Cytosolic phospholipase A2α and peroxisome proliferator-activated receptor γ signaling pathway counteracts transforming growth factor β–mediated inhibition of primary and transformed hepatocyte growth

Authors

  • Chang Han,

    Corresponding author
    1. Department of Pathology and Laboratory Medicine, Tulane University School of Medicine, New Orleans, LA
    2. Department of Pathology, University of Pittsburgh School of Medicine, Pittsburgh, PA
    • Department of Pathology and Laboratory Medicine, Tulane University School of Medicine, 1430 Tulane Avenue, SL-79, New Orleans, LA 70112
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  • William C. Bowen,

    1. Department of Pathology, University of Pittsburgh School of Medicine, Pittsburgh, PA
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  • Guiying Li,

    1. Department of Pathology, University of Pittsburgh School of Medicine, Pittsburgh, PA
    2. Key Laboratory for Molecular Enzymology and Engineering of Ministry of Education, Jilin University, Changchun, China
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  • Anthony J. Demetris,

    1. Department of Pathology, University of Pittsburgh School of Medicine, Pittsburgh, PA
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  • George K. Michalopoulos,

    1. Department of Pathology, University of Pittsburgh School of Medicine, Pittsburgh, PA
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  • Tong Wu

    Corresponding author
    1. Department of Pathology and Laboratory Medicine, Tulane University School of Medicine, New Orleans, LA
    2. Department of Pathology, University of Pittsburgh School of Medicine, Pittsburgh, PA
    • Department of Pathology and Laboratory Medicine, Tulane University School of Medicine, 1430 Tulane Avenue, SL-79, New Orleans, LA 70112
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    • fax: 504-988-7862


  • Potential conflict of interest: Dr. Demetris is a consultant for Wyeth, Novartis, and Bristol-Myers Squibb.

Abstract

Hepatocellular carcinoma often develops in the setting of abnormal hepatocyte growth associated with chronic hepatitis and liver cirrhosis. Transforming growth factor β (TGF-β) is a multifunctional cytokine pivotal in the regulation of hepatic cell growth, differentiation, migration, extracellular matrix production, stem cell homeostasis, and hepatocarcinogenesis. However, the mechanisms by which TGF-β influences hepatic cell functions remain incompletely defined. We report herein that TGF-β regulates the growth of primary and transformed hepatocytes through concurrent activation of Smad and phosphorylation of cytosolic phospholipase A2α (cPLA2α), a rate-limiting key enzyme that releases arachidonic acid for the production of bioactive eicosanoids. The interplays between TGF-β and cPLA2α signaling pathways were examined in rat primary hepatocytes, human hepatocellular carcinoma cells, and hepatocytes isolated from newly developed cPLA2α transgenic mice. Conclusion: Our data show that cPLA2α activates peroxisome proliferator-activated receptor γ (PPAR-γ) and thus counteracts Smad2/3-mediated inhibition of cell growth. Therefore, regulation of TGF-β signaling by cPLA2α and PPAR-γ may represent an important mechanism for control of hepatic cell growth and hepatocarcinogenesis. (HEPATOLOGY 2010;)

Transforming growth factor β (TGF-β) is a multifunctional cytokine that plays an important role in the regulation of cell proliferation, differentiation, migration, apoptosis, extracellular matrix production, angiogenesis, and neoplasia.1-4 The actions initiated by TGF-β are complex and often vary depending on individual cell or tissue types and the activation status of other intracellular signaling pathways. In the liver, TGF-β is well known to regulate stellate cell activation/liver fibrosis, hepatocyte proliferation/apoptosis, and hepatocarcinogenesis.

There are three mammalian TGF-β isoforms, TGF-β1, TGF-β2, and TGF-β3, all of which signal through a heteromeric complex of type I and type II TGF-β receptors.1, 5 Binding of ligand to the type II receptor results in the recruitment and activation of one of two type I receptors. The activated type I receptor phosphorylates Smad2 and Smad3. The phosphorylated Smad2 and Smad3 then associate with Smad4 and translocate to the nucleus. The mitoinhibition by TGF-β is predominantly mediated through activation of the Smad pathway. In addition, the activated TGF-βRII and TGF-βRI receptor complex can also signal independently of Smads, via extracellular signal-regulated kinase (ERK), c-Jun NH2-terminal kinase, p38 mitogen-activated protein kinase (MAPK), phosphatidylinositol-3 kinase, and Rho GTPases.1, 2, 6 Recent studies have revealed an intriguing link between TGF-β and interleukin-6/signal transducer and activator of transcription 3 signaling pathways in hepatocarcinogenesis.7, 8 However, it remains unclear whether TGF-β may also modulate hepatocarcinogenesis through interaction with other growth-regulating signaling pathways. In this study, we presented evidence that TGF-β regulates the growth of primary and transformed hepatocytes through concurrent activation of Smad-mediated gene transcription and phosphorylation of cytosolic phospholipase A2α (cPLA2α).

