Peroxisome proliferator-activated receptor δ (PPARδ) is a nuclear transcription factor that is recently implicated in tumorigenesis besides lipid metabolism. This study describes the cross-talk between the PPARδ and prostaglandin (PG) signaling pathways that coordinately regulate human hepatocellular carcinoma (HCC) cell growth. Activation of PPARδ by its pharmacologic ligand, GW501516, enhanced the growth of three human HCC cell lines (HuH7, HepG2, and Hep3B), whereas inhibition of PPARδ by small interfering RNA prevented growth. PPARδ activation up-regulates the expression of cyclooxygenase (COX)-2, a rate-limiting enzyme for PG synthesis, and tumor growth. PPARδ activation or PGE2 treatment also induced the phosphorylation of cytosolic phospholipase A2α (cPLA2α), a key enzyme that releases arachidonic acid substrate for PG production via COX. Activation of cPLA2α by the calcium ionophore A23187 enhanced PPARδ binding to PPARδ response element (DRE) and increased PPARδ reporter activity, which was blocked by the selective cPLA2α inhibitors. Consistent with this, addition of arachidonic acid to isolated nuclear extracts enhanced the binding of PPARδ to DRE in vitro, suggesting a direct role of arachidonic acid for PPARδ activation in the nucleus. Thus, PPARδ induces COX-2 expression and the COX-2–derived PGE2 further activates PPARδ via cPLA2α. Such an interaction forms a novel feed-forward growth-promoting signaling that may play a role in hepatocarcinogenesis. (Cancer Res 2006; 66(24): 11859-68)

Hepatocellular carcinoma (HCC) is the fifth most common cancer and the third leading cause of cancer-related mortality worldwide (1). The pathogenesis of hepatic carcinogenesis remains incompletely understood but is believed to involve sequential events, including chronic inflammation, hepatocyte hyperplasia, dysplasia, and, ultimately, malignant transformation (2). HCC usually develops in the presence of continuous inflammation and hepatocyte regeneration in the setting of chronic hepatitis and cirrhosis, although the molecular mechanisms linking chronic inflammation to malignant transformation remain to be further defined. Several lines of evidence suggest that mediators of inflammation, such as prostaglandins (PG), are implicated in hepatic carcinogenesis (see ref. 3 for review). For example, increased cyclooxygenase (COX)-2 expression has been found in human and animal HCCs (410). Elevated levels of PGs, most notably PGE2, have also been detected in HCC (11). Overexpression of COX-2 or treatment with exogenous PGE2 increases human HCC cell growth and invasiveness (9, 12). The COX inhibitors, nonsteroidal anti-inflammatory drugs (NSAID), inhibit the proliferation and induce apoptosis in cultured HCC cells and in animal models of hepatocarcinogenesis (3), although these inhibitors are known to mediate effects through both COX-dependent and COX-independent mechanisms.

The first step in the formation of PGs is the liberation of arachidonic acid (5,8,11,14-eicosatetraenoic acid) from membrane-bound phospholipids, usually by the action of phospholipase enzymes, primarily phospholipase A2s (PLA2). Although there exist multiple different isoforms of PLA2s in cells, it is the 85-kDa cytosolic PLA2α (cPLA2α) that most commonly supplies the arachidonic acid for PG production by COX (13, 14). Two isoforms of COXs have been identified, COX-1 and COX-2, both catalyzing the conversion of arachidonic acid into endoperoxide intermediates that are ultimately converted by specific synthases to prostanoids, including PGE2, the most abundant PG in human neoplastic epithelial cells (15, 16). Whereas COX-1 is constitutively expressed in most cells, COX-2 is highly induced by inflammatory cytokines/chemokines, growth factors, oncogene activation, and tumor promoters, thus contributing to the enhanced PG production when these signaling pathways are activated in inflammatory and neoplastic diseases (15, 16). PGs transduce signals mainly through binding to their specific G protein–coupled receptors along the plasma membrane. Recently, eicosanoids have been shown to regulate cell functions through activation of peroxisome proliferator-activated receptors (PPAR), which belong to the superfamily of nuclear receptors that function as ligand-activated transcription factors.

