-activated HSCs revealed a marked decrease in PPAR-␥ expression in activated cells. Conclusions: Activation of PPAR-␥ modulates profibrogenic and proinflammatory actions in HSCs. Reduced PPAR-␥ expression may contribute to confer an activated phenotype to HSCs.

epatic stellate cells (HSCs) represent the key cellular elements in the liver wound healing process and development of hepatic fibrosis.1 The ability of HSCs to modulate liver tissue repair is dependent on a process known as activation.1,2 Upon liver injury, HSCs acquire the ability to proliferate and migrate toward the damaged areas and increase the production of extracellular matrix components. In addition, activated HSCs regulate the recruitment of inflammatory cells via secretion of chemotactic factors, including chemokines, and immunomodulatory cytokines such as interleukin (IL)-10.3

H

These properties of the activated or myofibroblast-like phenotype of HSCs contribute to the morphologic and functional changes observed during chronic liver damage. The molecular mechanisms regulating the transition toward myofibroblast-like cells are only partially understood. Because the activation process is associated with de novo expression of specific genes, attention has focused on molecules capable of regulating gene transcription. Generation of lipid peroxidation products from damaged hepatocytes has been suggested to promote HSC activation via transcription factors such as c-myb and nuclear factor–␬B (NF-␬B).4 More recently, it has been reported that Zf9, a Kruppel-like transcription factor, is rapidly induced during the process of HSC activation in vivo and in vitro and transactivates the promoters of genes up-regulated during fibrogenesis, including type I collagen, transforming growth factor ␤, and its receptors.5,6 In addition, it is known that HSC activation is associated with loss of retinoids,7,8 a group of molecules with potent effects on gene transcription and cell differentiation. Peroxisome proliferator–activated receptors (PPARs) are a family of ligand-activated nuclear transcription factors belonging to the nuclear hormone receptor superfamily.9 Three mammalian subtypes have been identified, referred to as PPAR-␣, -␤ (or -␦), and -␥, which are encoded by separate genes.10 Transcriptional regulation by PPAR occurs through binding to specific regulatory Abbreviations used in this paper: 15d-PGJ2, 15-deoxy-⌬12,14-prostaglandin J2; ERK, extracellular signal–regulated kinase; HSC, hepatic stellate cell; IFN, interferon; IL, interleukin; MCP-1, monocyte chemotactic protein 1; NF-␬B, nuclear factor–␬B; PDGF, platelet-derived growth factor; PPAR-␥, peroxisome proliferator–activated receptor ␥; RXR, retinoid X receptor; SDS-PAGE, sodium dodecyl sulfate–polyacrylamide gel electrophoresis; TNF, tumor necrosis factor. © 2000 by the American Gastroenterological Association 0016-5085/00/$10.00 doi:10.1053/gast.2000.9365

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elements located in noncoding regions of the gene. Binding to the regulatory element requires the formation of a heterodimeric complex comprised of a PPAR and the retinoid X receptor (RXR), another member of the same family of transcription factors.11 The 3 PPAR subtypes differ in terms of ligand specificity. Agonists of PPAR-␥ include oxidative metabolites of polyunsaturated fatty acids,12 and prostaglandins (PG) of the J series, including 15-deoxy-⌬12,14-PGJ2 (15d-PGJ2), which is by far the most potent activator.13,14 Although the precise enzymatic pathway leading to 15d-PGJ2 is not completely understood, it has been shown that these compounds are produced in intact cells and organisms15 and are likely to represent physiologic ligands for PPAR-␥. PPAR-␥ is also bound and activated by antidiabetic drugs of the thiazolidinedione group, such as troglitazone, and by some nonsteroidal anti-inflammatory drugs.16,17 Activation of PPAR-␥ has been shown to play a leading role in the process of adipocyte differentiation and glucose metabolism, and thiazolidinedione analogues are used as antidiabetic drugs in the clinical practice.18,19 Studies show that this transcription factor regulates neoplastic cell growth and modulates monocyte activation and differentiation.20 –23 Because PPAR-␥ has the ability to control gene transcription and cellular differentiation, we evaluated whether PPAR-␥ agonists could modulate the biological actions that characterize the activated phenotype of HSCs. We report that treatment of cultured human HSCs with ligands of PPAR-␥ inhibits cell proliferation, migration, and chemokine expression, 3 actions relevant to the process of liver wound healing and fibrogenesis.

Materials and Methods Reagents 15d-PGJ2 was purchased from Cayman Chemical Co. (Ann Arbor, MI). Aliquots of the original solution in ethyl acetate were dried under a nitrogen stream and resuspended in dimethyl sulfoxide. Troglitazone was kindly provided by Dr. Toshihiko Hashimoto (Sankyo Co. Ltd., Tokyo, Japan) and dissolved in dimethyl sulfoxide. The final concentration of dimethyl sulfoxide was maintained equal to 0.1% (vol, vol) in all experimental conditions. Monoclonal, agarose-conjugated antiphosphotyrosine antibodies were purchased from Calbiochem (La Jolla, CA). Phosphospecific antibodies against extracellular signal–regulated kinase (ERK) and p38MAPK were purchased from New England Biolabs (Beverly, MA). Polyclonal antibodies directed against ERK, p38MAPK, and PPAR-␥ were purchased from Santa Cruz Biotechnology (Santa Cruz, CA), and monoclonal antiphosphotyrosine antibodies (clone 4G10) from Upstate Biotechnology Inc. (Lake Placid, NY). The rabbit antiserum against baboon monocyte chemotactic protein