Prostaglandin (PG) signaling is implicated in the growth control of various human cells and cancers.9-14 PG biosynthesis is tightly controlled by a series of enzymes including the group IVa cytosolic phospholipase A2 (cPLA2α) that selectively cleaves arachidonic acid (AA) from membrane phospholipids, and cyclooxygenase-2 (COX-2) that converts AA substrate to PGs.9-14 This signaling cascade is active in various human cancers including hepatocellular carcinoma and promotes tumor growth by enhancing tumor cell proliferation, survival, invasion, or angiogenesis.14-16 Whereas PGs regulate cell functions through activation of specific G protein–coupled receptors on plasma membrane, studies from our laboratory have shown that the cPLA2α-derived arachidonic acid can also regulate cellular functions through activation of nuclear receptors including peroxisome proliferator-activated receptor γ (PPAR-γ).17, 18

This study describes the interaction between TGF-β and prostaglandin signaling pathways in primary and transformed hepatocytes. Our data show that TGF-β phosphorylates and activates cPLA2α and the cPLA2α-derived AA subsequently activates PPAR-γ, leading to inhibition of Smad2/3. This phenomenon is further verified in hepatocytes isolated from newly developed transgenic mice with targeted overexpression of cPLA2α in the liver. Our findings suggest that the level and activation status of cPLA2α/PPAR-γ in hepatic cells may represent a key factor that determines the cellular response to TGF-β and modulates hepatocarcinogenesis.

Abbreviations:

AA, arachidonic acid; cDNA, complementary DNA; COX-2, cyclooxygenase-2; cPLA2α, cytosolic phospholipase A2α; ERK, extracellular signal-regulated kinase; MAPK, mitogen-activated protein kinase; PG, prostaglandin; PGE2, prostaglandin E2; PPAR-γ, peroxisome proliferator-activated receptor γ; PPRE, peroxisome proliferator response element; SD, standard deviation; siRNA, small interfering RNA; TGF-β, transforming growth factor β; WT, wild-type.

Materials and Methods

Materials.

Minimum essential medium with Earle's salts, fetal bovine serum, glutamine, antibiotics, Lipofectamine PLUS reagent, and Lipofectamine 2000 reagent were purchased from Invitrogen (Carlsbad, CA). Human TGF-β1 was purchased from R&D Systems, Inc. (Minneapolis, MN). AA and prostaglandin E2 (PGE2) were purchased from Calbiochem (San Diego, CA). The PPAR-γ agonists, ciglitazone and piglitazone, were purchased from Cayman Chemical (Ann Arbor, MI). The cPLA2α specific inhibitor pyrrolidine derivative, the COX2 inhibitor NS398, the p38 MAPK inhibitor SB203580 and the p42/44 MARK inhibitor PD98059 were purchased from Calbiochem (San Diego, CA). The antibodies against human cPLA2α, PPAR-γ, TGF-βRI (transforming growth factor β receptor I), TGF-βRII (transforming growth factor β receptor II), PAI-1 (the type 1 plasminogen activator inhibitor) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Antibodies against phospho-cPLA2α (Ser505), phospho-p38 MARP (Thr180/Tyr182), phospho-p42/44 MARP (Thr202/Tyr204), phospho-Smad2 (Ser465/467), phospho-Smad3 (Ser423/425), p38 MARP, p42/44 MARP, Smad2, Smad3, and Smad4 were purchased from Cell Signaling (Beverly, MA). Antibody against β-actin was purchased from Sigma (St. Louis, MO). Antibody against glyceraldehyde 3-phosphate dehydrogenase was purchased from Ambion (Austin, TX). Amersham ECL Plus western blotting detection reagents were purchased from GE Healthcare (Piscataway, NJ). The cell proliferation assay reagent WST-1 was purchased from Roche Molecular Biochemicals (Indianapolis, IN). The PPAR-γ expression plasmid and PPRE-Luc reporter vector were purchased from Addgene (Cambridge, MA). The small interfering RNAs (siRNAs) for cPLA2α, PPAR-γ, Smad2, or Smad3 were purchased from Dharmacon (Chicago, IL).

Isolation of Primary Hepatocytes.

Primary hepatocytes were isolated from the cPLA2α transgenic mice, matched wild-type (WT) mice, and Fisher 344 rats (Harlan, Indianapolis, IN) using the modified two-step collagenase perfusion technique as described.19, 20 Freshly isolated hepatocytes of >90% viability, as assessed by trypan blue exclusion, were placed on rat-tail collagen I–coated culture plates at a density of either 3 to 4 × 106 cells/100-mm plastic dish for western blotting or 1 × 105 cells/each well of six-well plates for the measurement of DNA synthesis in Williams' E medium supplemented with 5% calf serum. The cells were incubated at 37°C (5% CO2) and checked for adherence of monolayers after 2-4 hours. Once adhered, the cells were changed to serum-free Williams' E medium for 2 hours, then subjected to the treatment as indicated in the text.

Hepatocyte DNA Synthesis.

The primary hepatocyte cultures were treated with either different concentrations of TGF-β1 or PGE2 as indicated. To determine in vitro DNA synthesis, 1 μCi [3H]thymidine (PerkinElmer, Boston, MA) was added to each well of six-well plates. After overnight incubation, the hepatocytes were harvested and [3H]-thymidine incorporation was measured using a scintillation counter.

Cell Growth Assay.