PPARs regulate gene expression by binding with their heterodimeric partner retinoid X receptor to specific peroxisome proliferator response elements. Three different PPAR subtypes have been identified: PPARα, PPARδ (also termed as PPARβ), and PPARγ. PPARα is highly expressed in liver parenchymal cells and is implicated in lipid catabolism (1719). PPARγ is predominantly expressed in adipose tissue and plays an important role in adipocyte differentiation, insulin sensitization, and glucose homeostasis (20, 21). PPARβ/δ shows a ubiquitous expression in most tissues (18) and is implicated in fatty acid oxidation, cell differentiation, inflammation, cell motility, and cell growth (2232).

Recent studies suggest a potential role of PPARδ in carcinogenesis. For example, the expression of PPARδ is increased in colorectal cancer cells compared with normal colon epithelial cells (33, 34). Treatment of Apcmin mice with the PPARδ ligand GW501516 increased the number and size of intestinal polyps (35). On the other hand, disruption of PPARδ in human colon cancer cells by targeted homologous recombination reduced tumor growth when the PPARδ−/− cells were inoculated as xenografts in nude mice (36). These observations suggest a tumor-promoting role of PPARδ during intestinal carcinogenesis. Additionally, PPARδ has also been implicated in the growth of several other human cancers, including prostate and lung cell lines (37) and HCC cells (38). Activation of PPARδ by its synthetic agonist increases COX-2 expression in human HCC cells (38). Furthermore, PPARδ is a downstream gene of Wnt-β-catenin signal pathway and the target of NSAIDs (33, 39). PPARδ has also been shown to mediate the PGE2-induced intestinal adenoma growth (40). However, in spite of the documented tumor-promoting effect of PPARδ, there is also evidence suggesting that PPARδ might inhibit intestine tumor development (41). Thus, the precise role of PPARδ in tumorigenesis remains to be further defined.

This study was designed to evaluate the effect and mechanism of PPARδ in HCC cell growth. Our data show that PPARδ promotes human HCC cell growth through up-regulation of COX-2 gene transcription and PGE2 production. More importantly, the PPARδ-induced PGE2 subsequently phosphorylates and activates cPLA2α, thereby providing arachidonic acid for further PPARδ activation and PGE2 production. Consequently, the interplay between PPARδ and cPLA2α/COX-2/PGE2 signaling pathways forms a positive feedback loop promoting HCC cell growth.

Materials. Cell culture medium and LipofectAMINE Plus reagent were purchased from Invitrogen (Carlsbad, CA). Cell proliferation reagent WST-1 was purchased from Roche Applied Science (Indianapolis, IN). Luciferase Assay System and reporter lysis buffer were from Promega Corp. (Madison, WI). Antibody providers are as follows: anti-COX-2 (Cayman Chemical Co., Ann Arbor, MI); anti-cPLA2, anti-PPARδ, and anti-epidermal growth factor receptor (EGFR; Santa Cruz Biotechnology, Santa Cruz, CA); anti-phosphorylated cPLA2α (Ser505) and Akt Kinase Assay kit (Cell Signaling Technology, Danvers, MA); anti-phosphorylated EGFR (BD Biosciences, San Jose, CA); and anti-β-actin (Sigma, St. Louis, MO). Chemiluminescence detection reagent was from Amersham Biosciences (Piscataway, NJ). PPARδ agonist GW501516 was purchased from Cayman Chemical. PGE2, indomethacin, arachidonic acid, A23187, the cPLA2 inhibitors pyrrolidine derivatives and AACOCF3, the EGFR tyrosine kinase inhibitor AG1478, the p38 kinase inhibitor SB203580, the protein kinase C (PKC) inhibitor bisindolylmaleimide I, the phosphatidylinositol 3-kinase (PI3K) inhibitor LY294002, and the p44/42 mitogen-activated protein kinase (MAPK) inhibitor PD98059 were purchased from Calbiochem (San Diego, CA). The PGE2 enzyme immunoassay system was purchased from Amersham Biosciences. The PPARδ Transcription Factor Assay kit was from Cayman Chemical. The nuclear extraction kit was purchased from Sigma. Small interfering RNA (siRNA)-PPARδ, siRNA-COX-2, and siRNA-control were from Dharmacon, Inc. (Lafayette, CO). The 5′-biotinylated PPARδ response element (DRE) oligonucleotides were synthesized by Sigma-Genosys (Woodland, TX), and the unlabeled DRE oligonucleotides were from Integrated DNA Technologies, Inc. (Coralville, IA). The immobilized streptavidin beads were purchased from Pierce (Rockford, IL). Poly(dI-dC).poly(dI-dC) was from Amersham Biosciences.