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(MCP)-1 (100% cross-reacting with human MCP-1) was a kind gift of Dr. Anthony J. Valente (University of Texas Health Science Center at San Antonio, TX). The luciferase reporter plasmid under the control of PPAR-␥ (ACO-PPRE) was generously provided by Dr. Brian Seed (Massachusetts General Hospital, Boston, MA). Phosphatidylinositol was from Sigma Chemical Co. (St. Louis, MO). Protein A–Sepharose was purchased from Pharmacia (Uppsala, Sweden). [␥-32P]Adenosine triphosphatase (3000 Ci/mmol) and [methyl3H]thymidine were from New England Nuclear (Milan, Italy). Human recombinant platelet-derived growth factor (PDGF)BB, IL-1␣, tumor necrosis factor (TNF)-␣, and interferon (IFN)-␥ were from Peprotech (London, England). All other reagents were of analytical grade.

Isolation and Culture of HSCs Human HSCs were isolated from wedge sections of liver tissue unsuitable for transplantation by collagenase/pronase digestion and centrifugation on stractan gradients. Procedures used for cell isolation and characterization have been described extensively.24 Purity of the cell was ⬎95%. Unless indicated otherwise, all the experiments were conducted on cells cultured on uncoated plastic dishes (passage 3– 6), showing an activated or myofibroblast-like phenotype. To assess the levels of PPAR-␥ in the activation process, HSCs were washed and the pellet was frozen in liquid nitrogen and stored at ⫺80°C until lysis and quantified for protein concentration as indicated later. Aliquots of freshly isolated cells (4 ⫻ 106 cells) were seeded onto 100-mm petri dishes and cultured in complete medium for 3 days or until the second passage. Cellular proteins were prepared and quantified as described later.

Measurement of DNA Synthesis Confluent HSCs in 24-well dishes were washed with phosphate-buffered saline (PBS) and incubated in serum-free medium for 48 hours. The cells were incubated with PPAR-␥ ligands for 15 minutes, and then with different mitogens for an additional 24 hours. DNA synthesis was measured as the incorporation of [3H]thymidine, as described previously.25

Cell Migration Assay Confluent HSCs were serum-starved for 48 hours and then exposed to PPAR-␥ ligands for 15 minutes. The cells were then washed, trypsinized, and resuspended in serum-free medium containing 1% albumin at a concentration of 3 ⫻ 105 cells/mL. Chemotaxis was measured in modified Boyden chambers equipped with 8-␮m-pore filters (Poretics, Livermore, CA) coated with rat tail collagen (Collaborative Biomedical Products, Bedford, MA) as previously described.26 At the end of the incubation, the filters were fixed, stained with Giemsa (Merck, Darmstadt, Germany), mounted, and viewed at 450⫻ magnification. Data are expressed as the average of cell counts obtained in 10 randomly chosen high-power fields.

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Analysis of MCP-1 Secretion Confluent HSCs in 24-well plates were deprived of serum for 24 hours. After replacement of the medium with fresh serum-free medium, the cells were exposed to PPAR-␥ agonists for 15 minutes and then to IL-1, TNF, or IFN-␥ for 24 hours. At the end of the incubation, the medium was collected and stored at ⫺20°C until assaying. MCP-1 secretion in the conditioned medium was measured using Western blot analysis as previously described.27 Briefly, 50 –100 ␮L of conditioned medium was dried, resuspended in Laemmli buffer,28 separated by 15% sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE), and electroblotted on a polyvinylidene-difluoride membrane. The membranes were blocked overnight at 4°C with 2% bovine serum albumin in 0.1% PBS–Tween, and then sequentially incubated at room temperature with an antiserum against baboon MCP-1 (1:1000) and with a horseradish peroxidase– conjugated secondary antibody. Detection was performed using chemiluminescence according to the manufacturer’s protocol (Amersham, Arlington Heights, IL). This technique has been shown to detect as little as 1 ng of human MCP-1 and has been previously validated using a commercially available enzyme-linked immunosorbent assay kit.27

Preparation of Cell Lysates Confluent, serum-starved HSCs were treated with the appropriate conditions, quickly placed on ice, and washed with ice-cold PBS. The monolayer was lysed in RIPA (radioimmunoprecipitation assay) buffer (20 mmol/L Tris-HCl [pH 7.4], 150 mmol/L NaCl, 5 mmol/L EDTA, 1% Nonidet P-40, 1 mmol/L Na3VO4, 1 mmol/L phenylmethylsulfonyl fluoride, and 0.05% [wt/vol] aprotinin). Insoluble proteins were discarded by high-speed centrifugation at 4°C. Protein concentration in the supernatant was measured in triplicate using a commercially available assay (Pierce, Rockford, IL).

ERK Assay ERK was immunoprecipitated from 25 ␮g of total cell lysate using polyclonal anti-ERK antibodies and protein A–Sepharose. After washing, the immunobeads were incubated in a buffer containing 10 mmol/L HEPES (pH 7.4), 10 mmol/L MgCl2, 0.5 mmol/L dithiothreitol, 0.5 mmol/L Na3VO4, 25 ␮mol/L adenosine triphosphatase, 1 ␮Ci [␥-32P]adenosine triphosphatase, and 0.4 mg/mL myelin basic protein for 30 minutes at 30°C. At the end of the incubation, the reaction was stopped by addition of Laemmli buffer and run on 15% SDS-PAGE. After electrophoresis, the gel was dried and autoradiographed.