Cell growth was determined using the cell proliferation reagent WST-1, a tetrazolium salt that is cleaved by mitochondrial dehydrogenases in viable cells. Briefly, 100 μL of cell suspension (containing 0.5 to 2 × 104 cells) were plated in each well of 96-well plates. After overnight culture to allow reattachment, the cells then were treated with specific reagents such as different concentrations of TGF-β1 in serum-free medium for indicated time points. At the end of each treatment, the cell proliferation reagent WST-1 (10 μL) was added to each well, and the cells were incubated at 37°C for 0.5-5 hours. A450 nm was measured using an automatic enzyme-linked immunosorbent assay plate reader.

Cell Culture and Transient Transfection.

Three different human hepatocellular carcinoma cell lines (Hep3B, HepG2, and Huh7) were cultured according to our described methods.18, 21 For transient transfection assays, the cells with 80% confluence were transfected with the cPLA2α expression plasmid (with MT-2 as control plasmid) or the PPAR-γ expression plasmid (with pcDNA as control plasmid) using Lipofectamine PLUS reagent. The cells with optimal overexpression of either cPLA2α or PPAR-γ were confirmed by immunoblotting and subsequently used for further experiments.

Luciferase Reporter Assay.

The cells with 80% confluence were transiently transfected with either p3TP or PPRE reporter vector using Lipofectamine PLUS reagent. After transfection, the cells were treated with specific reagent such as PPAR-γ agonists ciglitazone and piglitazone in serum-free medium for 24 hours. The cell lysates were then obtained with 1× reporter lysis buffer (Promega). The luciferase activity was assayed in a Berthold AutoLumat LB 953 luminometer (Nashua, NH) using the luciferase assay system from Promega. The relative luciferase activity was calculated after normalization of cellular proteins. All values are expressed as fold induction relative to basal activity.

Phosphorylation of cPLA2α.

Analysis for cPLA2α phosphorylation was performed as described.22 Equal amounts of cell lysate were preincubated with 5 μg/mL mouse anti-human cPLA2α monoclonal antibody for 1 hour followed by addition of 20 μL of protein A/G-agarose (Santa Cruz Biotechnology) for overnight at 4 °C. The cell lysate preincubated with mouse immunoglobulin G was used as the negative control. After three washes with the same hypotonic buffer, the pellet was used for immunoblotting using rabbit anti–phospho-cPLA2α (Ser505) antibody.

DNA–Protein Binding.

DNA–protein binding was performed by the biotinylated oligonucleotides precipitation assay as described,23 with minor modifications. Briefly, 1 μg 5′-biotinylated, double-stranded oligonucleotides that corresponded to the Smad3 binding site from the PAI-1 promoter region (forward, 5′- CAACCTCAGCCAGACAAGGTTGTTGACACA AGAGAGCCCTCAGGGGCACAGAGAGAGTCTGG ACACGTGGGGAGTCAGCCGTGTATCATCGGAG GCGGCCGGGC-3′; reverse, 5′- GCCCGGCCGCC TCCGATGATACACGGCTGACTCCCCACGTGTC CAGACTCTCTCTGTGCCCCTGAGGGCTCTCT TGTGTCAACAACCTTGTCTGGCTGAGGTTG-3′; synthesized by Sigma-Genosys in Woodland, Texas) were mixed with equal amount of the cell extract and 10 μg poly(dl-dC)/poly(dl-dC) overnight at 4°C. After precipitation with ImmunoPure streptavidin-agarose beads (Pierce, Rockford, IL) at 4°C for another 1 hour, the DNA-bound proteins were subjected to immunoblotting to detect Smad3.

RNA Interference.

The cells with 50% confluence were transfected with either cPLA2α siRNA or PPAR-γ siRNA or Smad2/3 siRNA, or a 21-nucleotide irrelevant RNA duplex as a control siRNA using Lipofectamine 2000. After transfection, the cells were cultured in serum-free medium for 48 hours. Depletion of cPLA2α or PPAR-γ or Smad2/3 was confirmed by immunoblotting for further experiments.

AA Release and PGE2 Production.

To measure AA release, the cells with 80% confluence in six-well plates were labeled with 0.5 μCi/mL [3H]-AA (218 Ci/mmol) in serum-supplemented medium for 18 hours. After washing three times with serum-free medium, the cells were exposed to TGF-β1 in the absence or presence of the cPLA2α-specific inhibitor pyrrolidine derivative, the p38 MAPK inhibitor SB203580, or the p42/44 MAPK inhibitor PD98059. At the end of each treatment, the media were collected and centrifuged to remove the suspending cells. In addition, 0.5 mL media was used for scintillation counting to measure 3H activity.

To measure PGE2 production, the serum-starved cells with 80% confluence in six-well plates were exposed to TGF-β1 in the absence or presence of the inhibitors of cPLA2α, COX-2, p38 MAPK, or p42/44 MAPK in serum-free medium. At the end of each treatment, the spent medium was collected. A 100-μL centrifuged sample was analyzed for PGE2 production using the PGE2 enzyme immunoassay system (Amersham Biosciences) according to the manufacturer's protocol.

Generation of cPLA2α Transgenic Mice.

Transgenic mice with targeted expression of cPLA2α in the hepatocytes were developed by using the well-established albumin promoter-enhancer driven vector. To construct the albumin promoter-cPLA2α transgene, a 2.8-kb human cPLA2α complementary DNA (cDNA) containing the entire coding region of human cPLA2α was inserted into the first exon of the human growth hormone gene controlled by the mouse albumin ehancer/promoter.24, 25 After the confirmation of its overexpression in vitro, this transgene was microinjected into mouse zygotes (B6SJL/F1 eggs) at the transgenic core facility of the University of Pennsylvania according to our contract. The produced transgenic lines were brought back to the University of Pittsburgh animal facility for propagation. The transgenic lines were maintained by back-crossing to the C57Bl/6 wild-type mice, and the transgenic mice were identified with genotyping from the genomic DNA of tails. The cPLA2α transgenic mice develop normally with no significant liver inflammation or histological abnormality under normal housing conditions, although the cPLA2α transgenic mice show slightly higher body weight and liver weight compared with WT mice (body weight 25.82 ± 1.23 in cPLA2α-Tg versus 22.63 ± 0.68 in WT; liver weight 1.30 ± 0.10 in cPLA2α-Tg versus 1.13 ± 0.06 in WT). The animals were kept at 22°C under a 12-hour light/dark cycle and received food and water ad libitum. The handling of mice and experimental procedures were conducted in accordance with experimental animal guidelines. The cPLA2α transgenic mice used in this study were derived from one transgenic line that was back-crossed to C57Bl/6 WT mice for 10 consecutive generations.

Statistical Analysis.

All values were expressed as the mean and standard deviation (SD). The statistical significance of differences between groups was analyzed with the homoscedastic Student t test, and P < 0.05 was considered statistically significant. Single and double asterisks represent P < 0.05 and P < 0.01, respectively.

Results

We examined the effect of TGF-β1 on cPLA2α phosphorylation and protein expression in three human hepatocellular carcinoma cell lines (Hep3B, Huh7, and HepG2). Treatment of these cells with TGF-β1 (5 ng/mL) induced a rapid phosphorylation of cPLA2α, occurring within 15 minutes (Fig. 1A). In contrast, TGF-β1 treatment had no effect on the expression of cPLA2α and COX-2 proteins (Supporting Fig. 1). Consistent with its effect on cPLA2α phosphorylation, TGF-β1 treatment significantly increased AA release and PGE2 production in these cells (Fig. 1B). These results show that TGF-β1 activates cPLA2α phosphorylation and increases AA release and PGE2 synthesis in these cells.

Figure 1.

TGF-β1 activates AA signaling cascade through cPLA2α phosphorylation in transformed human hepatocytes. (A) Effect of TGF-β1 on cPLA2α phosphorylation. Left: TGF-β1 increases phosphorylation of cPLA2α. Hep3B, Huh7, and HepG2 cells were treated with 5 ng/mL TGF-β1 for the indicated periods. Cell lysates were then collected to determine the cPLA2α phosphorylation by immunoprecipitation and western blot analysis. Right: The p38 MAPK inhibitor SB203580 and the p42/44 MAPK inhibitor PD98059 inhibited TGF-β1–induced cPLA2α phosphorylation. Hep3B, Huh7, and HepG2 cells were treated with either 10 μM SB203580 or 10 μM PD98059 in serum-free medium for 2 hours prior to the stimulation with 5 ng/mL TGF-β1 for 15 minutes. Cell lysates were then collected to determine the cPLA2α phosphorylation by immunoprecipitation and western blot analysis. (B) AA release and PGE2 production. Left: The cPLA2α inhibitor pyrrolidine derivative, the p38 MAPK inhibitor SB203580, and the p42/44 MAPK inhibitor PD98059 inhibited TGF-β1–induced AA release. Hep3B, Huh7, and HepG2 cells prelabeled with 0.5 μCi/mL [3H]-AA were treated with 5 ng/mL TGF-β1 for 60 minutes in the absence or presence of 2 μM pyrrolidine derivative, 10 μM SB203580, or 10 μM PD98059. Media were then collected for the measurement of AA release as indicated in the Materials and Methods. Data are presented as the mean ± standard deviation of four experiments (*P < 0.01 versus control; **P < 0.05 versus TGF-β1 treatment). Right: The cPLA2α inhibitor pyrrolidine derivative, the COX-2 inhibitor NS 398, the p38 MAPK inhibitor SB203580, and the p42/44 MAPK inhibitor PD98059 inhibited TGF-β1–induced PGE2 production. Hep3B, Huh7, and HepG2 cells were treated with 5 ng/mL TGF-β1 for 8 hours in the absence or presence of 2 μM pyrrolidine derivative, 25 μM NS398, 10 μM SB203580, or 10 μM PD98059 in serum-free medium. Media were then collected for the measurement of PGE2 production as indicated in the Materials and Methods. Data are presented as the mean ± standard deviation of four experiments (*P < 0.01 versus control; **P < 0.05 versus TGF-β1 treatment). (C) TGF-β1 activates p38 MAPK, p42/44 MAPK, Smad2, and Smad3. Hep3B, Huh7, and HepG2 cells were treated with 5 ng/mL TGF-β1 for the indicated periods. Cell lysates were then collected to determine phosphorylated and total p38 MAPK, phosphorylated and total p42/44 MAPK, or phosphorylated and total Smad2/3 by western blot analysis.