Phosphatidylinositol 3-Kinase Assay This assay was performed after immunoprecipitation with antiphosphotyrosine antibodies, as described previously.26,29 Radioactive lipids were separated by thin-layer chromatography, using chloroform/methanol/30% ammonium hydroxide/ water (46/41/5/8). After drying, the plates were autoradio-

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graphed. The radioactive spots were then scraped and counted in a ␤-counter.

Western Blot Analysis of Cellular Proteins Equal amounts of total cellular proteins were separated by SDS-PAGE and analyzed by Western blot as described for the MCP-1 assay. Immunoblot analysis of platelet-derived growth factor (PDGF) receptor tyrosine phosphorylation was conducted after immunoprecipitation. Briefly, 150 ␮g of total cellular proteins was incubated with anti–PDGF-␤ receptor antibodies and protein A–Sepharose for 2 hours at 4°C. The immunobeads were washed twice in lysis buffer and once in 20 mmol/L Tris-HCl (pH 7.4) and 1 mmol/L Na3VO4, resuspended in Laemmli buffer, and analyzed by Western blot as described earlier.

Analysis of Cell Cycle The cell cycle was analyzed using a previously described technique with minor modifications.30 Subconfluent HSCs (60%–70% density) were serum-deprived and preincubated with 5 ␮mol/L 15d-PGJ2 or its vehicle for 15 minutes, and then exposed to 50 ng/mL PDGF for 20 hours. At the end of incubation, cells were harvested using PBS/EDTA and washed twice with PBS. Cells were suspended in a solution containing 50 ␮g/mL propidium iodide, 0.02% Nonidet P-40, and 0.5 mg/mL ribonuclease A in PBS. Samples were incubated in the dark at room temperature for 30 minutes and stored at 4°C until analysis. Cell fluorescence was measured by FACScan (BD, Franklin Lakes, NJ) and analyzed by the ModFit LT 2.0 software (Verity Software House, Topsham, ME) to determine the distribution of cells in the various phases of the cell cycle.

Preparation of Nuclear Extracts and Gel Mobility Shift Assay Nuclear extracts were prepared according to Andrew and Faller.31 Gel mobility shift analysis was performed using AP-1 or NF-␬B consensus oligonucleotides (Promega, Madison, WI), as described previously.32 Briefly, 5–10 ␮g of nuclear extracts was incubated for 30 minutes at room temperature in a buffer containing 35 mmol/L HEPES, pH 7.8, 0.5 mmol/L EDTA, 0.5 mmol/L dithiothreitol, 10% glycerol, 10 ␮g/mL polydI-dC, 0.28 mmol/L spermidine, and 50,000 – 100,000 cpm of 32P-labeled oligonucleotide probe. The DNAprotein complexes were separated by PAGE in 0.5⫻ Trisborate-EDTA. At the end of the run, the gel was dried and autoradiographed.

Northern Blot Analysis Isolation of total RNA and Northern blot analysis were performed using previously described methods.33 After transfer, the blots were sequentially hybridized with radiolabeled complementary DNA probes encoding for c-fos, c-myc, MCP-1, and the ribosomal protein 36B4 (control gene).

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Analysis of Transcriptional Activity Using Reporter Plasmids HSCs were seeded in 6-well dishes at a density of 300,000 cells/well and grown in complete medium for 24 hours. The cells were then washed and transfected with 2.5 ␮g of the reporter plasmid ACO-PPRE and 1.5 ␮g of a ␤-galactosidase expression vector using 4 ␮L of Lipofectamine (Life Technologies, Rockville, MD). The ACO-PPRE reporter plasmid contains 3 copies of PPAR response element from the promoter of rat acyl coenzyme A oxidase.23 After 5 hours, complete medium was added overnight and then replaced with fresh complete medium for 12 hours. The cells were then serum-starved overnight and incubated with 15d-PGJ2 or dimethyl sulfoxide for an additional 24 hours. At the end of the incubation, the cells were lysed using 1⫻ reporter lysis buffer (Promega) and the protein concentration was measured. Luciferase activity was measured in a buffer containing 25 mmol/L Gly-Gly buffer, 5 mmol/L adenosine triphosphatase, 15 mmol/L MgSO4, and 0.6 mmol/L coenzyme A. Light was measured for 10 seconds immediately after addition of luciferin (0.2 mmol/L). ␤-Galactosidase activity was measured using a commercially available kit (Promega).

Data Analysis Unless otherwise indicated, all data are representative of at least 3 experiments with similar results. Statistical analysis was performed by the Student t test or by 1-way analysis of variance, when appropriate. P values of 0.05 were considered significant.

Results PPAR-␥ Ligands Inhibit Proliferation and Migration of HSCs We first investigated whether PPAR-␥ agonists could modulate DNA synthesis of cultured human HSCs as an index of cell proliferation. As expected, incubation of serum-starved HSCs with PDGF induced a severalfold increase in the uptake of [3H]thymidine (Figure 1A). Preincubation of the cells with increasing concentrations of the PPAR-␥ agonists 15d-PGJ2 or troglitazone resulted in a marked and dose-dependent inhibition of DNA synthesis. At concentrations as high as 5 ␮mol/L 15d-PGJ2, or 10 ␮mol/L troglitazone, the stimulation of DNA synthesis by PDGF was virtually eliminated. To confirm the inhibitory effects of PPAR-␥ agonists on cell proliferation, we also tested the effects of 15d-PGJ2 and troglitazone on DNA synthesis in response to fetal bovine serum (Figure 1B). In this case also, the marked increase in cell proliferation was inhibited by PPAR-␥ ligands in a dose-dependent fashion, although the effects were less evident than in cells treated with PDGF. We