Because cPLA2α is phosphorylated by protein kinases including p38 MAPK and ERK1/2 (p44/42 MAPK), we also examined the effect of TGF-β1 on p38 MAPK and ERK1/2 activation in these cells. TGF-β1 treatment induced the phosphorylation of p38 MAPK and ERK1/2 as well as Smad2/3 (Fig. 1C). These findings, along with the significant increase of Smad2/3 reporter activity by TGF-β1 (Fig 4C), indicate intact TGF-β–initiated signaling in these cells. The involvement of p38 MAPK and ERK1/2 in TGF-β1–induced cPLA2α phosphorylation is demonstrated by the fact that TGF-β1–induced cPLA2α phosphorylation in these cells was inhibited by the p38 MAPK inhibitor SB203580 and by the MEK1/2 inhibitor PD98059 (Fig. 1A). Consistent with this, TGF-β1–induced AA release and PGE2 production were also inhibited by SB203580, PD98059, and the cPLA2α inhibitor pyrrolidine (Fig. 1B). These findings demonstrate the role of p38 MAPK and ERK1/2-mediated cPLA2α activation in TGF-β1–induced AA release and PGE2 synthesis. In addition, the TGF-β1–induced PGE2 production was also inhibited by the selective COX-2 inhibitor NS-398 (Fig. 1B), although the level of COX-2 expression was not altered (Supporting Fig. 1). Taken together, these findings suggest that TGF-β1 induces AA release for PGE2 production via p38 MAPK and ERK1/2-mediated cPLA2α phosphorylation.

Further experiments were performed to determine whether cPLA2α overexpression or PGE2 treatment could prevent TGF-β1–induced inhibition of cell growth. As shown in Fig. 2, overexpression of cPLA2α in Hep3B cells prevents TGF-β1–induced inhibition of growth; PGE2 treatment of Hep3B cells as well as rat primary hepatocytes also prevented TGF-β1–induced inhibition of cell growth. The observation that cPLA2α overexpression prevents TGF-β1–induced caspase-3 cleavage in Hep3B cells suggests the role of cPLA2α for prevention of TGF-β–induced apoptosis. These data indicate that cPLA2α signaling pathway is able to counteract the growth inhibitory effect of TGF-β.

Figure 2.

cPLA2α signaling prevents TGF-β1–induced inhibition of cell growth. (A) Overexpression of cPLA2α prevents TGF-β1–induced inhibition of Hep3B cell growth. Hep3B cells were transfected with either MT2 control vector or cPLA2α in MT2. After transfection, the cells were treated with different concentrations of TGF-β1 in serum-free medium for 48 hours. Cell growth was determined with WST-1 reagent. Data are presented as the mean ± SD of six independent experiments (*P <0.01 versus MT2 control vector cells without TGF-β1 treatment; **P <0.05 versus MT2 control vector cells treated with the same concentration of TGF-β1). Western blots in the right panel show successful overexpression of cPLA2α in Hep3B cells transfected with the cPLA2α expression vector; the protein levels of PPAR-γ, TGF-βRI, TGFβRII, Smad2, Smad3, and Smad4 were not altered. (B) PGE2 effect in Hep3B cells. Hep3B cells were treated with 5 ng/mL TGF-β1 in the absence or presence of 10 μM PGEmath image in serum-free medium for 48 hours. Cell growth was determined with the WST-1 reagent. Data are presented as the mean ± SD of six independent experiments. PGE2 increased the growth of Hep3B cells (*P < 0.05). TGF-β1 significantly inhibited the growth of Hep3B cells (**P < 0.01); this effect was partially blocked by cotreatment with 10 μM PGEmath image (***P < 0.05). (C) PGE2 effect in rat primary hepatocytes. Rat primary hepatocytes were treated with 5 ng/mL TGF-β1 in the absence or presence of 10 μM PGEmath image in serum-free medium for 48 hours. Cell growth was determined with [3H]-thymidine incorporation assay (left panel). Data are presented as the mean ± SD of three independent experiments (*P <0.01 versus control; **P <0.05 versus TGF-β1 treatment). Western blots in the right panel show the protein levels of cPLA2α, PPAR-γ, TGFβRI, TGFβRII, Smad2, and Smad3 in rat primary hepatocytes. (D) cPLA2α overexpression prevents TGF-β1–induced caspase-3 activation. After transfection with the cPLA2α expression plasmid or the MT-2 control vector, Hep3B cells were cultured in serum-free medium for 24 hours and then treated with either vehicle or 5 ng/mL TGFβ1 for 12 hours. The cell lysates were then obtained for western blotting analysis to determine caspase-3 activation (β-actin was used as the loading control).

We next investigated whether cPLA2α-mediated AA release might influence Smad transcriptional activity. Hep3B cells were transiently transfected with the cPLA2α expression plasmid or the control plasmid MT-2 with cotransfection of the p3TP-Lux reporter construct (containing Smad2/3-responsive element) and the cell lysates were obtained to determine the luciferase reporter activity. As shown in Fig. 3A, overexpression of cPLA2α significantly inhibited Smad2/3 transcriptional activity. Accordingly, depletion of cPLA2α by siRNA significantly enhanced Smad2/3 transcriptional activity (Fig. 3B). These findings reveal an important role of cPLA2α for modulation of Smad2/3 transcription activity.

Figure 3.

Effect of cPLA2α on p3TP and PPRE reporter activities. (A) cPLA2α overexpression. Hep3B, HepG2, and Huh7 cells were transiently transfected with the cPLA2α expression plasmid or MT2 control plasmid with cotransfection of either the p3TP-Luc reporter vector as shown in the upper left panel or the PPRE-Luc reporter vector as shown in the upper right panel. Cell lysates were obtained to determine luciferase activity. Data are presented as the mean ± SD of three independent experiments (*P <0.01 versus corresponding control). (B) cPLA2α siRNA. Hep3B, HepG2, and Huh7 cells were transiently transfected with either cPLA2α siRNA or control siRNA with cotransfection of p3TP-Luc reporter vector. Cell lysates were then obtained to determine luciferase activity. Data are presented as the mean ± SD of three independent experiments (*P <0.01 versus corresponding control siRNA). RNA suppression of cPLA2α expression significantly increased the expression PAI-1, a Smad2/3 target gene.

PPAR-γ is a ligand-activated nuclear transcription factor regulating the expression of target genes by binding to specific peroxisome proliferator response elements (PPREs) or by interacting with other intracellular signaling molecules.26 The activity of PPAR-γ is regulated by several ligands, including thiazolidinediones, 15-deoxy-Δ,12, 14 prostaglandin J2, and AA, among others. Consistent with our previous study showing that cPLA2α is able to activate PPAR-γ in other cells,17 cPLA2α overexpression was found to increase PPRE reporter (containing PPAR response element) activity in all three hepatocellular carcinoma cell lines used in this study (Hep3B, Huh7, and HepG2) (Fig. 3A). Because PPAR-γ is known to bind and inhibit Smad3 in vitro27 and cPLA2α is able to activate PPAR-γ, we postulated that cPLA2α might inhibit Smad3 through activation of PPAR-γ.

To document the direct effect of PPAR-γ on Smad activation, Hep3B, HepG2, and Huh7 cells were cotransfected with the PPAR-γ expression plasmid and the p3TP-Lux reporter construct containing the Smad2/3 response element. As shown in Fig. 4A, overexpression of PPAR-γ partially inhibited Smad transcriptional activity in those cells. Accordingly, activation of PPAR-γ by its ligands (ciglitazone and piglitazone) significantly inhibited TGF-β1–induced Smad activation in Hep3 cells; this effect was observed with or without Smad3 overexpression (Fig. 4B,C). In contrast, siRNA inhibition of PPAR-γ augmented TGF-β1–mediated Smad transcription (Fig. 5A). In the transfection experiments with a reporter construct containing PPRE, we observed an approximately two-fold increase of PPRE reporter activity in Hep3B cells after PPAR-γ ligand treatment (Fig. 4B), suggesting that the endogenous PPAR-γ protein in these cells is functional.

Figure 4.

The effect of PPAR-γ overexpression and ligands on p3TP and PPRE reporter activities. (A) PPAR-γ overexpression. Hep3B, HepG2, and Huh7 cells were transiently transfected with the PPAR-γ expression plasmid or pcDNA control plasmid with cotransfection of either PPRE-Luc reporter vector as shown in the left upper panel or the p3TP-Luc reporter vector as shown in the right upper panel. Cell lysates were obtained to determine luciferase activity. Data are presented as the mean ± SD of three independent experiments (*P <0.01 versus corresponding vector control). (B) PPAR-γ ligands. Left: Hep3B cells were transiently transfected with PPRE-Luc reporter vector and then were treated with either 5 μM ciglitazone or 10 μM piglitazone in serum-free medium for 24 hours. Cell lysates were obtained to determine luciferase activity. Data are presented as the mean ± SD of three independent experiments (*P <0.01 versus control). Right: Hep3B cells were transiently transfected with p3TP-Luc reporter vector. After transfection, the cells were treated with 5 ng/mL TGF-β1 in the absence or presence of either ciglitazone or piglitazone in serum-free medium for 24 hours. Cell lysates were then obtained to determine luciferase activity. Data are presented as the mean ± SD of three independent experiments (*P <0.01 versus no TGF-β1 treatment; **P <0.05 versus TGF-β1 treatment alone). (C) Effect of ciglitazone on TGF-β1–induced p3TP reporter activity in cells with or without Smad3 overexpression. Hep3B cells were transiently transfected with either pcDNA control vector or Smad3 in pcDNA overexpression vector with cotransfection of the p3TP-Luc reporter vector. After transfection, the cells were treated with 5 ng/mL TGF-β1 in the absence or presence of 5 μM ciglitazone in serum-free medium for 24 hours; cell lysates were then obtained to determine luciferase activity. Data are presented as the mean ± SD of three independent experiments. Overexpression of Smad3 significantly increased the p3TP reporter activity compared with pcDNA control vector (*P < 0.01). TGF-β1 significantly increased the p3TP reporter activity in pcDNA- or Smad3-transfected cells compared with each own control (**P < 0.01). Ciglitazone significantly blocked TGF-β1–induced increase of p3TP reporter activity in cells transfected with either pcDNA or Smad3 in pcDNA (***P <0.01).