Figure 1. PPAR-␥ ligands inhibit proliferation of HSCs. Serum-starved HSCs were preincubated with increasing concentrations of 15d-PGJ2 or troglitazone for 15 minutes, before exposure to (A) 10 ng/mL PDGF or (B) 10% fetal bovine serum. (C) HSCs were preincubated with 5 ␮mol/L 15d-PGJ2 or its vehicle before exposure to 100 ng/mL epidermal growth factor (EGF) or 5 U/mL thrombin (THR). Fetal bovine serum (10%) was used as a positive control. After 24 hours, DNA synthesis was measured as the incorporation of [3H]thymidine. Mean ⫾ SD of a representative experiment.

also tested the effects of 15d-PGJ2 on the proliferative response induced by other mitogens, such as epidermal growth factor and thrombin.25,34 The stimulation of thymidine uptake was less pronounced than in response to fetal bovine serum, but exposure to the PPAR-␥ agonist blocked cell proliferation (Figure 1C). These data

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Effects of PPAR-␥ Ligands on PDGF’s Signaling, Proto-oncogene Activation, and Cell Cycle

Figure 2. PPAR-␥ ligands inhibit HSC chemotaxis. Serum-starved HSCs were preincubated with increasing concentrations of 15d-PGJ2 or troglitazone for 15 minutes. The cells were then trypsinized, and cell migration to 10 ng/mL PDGF was measured in modified Boyden chambers. Results are expressed as the mean number of cells (⫾SD) migrated to the underside of the filter in a representative experiment.

indicate that exposure of HSCs to PPAR-␥ ligands inhibits HSC proliferation independently of the mitogen used. Because chemotaxis represents an important characteristic of cells involved in the wound healing response in several tissues, we tested the effects of 15d-PGJ2 and troglitazone on PDGF-induced migration of HSCs. PDGF stimulated an 8-fold induction of HSC migration (Figure 2), as reported previously.32 Incubation with 15d-PGJ2 or troglitazone resulted in a marked inhibition of HSC chemotaxis. Remarkably, low concentrations of PPAR-␥ ligands resulted in ⬍75% reduction of cell migration, indicating that this parameter is very sensitive to the effects of these compounds.

We next investigated whether the observed effects of PPAR-␥ ligands could be mediated by changes in PDGF-dependent postreceptor signaling or proto-oncogene activation. Although PDGF activates several intracellular pathways,35 we focused on the signaling intermediates that have been shown to be necessary for PDGF-induced mitogenic or motogenic action in HSCs.27,32 Autophosphorylation on tyrosine residues immediately follows the binding of PDGF dimers to the cognate receptors. Exposure of serum-starved HSCs to PDGF resulted in the appearance of an evident increase in receptor tyrosine phosphorylation, as evaluated by immunoprecipitation followed by immunoblot analysis (Figure 3A). Preincubation with PPAR-␥ ligands, at concentrations that markedly reduce DNA synthesis or cell migration, did not inhibit PDGF-receptor phosphorylation. Activation of phosphatidylinositol 3-kinase (PI 3-K) is mediated by the recruitment of this enzyme by the activated PDGF receptor,36 and inhibitors of PI 3-K activity completely block PDGF-induced proliferation and chemotaxis.27 However, the increase in PI 3-K activity induced by PDGF (Figure 3B) was unchanged in cells treated with troglitazone or 15d-PGJ2, indicating that the inhibition of cell proliferation and chemotaxis caused by PPAR-␥ ligands is independent of any interference with PI 3-K activation. We reported recently that in PDGF-stimulated HSCs, activation of ERK is necessary for mitogenesis and contributes to chemotaxis.32 In addition, troglitazone has

Figure 3. Effects of PPAR-␥ ligands on PDGF-induced tyrosine phosphorylation and PI 3-K activation. (A) Serum-starved HSCs were preincubated with 3 ␮mol/L troglitazone or with different concentrations of 15d-PGJ2 before addition of 10 ng/mL PDGF for 10 minutes. Total cell lysates were immunoprecipitated with anti–PDGF-␤ receptor antibodies, separated by SDS-PAGE, and blotted with antiphosphotyrosine antibodies. Migration of the molecular-weight marker is shown on the left. (B) Serum-starved HSCs were preincubated with different concentrations of troglitazone or 15d-PGJ2 before addition of 10 ng/mL PDGF for 10 minutes. Analysis of PI 3-K activity was performed on antiphosphotyrosine immunoprecipitates as described in Materials and Methods. Migration of 3-OH–phosphorylated phosphatidylinositol is shown on the left.

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Figure 4. Effects of PPAR-␥ ligands on ERK phosphorylation and activation. (A) Serum-starved HSCs were preincubated with 3 ␮mol/L troglitazone or 5 ␮mol/L 15d-PGJ2 before addition of 10 ng/mL PDGF for 10 minutes. Total cell lysate (40 ␮g) was separated by SDS-PAGE and sequentially immunoblotted with antibodies specifically recognizing the phosphorylated form of ERK (top) or directed against ERK (bottom). Migration of the molecular-weight marker is shown on the left. (B) Serum-starved HSCs were preincubated with 10 ␮mol/L troglitazone or 5 ␮mol/L 15d-PGJ2 before addition of 10 ng/mL PDGF for 10 minutes. Total cell lysate (25 ␮g) was immunoprecipitated with anti-ERK antibodies, and ERK activity was measured as the ability to phosphorylate myelin basic protein as described in Materials and Methods.

been shown to interfere with the ERK pathway in vascular smooth muscle cells.37,38 To assess whether inhibition of ERK could contribute to the effects of PPAR-␥ ligands on HSCs, we analyzed the activation of ERK in cells exposed to PDGF in the presence or absence of troglitazone or 15d-PGJ2. ERK activation is associated with phosphorylation of specific threonine and tyrosine residues by the upstream kinase MEK. We found that neither of the 2 PPAR-␥ ligands affected ERK activation when we used antibodies specifically recognizing the phosphorylated form of ERK (Figure 4A). To confirm and extend this observation, we immunoprecipitated ERK from total cell lysates and measured ERK activity in an immune complex kinase assay using myelin basic protein as a substrate (Figure 4B). Also in this case, the evident increase induced by incubation with PDGF was

unaffected by pretreatment with the 2 PPAR-␥ ligands. Together, these experiments show that agonists of PPAR-␥ modulate PDGF action without interfering with postreceptor signaling pathways known to be involved in mitogenesis or chemotaxis. Activation of proto-oncogenes such as c-fos or c-myc has been shown to be implicated in the mitogenic signaling of several growth factors including PDGF.39,40 Expression of c-fos is mediated, at least in part, by molecules located downstream of ERK and has been shown to be inhibited by troglitazone in smooth muscle cells. In HSCs, PDGF markedly increased the expression of c-fos, but neither 15d-PGJ2 or troglitazone affected the steady-state messenger RNA (mRNA) levels for this proto-oncogene (Figure 5A). Similarly, exposure of HSCs to PDGF increased c-myc expression, but PPAR-␥ li-

Figure 5. Effects of PPAR-␥ ligands on proto-oncogene expression in HSCs. (A) Serum-starved HSCs were preincubated with 3 ␮mol/L troglitazone or 5 ␮mol/L 15d-PGJ2 before addition of 10 ng/mL PDGF for the indicated time points. Total RNA (15 ␮g) was sequentially analyzed by Northern blotting with probes encoding for c-fos and the ribosomal protein 36B4 (control gene). (B) Serum-starved HSCs were preincubated with 3 ␮mol/L troglitazone before addition of 10 ng/mL PDGF for 60 minutes. Total RNA (15 ␮g) was sequentially analyzed by Northern blotting with probes encoding for c-myc and the ribosomal protein 36B4.

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PPAR-␥ Ligands Inhibit Expression and Secretion of MCP-1

Figure 6. Effects of PPAR-␥ ligands on cell cycle. Serum-starved HSCs were preincubated with 5 ␮mol/L 15d-PGJ2 or its vehicle and either left untreated or incubated with 50 ng/mL PDGF for 20 hours as indicated. At the end of the incubation, the cells were detached and cell cycle analysis was performed as described in Materials and Methods. Data show the percentage of cells in each phase of the cell cycle in a representative experiment.

gands did not inhibit this effect (Figure 5B, and data not shown). These results show that PPAR-␥ acts downstream of early nuclear signaling and proto-oncogene activation mediated by PDGF. To better define the level of action of PPAR-␥ agonists on PDGF-induced mitogenesis, we analyzed the cell cycle in HSCs in the presence or absence of 15d-PGJ2 (Figure 6). As expected, exposure to PDGF was associated with a marked decrease in the percentage of cells in the G0/G1 phase, together with an increase in the number of cells in the S phase. Addition of the PPAR-␥ agonist before PDGF reduced the number of cells in the S phase, in agreement with the effects on [3H]thymidine uptake, and the percentage of cells in the G0/G1 phase was similar to the percentage observed in untreated cells. Thus, addition of 15d-PGJ2 inhibits PDGF-induced progression of the cell cycle beyond the G1 phase.

Activated HSCs acquire the ability to modulate the recruitment and activation of inflammatory cells. The proinflammatory role of HSCs is well exemplified by the expression of MCP-1, a potent chemoattractant for monocytes and T lymphocytes, which is secreted at high levels by HSCs in their myofibroblast-like phenotype.3 Secretion of MCP-1 in HSC-conditioned medium was detectable by immunoblotting and appeared as 2 strong bands and a weaker band because of differences in glycosylation.3 MCP-1 secretion is stimulated by several proinflammatory cytokines, including IL-1, TNF-␣, and IFN-␥ (Figure 7).3 Exposure of HSCs to PPAR-␥ ligands before the addition of the cytokines resulted in a dosedependent reduction of the amount of immunoreactive MCP-1 detectable in the conditioned medium (Figure 7A, B, and C). Whereas 0.5–1 ␮mol/L PGJ2 generally resulted in an inhibition of approximately 50%, as evaluated by densitometric analysis, concentrations of 5 ␮mol/L 15d-PGJ2 or 10 ␮mol/L troglitazone virtually abolished the cytokine-mediated induction of MCP-1 secretion. We have previously shown that the up-regulation of MCP-1 secretion in response to proinflammatory cytokines in HSCs is accompanied by an increase in MCP-1 gene expression. Accordingly, IL-1, TNF, and IFN-␥ increased the abundance of MCP-1 mRNA (Figure 8A and 8B). The expression levels of MCP-1 in response to all 3 agonists were reduced by 15d-PGJ2 (Figure 8A). Troglitazone had similar effects, although the inhibition of MCP-1 mRNA expression was somehow less impressive (Figure 8B, and data not shown).