Figure 5.

The effect of PPAR-γ depletion on p3TP reporter activity and cell growth. (A) PPAR-γ siRNA. Hep3B, HepG2, and Huh7 cells were transiently transfected with either PPAR-γ siRNA or control siRNA with cotransfection of p3TP-Luc reporter vector. Cell lysates were obtained to determine luciferase activity. Data are presented as the mean ± SD of three independent experiments (*P < 0.01 versus control siRNA). RNAi suppression of PPAR-γ expression significantly increased expression of the Smad2/3 target gene, PAI-1. (B) Depletion of cPLA2α and PPAR-γ inhibits cell growth and involvement of Smad2/3. Hep3B cells were transfected with cPLA2α siRNA or PPAR-γ siRNA, with or without Smad2/3 siRNA. After transfection, the cells were cultured in serum-free medium for 48 hours. Cell growth was determined with WST-1 reagent. Data are presented as the mean ± SD of six independent experiments (*P < 0.01 versus control siRNA; **P <0.05 versus cPLA2α siRNA or PPAR-γ siRNA). Western blots in the lower panel show successful depletion of cPLA2α, PPAR-γ or Smad2/3 in cells transfected with the corresponding siRNA.

To further evaluate the role of cPLA2α and PPAR-γ in Smad activation and hepatic cell growth, additional experiments were performed to determine whether depletion of endogenous cPLA2α and PPAR-γ might inhibit cell growth via Smad2/3. As shown in Fig. 5B, depletion of either cPLA2α or PPAR-γ significantly reduced cell growth, and this effect was blocked by Smad2/3 siRNA. These findings suggest the involvement of Smad2/3 in cPLA2α/PPAR-γ depletion-induced inhibition of cell growth. Furthermore, the cPLA2α product (AA) and the PPAR-γ ligands (ciglitazone and piglitazone) inhibited TGF-β1–induced binding of Smad3 to its DNA response element (Fig. 6A). Therefore, Smad3 is a downstream target of cPLA2α/PPAR-γ in hepatic cells.

Figure 6.

AA and PPAR-γ ligands block TGF-β1–induced Smad3 binding to its DNA response element. A comparison of cPLA2α and PPAR-γ expression in different cell lines is shown. (A) Hep3B cells were treated with AA, ciglitazone, or piglitazone in the absence or presence of TGF-β1 for 30 minutes. The binding of Smad3 to its DNA response element was analyzed by DNA–protein binding assay. (B) Hep3B, Huh7, and HepG2 cells were treated with different concentrations of TGF-β1 for different periods as indicated. Cell growth was determined with WST-1 reagent. Data are presented as the mean ± SD of six independent experiments. TGF-β1 inhibited the growth of Hep3B, but not Huh7 and HepG2 cells. The Western blots in the lower panel showed the protein levels of cPLA2α, PPAR-γ, TGFβRI, TGFβRII, and Smad2/3 in these cells.

We have found that cPLA2α activates PPAR-γ in all three hepatic cell lines used in this study. However, these cell lines respond differently to TGF-β treatment; whereas TGF-β1 significantly inhibited the growth of Hep3B cells, it had minimal growth inhibitory effect in Huh7 or HepG2 cells under the same experimental condition (Fig. 6B). The exact mechanism for such a differential effect among different cell lines is complex; however, it is possible that this may relate partly to the low level of PPAR-γ expression in Hep3B cells (hence a sensitivity to TGF-β–induced inhibition of growth) and the high level of PPAR-γ in Huh7 and HepG2 cells (hence a resistance to TGF-β–induced inhibition of growth).

To further address the role of cPLA2α in TGF-β–induced hepatocyte growth regulation, we generated transgenic mice with targeted expression of the cPLA2α in the liver (Fig. 7) and the produced animals were used to determine TGF-β–induced inhibition of hepatocyte growth. Primary hepatocytes were isolated from the cPLA2α transgenic or WT mice and the cultured cells were treated with different concentrations of TGF-β1 in serum-free medium to determine [3H]-thymidine incorporation. As shown in Fig. 7D, although TGF-β1 significantly inhibited the growth of hepatocytes from WT mice, this effect was attenuated in cPLA2α overexpressed hepatocytes. Thus, overexpression of cPLA2α in hepatocytes renders the cells resistant to TGF-β1–induced inhibition of growth.

Figure 7.