Figure 7. PPAR-␥ ligands inhibit cytokine-induced secretion of MCP-1. Serum-starved HSCs were preincubated with the indicated concentrations of 15dPGJ2 or with troglitazone (B, 10 ␮mol/L; C, 3 ␮mol/L) before addition of IL-1 (A, 4 ng/mL), TNF-␣ (B, 100 ng/mL), or IFN-␥ (C, 1000 U/mL) for 24 hours. Aliquots of the conditioned medium were separated by SDSPAGE and immunoblotted with anti–MCP-1 antibodies. Migration of the molecular-weight marker is shown on the left.

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Figure 8. Inhibition of MCP-1 gene expression by PPAR-␥ ligands. (A) Serum-starved HSCs were preincubated with 5 ␮mol/L 15d-PGJ2 for 15 minutes before addition of IL-1 (4 ng/mL), TNF-␣ (100 ng/ mL), or IFN-␥ (1000 U/mL) for 4 hours. (B) Serumstarved HSCs were preincubated with 10 ␮mol/L troglitazone for 15 minutes before addition of 100 ng/mL TNF-␣ for 4 hours. Total RNA (15 ␮g) was sequentially analyzed by Northern blotting with probes encoding for MCP-1 and for the ribosomal protein 36B4.

Inhibition of MCP-1 Occurs Independently of Changes in the Activity of p38MAPK or in the Activation of NF-␬B or AP-1 The intracellular signaling pathways leading from receptor activation to MCP-1 expression are only partially known. Activation of p38MAPK, a member of the stress-activated protein kinase family, has been shown to be necessary for induction of many inflammation-related genes, including MCP-1.41 As described earlier for ERK, activation of p38MAPK requires phosphorylation on threonine and tyrosine residues by a dual-specificity kinase, and the activation of the molecule may be detected using phosphospecific antibodies. Exposure of HSCs to TNF-␣ resulted in increased phosphorylation of p38MAPK (Figure 9). However, neither 15d-PGJ2 or troglitazone, at concentrations that markedly reduce MCP-1 expression and secretion, modified the activation of this kinase. The inhibition of monocyte activation by PPAR-␥ ligands has been shown to be associated with reduced transcriptional activity of NF-␬B or AP-1– driven pro-

Figure 9. Effects of PPAR-␥ ligands on phosphorylation of p38MAPK. Serum-starved HSCs were preincubated with 3 ␮mol/L troglitazone or 5 ␮mol/L 15d-PGJ2 before addition of 100 ng/mL TNF-␣ for 15 minutes. Total cell lysate (40 ␮g) was separated by SDS-PAGE and sequentially immunoblotted with antibodies specifically recognizing the phosphorylated form of p38MAPK (top) or directed against p38MAPK (bottom).

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Table 1. Evaluation of the Toxic Effects of PPAR-␥ Agonists on Cultured HSCs Living cells (% of total) DMSO 15d-PGJ2 Troglitazone

96.7 ⫾ 1.7 95.1 ⫾ 1.4 98.4 ⫾ 1.6

NOTE. Subconfluent HSCs were incubated with 0.1% dimethylsulfoxide (vehicle), 5 ␮mol/L 15d-PGJ2, or 10 ␮mol/L troglitazone for 24 hours. At the end of incubation, floating cells were collected and pooled with adherent cells after trypsinization. The number of alive cells was measured by trypan blue exclusion; results are expressed as mean ⫾ SEM (n ⫽ 3). No significant differences among the 3 groups were observed by 1-way ANOVA.

Figure 10. Effects of PPAR-␥ ligands on activation of NF-␬B and AP-1. Serum-starved HSCs were preincubated with 5 ␮mol/L 15d-PGJ2 before addition of 100 ng/mL TNF-␣ for 45 minutes. Total nuclear extract (5 ␮g) was used in gel mobility shift assays using consensus oligonucleotides for (A) NF-␬B or (B) AP-1. Migration of the shifted complex is indicated by arrows.

moters.22 Because NF-␬B and AP-1 are also important for MCP-1 transcription, we analyzed the effects of PPAR-␥ agonists on the activation of these factors. TNF increased the DNA-binding activity present in HSC nuclear extracts to consensus oligonucleotides recognizing NF-␬B (Figure 10A) or AP-1 (Figure 10B), as shown by electrophoretic mobility shift assays. However, preincubation of HSCs with 5 ␮mol/L 15d-PGJ2 did not affect the activation of these transcriptional regulators, indicating that the inhibitory effect of PPAR-␥ ligands on MCP-1 expression is independent of these pathways.

a luciferase reporter plasmid under the control of PPAR-␥ (ACO-PPRE),23 exposure to 15d-PGJ2 induced a 2-fold, statistically significant increase in the activity of the reporter gene (Figure 11), indicating that PPAR-␥ agonists are transcriptionally active in HSCs. Reduced Expression of PPAR-␥ in Activated HSCs The levels of PPAR-␥ are markedly variable in different tissues and cultured cells. We evaluated whether the phenotypic changes observed in the process of HSC activation are accompanied by modifications in the levels of PPAR-␥. We compared cell lysates obtained from HSCs immediately after isolation from normal liver tissue, and, therefore, showing a quiescent phenotype, with those from cells cultured for 3 days or after 2 passages in culture, when transition to a myofibroblastlike phenotype is complete (Figure 12). In freshly isolated HSCs a clearly detectable band of an apparent molecular weight corresponding to that of PPAR-␥ was

The Effects of PPAR-␥ Ligands on Cultured HSCs Are Not Caused by Cell Toxicity To rule out that the actions of PPAR-␥ ligands on HSCs could be mediated by cell toxicity, we evaluated the effects of these compounds on cell viability. As shown in Table 1, no differences in cell viability were observed using the highest concentration of PPAR-␥ ligands and the longest time point (24 hours) used in the present study. In addition, when PPAR-␥ agonists were added to HSC cultures for 45 minutes, and subsequently withdrawn, the inhibitory effects on MCP-1 secretion were partially reverted (data not shown), further confirming the lack of toxic effects in this system. We also investigated whether PPAR-␥ exerts transcriptional effects in cultured human HSCs. In cells transfected with

Figure 11. 15d-PGJ2 activates PPAR-␥–induced transcription in HSCs. Subconfluent HSCs were transfected with a reporter plasmid under the control of PPAR-␥ and an expression vector for ␤-galactosidase, as described in Materials and Methods. The cells were then incubated with 15d-PGJ2 or its vehicle for 24 hours, and the levels of luciferase and ␤-galactosidase activity were determined. The data represent luciferase activity (arbitrary units) normalized for ␤-galactosidase activity and protein concentration. Mean ⫾ SEM of 3 experiments. *P ⬍ 0.05.

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Figure 12. Reduced expression of PPAR-␥ in activated HSCs. Total cell lysate (40 ␮g) from freshly isolated human HSCs (lane 1), from HSCs cultured on plastic for 3 days (lane 2), or after subculture (lane 3) was analyzed by immunoblotting using an antibody directed against PPAR-␥ (top panel). The membrane was stripped and reblotted with an antibody against ␣–smooth muscle actin (middle panel). The lower panel shows Ponceau red staining of the membrane indicating comparable protein loading. Migration of the molecular-weight marker is indicated on the left.

observed. Culture of HSCs for 3 days led to a marked reduction (⬃60% by densitometric analysis) of PPAR-␥ expression, which became barely detectable after cell subculture, a condition associated with a fully activated phenotype, as shown by expression of ␣–smooth muscle actin (Figure 12).

Discussion Understanding the molecular mechanisms that regulate the biology of HSCs has a major relevance for the pathophysiology of liver fibrosis.2,42 The data of the present study indicate that activation of PPAR-␥, a member of the nuclear hormone–receptor superfamily, modulates different biological actions of HSCs that contribute to the process of liver inflammation and fibrogenesis. Exposure of HSCs to 2 well-established ligands of PPAR-␥, the endogenously produced prostanoid 15dPGJ2 and troglitazone, resulted in complete inhibition of HSC proliferation, migration, and expression of the chemokine MCP-1, 3 biological actions associated with the activated phenotype. We also found a striking decrease in the expression of PPAR-␥ in activated vs. quiescent HSCs, as established using freshly isolated and cultureactivated human HSCs. Interestingly, PPAR-␥ levels

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were reduced as early as after 3 days in culture, when the expression of ␣–smooth muscle actin is not yet evident. The decrease in PPAR-␥ at the early stages of HSC activation and the fact that PPAR-␥ ligands inhibit at least some of the characteristics associated with the activated phenotype of HSCs suggest that PPAR-␥ may be involved in the maintenance of a quiescent phenotype. Along these lines, the ability of PPAR-␥ to regulate differentiation and phenotypic modulation has been shown in several systems, as in the cases of the transition of preadipocytes to adipocytes,19 differentiation of monocytes to macrophages and foam cells,43 or reversal of the transformed phenotype in cancer cells.20,21 The molecular mechanisms responsible for the effects of PPAR-␥ agonists are still controversial. In rat smooth muscle cells, troglitazone interfered with the Ras/ERK pathway at the level of ERK activation or of c-fos expression, depending on the agonist used.37,38 Therefore, inhibition of the ERK cascade has been suggested to mediate the inhibition of proliferation and migration in these cells.37,38 In contrast, the results of this study show that neither troglitazone nor 15d-PGJ2 inhibits ERK phosphorylation or activation at concentrations that completely block proliferation and migration of HSCs. In addition, the PDGF-induced increase in c-fos expression, which is at least partly dependent on ERK activation, was unchanged in HSCs exposed to PPAR-␥ agonists. We also explored other pathways that have been shown to contribute to transduce PDGF mitogenic or motogenic signals. However, neither activation of PI 3-K nor expression of c-myc was affected by PPAR-␥ ligands. These findings indicate that in HSCs the effects of PPAR-␥ activators on cell proliferation and migration occur downstream of the activation of postreceptor-signaling pathways and of proto-oncogene expression. This hypothesis is supported by the observation that the inhibition of HSC proliferation was observed irrespective of the agonist used because PPAR-␥ agonists blocked mitogenesis also in response to epidermal growth factor and thrombin. In addition, analysis of the cell cycle in HSCs exposed to PPAR-␥ agonists showed the inability to proceed beyond the G1 phase when stimulated with PDGF. In adipocytes, where PPAR-␥ agonists induce terminal differentiation and withdrawal from the cell cycle, PPAR-␥ bypasses the requirement for Rb hyperphosphorylation,44 and the growth-inhibitory effect of PPAR-␥ on fibroblasts is mediated by the inhibition of E2F/DP DNA-binding activity.45 These data in other systems are in keeping with an action downstream of ERK activation or c-fos expression and with a block in the G1 phase of the cell cycle.