Overexpression of cPLA2α in hepatocytes prevents TGF-β1–induced mitoinhibition. (A) Schematic representation of the human cPLA2α transgene. Transgenic mice with targeted expression of cPLA2α in hepatocytes were developed using the well-established albumin promoter-enhancer driven vector. The 8-kb ALB-cPLA2α transgene consists of the 2.8-kb human cPLA2α cDNA (white box) inserted into the first exon of the human growth hormone gene (black boxes) controlled by the mouse albumin enhancer/promoter (cross-hatched ovals), and possessing a human growth hormone polyadenylation site (cross-hatched box). (B) PCR analysis of tail genomic DNA for cPLA2α transgene. cPLA2α cDNA served as the positive control; the genomic DNA from WT mice served as a negative control. An 860-bp cPLA2α transgene product was detected by polymerase chain reaction using specific primer pairs (forward primer, cPLA2αF, 5′-TGGCCAACATCAACTTCAGA-3′; reverse primer, GHE1R, 5′TTACCTGCAGCCATTGCCGCTAGTGAG-3′) derived from the pALB-cPLA2α transgene. (C) Western blotting analysis of cPLA2α protein levels in liver tissues from cPLA2α transgenic mice. Equal amounts of the liver tissue proteins from WT and cPLA2α transgenic (TG) mice littermates underwent sodium dodecyl sulfate–polyacrylamide gel electrophoresis and western blot analysis using anti-human cPLA2α antibody (with glyceraldehyde 3-phosphate dehydrogenase as a loading control). (D) TGF-β1–mediated mitoinhibition of primary hepatocytes from cPLA2α transgenic and WT mice. Primary hepatocytes isolated from WT or cPLA2α transgenic mice were treated with different concentrations of TGF-β1 in serum-free medium for 48 hours. Cell proliferation was determined by [3H]-thymidine incorporation assay. Data are presented as the mean ± SD of three independent experiments (*P < 0.01 versus WT cells without TGF-β1 treatment; **P <0.05 versus the same TGF-β1 treatment of WT cells).

Discussion

In the liver, TGF-β is well-known to inhibit hepatocyte proliferation and induce hepatocyte apoptosis.28-34 Quiescent liver usually contains only modest amounts of TGF-β but injury to the liver results in the production of TGF-β, most prominently by nonparenchymal cells, including hepatic stellate cells and Kupffer cells. TGF-β1 regulates hepatocyte growth by inducing cell cycle arrest or apoptosis, both in vitro and in vivo.28-34 In primary hepatocyte cultures, TGF-β inhibits DNA synthesis from normal and regenerating livers by blocking the transition from the G1 to the S phase of the cell cycle. After a two-thirds partial hepatectomy, TGF-β1 messenger RNA expression increases, and TGF-β is the ostensible inhibitory peptide for hepatocyte replication and liver regeneration.35 Intravenous administration of mature TGF-β to rats has been shown to reduce [3H]thymidine incorporation in hepatocytes after partial hepatectomy, although inhibition of hepatocyte DNA synthesis was transient, because complete regeneration of the liver still occurred by 8 days.30 On the other hand, intravenous administration of adenoviral vector expressing active TGF-β1 potently inhibits liver regeneration in rats after standard two-thirds partial hepatectomy, ultimately leading to animal death.36 Thus, high levels of expression of active TGF-β1 is able to effectively inhibit hepatocyte growth in vivo.

In consideration of the key role of TGF-β in the regulation of hepatocyte growth and liver fibrosis, it is not surprising that TGF-β is causatively involved in hepatocarcinogenesis. Given its antiproliferative and proapoptotic role in the liver, TGF-β1 could be expected to act as a tumor suppressor. Indeed, mice heterozygous for the deletion of TGF-β1 or TGFβRII were found to be more susceptible to DEN-induced hepatocarcinogenesis when compared with their WT littermates37, 38 (mice homozygous for the deletion of TGF-β1 or TGFβRII are lethal). Enhanced hepatocarcinogenesis is also observed in transgenic mice overexpressing a dominant negative TGFβRII39 or in mice heterozygous for deletion of the Smad adaptor protein, embryonic liver fodrin.40 Consistent with these observations, forced expression of Smad3 in the liver has been shown to inhibit DEN-induced hepatocarcinogenesis.41 However, there is also evidence suggesting that TGF-β may promote hepatic carcinogenesis42-44 and that late stage human liver cancers often show overexpression of TGF-β.45, 46 The opposing effects of TGF-β on liver cancer growth may be explained by the fact that TGF-β functions as a tumor suppressor early in tumorigenesis when epithelial cell responsiveness to TGF-β is still relatively normal. It is possible that during multistage tumorigenesis, the mitoinhibitory effect of TGF-β becomes lost, either through mutation of the TGF-β signaling molecules or by subversion of the normal signaling pathway due to activation of other molecules.1, 2, 47-49 Because mutation of TGF-β signaling molecules occurs only in a minority of human hepatocellular cancers and the prostaglandin signaling pathway is active in tumor cells,15, 50 we postulate that disruption of TGF-β–mediated inhibition of cell growth by PG cascade may be an important mechanism for the regulation of cell growth and carcinogenesis.

The current study shows that TGF-β regulates the growth of primary and transformed hepatocytes through concurrent activation of Smad-mediated gene transcription and phosphorylation of cPLA2α (Fig. 8). Our findings suggest that the level and activation status of cPLA2α/PPAR-γ signaling in hepatic cells likely represents a key factor that determines the cellular response to TGF-β. It is possible that activation of cPLA2α/PPAR-γ signaling may in part explain the loss of responsiveness of neoplastic cells to the antiproliferative actions of TGF-β (due to suppression of Smad2/3 activity). Further studies are warranted to determine the exact role of this interaction in different stages of hepatocarcinogenesis.

Figure 8.

Schematic illustration of the interaction between cPLA2α and TGF-β signaling pathways in hepatic cells.

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