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The ability of 15d-PGJ2 and troglitazone to block the production of MCP-1 in HSCs indicates that PPAR-␥ may interfere with the expression of members of the chemokine family. We focused our attention on MCP-1, as a paradigm chemokine, in consideration of its emerging importance in the regulation of leukocyte trafficking within the liver. MCP-1 expression is directly related to the number of monocytes infiltrating the portal tract during chronic hepatitis,46 and its expression precedes leukocyte recruitment in a rodent model of liver damage.47 Expression of MCP-1, and of other chemokines, is markedly increased in activated HSCs and may be induced by several soluble mediators, including proinflammatory cytokines.3 Exposure of HSCs to PPAR-␥ ligands resulted in complete inhibition of MCP-1 gene and protein expression, indicating that this class of compounds modulates a key mechanism leading to hepatic inflammation. Interestingly, recent data indicate that monocyte activation may be inhibited by PPAR-␥ ligands.22,23 The ability of these compounds to modulate monocyte gene expression is associated with inhibition of transcription driven by NF-␬B and AP-1 in reporter assays. Because NF-␬B and AP-1 play an important role in the regulation of transcription of the MCP-1 gene,48,49 we investigated whether activation of these transcriptional regulators was affected by PPAR-␥ ligands. However, the DNA-binding activity of both NF-␬B and AP-1 was not changed in cells exposed to effective concentrations of a PPAR-␥ agonist. This is in agreement with data by Peraldi et al.,50 who showed that PPAR-␥ agonists block insulin resistance induced by TNF-␣ but do not affect the ability of this cytokine to activate NF-␬B. PPAR-␥ may modulate transcription at a level downstream of the interaction of transcription factors with their responsive elements. Alternatively, the inhibitory effect on chemokine expression may be the result of the effect on specific regulatory sequences in the MCP-1 gene. This latter hypothesis is indirectly supported by the observation that PPAR-␥ inhibits MCP-1 expression elicited by 3 different agonists: IL-1, TNF, and IFN-␥. Although IL-1 and TNF are believed to activate MCP-1 transcription through partially overlapping pathways, it has been recently shown that IFN-␥–induced transcription of MCP-1 occurs through specific regulatory elements in the 5⬘-flanking region.51 These data suggest that the action of PPAR-␥ is mediated by a specific effect on the MCP-1 gene independent of the activating cytokine used. An intriguing aspect of the present results is related to the interaction between PPAR-␥ and the retinoids. The retinoids activate RXR and the retinoid acid receptor,

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which are members of the same family of nuclear hormone receptors that include PPAR-␥, and PPAR-␥ requires RXR to form a transcriptionally active heterodimer.11 The HSCs play an important role in the storage and metabolism of retinoids, and it has been extensively shown that acquisition of the activated phenotype is associated with loss of retinoid content.7,8 Treatment of HSCs with retinoids inhibits several biological actions that accompany the activated phenotype, such as cell proliferation or expression of extracellular matrix.52 A recent study showed that exposure of HSC to 9-cis retinoic acid, a metabolite that selectively activates RXR, results in the reduction of cell proliferation.53 Moreover, the mRNA levels for RXR-␣ were decreased in HSC isolated from bile duct–ligated rats.54 These findings are particularly interesting because ligands of RXR have been shown to exert biological actions similar to those elicited by PPAR-␥ agonists, and the simultaneous addition of the 2 types of ligands is associated with a synergistic effect.55 These data, together with our results, suggest that vitamin A metabolites and PPAR-␥ ligands may cooperate in maintaining a quiescent HSC phenotype in the normal liver. The identification of PPAR-␥ as a novel modulator of HSC biology raises a number of questions on its possible regulatory mechanisms. In quiescent HSCs, where high expression is observed, the molecules responsible for PPAR-␥ activation need to be identified. Besides PG of the J series, other endogenous activators of PPAR-␥ are being discovered, including oxidative metabolites of linoleic acid.12 It will be important to establish if these or other compounds are synthesized in quiescent cells and whether their abundance decreases along with the activation process. Recently, rat liver myofibroblasts have been characterized as another matrix-producing cell partially distinct from activated HSCs.56 Although these cells have not yet been characterized in the human liver, it will be relevant to understand whether they exhibit different levels of PPAR-␥ or are differentially modulated by agonists of this transcription factor. An important consequence of our present data is the potential application of PPAR-␥ agonists in the treatment of liver fibrosis. Most drugs capable of blocking the biological actions of HSCs in vitro may not be used in vivo unless a delivery system is developed to selectively target these cells. On the other hand, thiazolidinediones have been approved for use in type II diabetes and are generally well tolerated. The in vitro data obtained in this study and the relative safety of thiazolidinediones call for in vivo studies aimed at establishing whether interference with

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the PPAR-␥ pathway is beneficial for the treatment of liver fibrosis.

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Received August 26, 1999. Accepted March 3, 2000. Address requests for reprints to: Fabio Marra, M.D., Ph.D., Dipartimento di Medicina Interna, Viale Morgagni, 85, I-50134 Florence, Italy. e-mail: [email protected]; fax: (39) 055-417-123. Supported by a MURST grant (project: molecular and cellular biology of hepatic fibrosis) and by the Italian Liver Foundation. The authors thank Wanda Delogu and Chiara Sali for skillful technical help and Drs. T. Hashimoto, A. J. Valente, and B. Seed for providing some of the reagents used in this study.