Cell culture and proliferation assay. Three hepatocarcinoma cell lines, HuH7, HepG2, and Hep3B, were cultured in Eagle's Minimum Essential Medium with 10% fetal bovine serum (FBS). Cell growth was determined using the cell proliferation reagent WST-1, which is a tetrazolium salt cleaved by mitochondrial dehydrogenases in viable cells. Briefly, the cells (3,000 per well) were seeded on 96-well plate and incubated at 37°C overnight. The cells were then treated with GW501516 or transfected with siRNAs for indicated times. WST-1 (10 μL) was subsequently added to each well, and the culture was continued for 30 minutes to 4 hours before measurement of A450 nm using an automatic ELISA plate reader.

Transient transfection and luciferase reporter assay. Cells were seeded in six-well plate in culture medium containing 10% FBS the day before transfection. On the following day, the cells in each well (70–80% confluence) were transfected with 1 μg of plasmid using LipofectAMINE Plus reagent (Plus reagent, 6 μL; LipofectAMINE, 4 μL) or with siRNA using LipofectAMINE 2000 reagent in serum-free medium. After 3 hours of transfection, the transfection medium was replaced with culture medium containing 10% FBS. At the end of indicated treatment, the cells were washed twice in ice-cold PBS and lysed with reporter lysis buffer on ice for 20 minutes. The cells were then scraped down and spun at 14,000 rpm for 10 minutes in cold room. The supernatant was collected for luciferase activity assay using a Berthold AutoLumat LB 953 luminometer (Berthold, Nashua, NH).

Preparation of whole-cell lysate and immunoblotting. HuH7 cells were grown on six-well plates and treated with different concentration of GW501516 or other indicated reagents in FBS-free medium. The vehicle, DMSO, was added to the control culture. Following treatment for indicated times, the cells were washed twice with cold PBS and scraped down. The cell pellets were washed two more times with cold PBS and then resuspended in homogenization buffer containing 50 mmol/L HEPES (pH 7.55), 1 mmol/L EDTA, 1 mmol/L DTT, and 1 mmol/L mammalian protease inhibitor cocktail (Sigma). The cell suspension was placed on ice and sonicated four times for 15 seconds. The samples were then centrifuged at 14,000 rpm for 10 minutes at 4°C, and the supernatants were collected as whole-cell lysate. Total protein concentration was measured by bicinchoninic acid (BCA) reagent (Pierce). Cell lysate was aliquoted and frozen at −80°C until use. For immunoblotting, 20 to 35 μg protein was separated on 4% to 20% Tris-glycine gels and the separated proteins were electrophoretically transferred onto the nitrocellulose membrane (Bio-Rad, Hercules, CA). Nonspecific binding was blocked with 5% nonfat milk dissolved in 0.5% Tween 20 in PBS (PBST) for 1 hour at room temperature. The membrane was then incubated overnight with primary antibodies (1:1,000 dilution for COX-2, EGFR, phosphorylated EGFR, Akt, phosphorylated Akt, and β-actin; 1:2,000 dilution for PPARδ) in 5% milk PBST. Following repeated washing with PBST the next day, the membranes were incubated with the horseradish peroxidase (HRP)-conjugated secondary antibody (1:10,000 dilution) for 1 hour at room temperature. After washing, the blots were developed using the enhanced chemiluminescence (ECL) Western blotting detection system and exposed to Eastman Kodak (Rochester, NY) MR radiographic films.

Immunoprecipitation and Western blotting for cPLA2α phosphorylation. To immunoprecipitate cPLA2α, 300 μL of whole HuH7 cell lysate (∼50 μg protein) in a 1.5 mL Eppendorf tube were precleared with 20 μL protein A/G agarose (Santa Cruz Biotechnology) for 1 hour at 4°C. The cleared cell lysate was then incubated with 5 μL mouse anti-human cPLA2α monoclonal antibody at 4°C for 3 hours with gentle agitation. Protein A/G agarose (20 μL) was then added, and the sample was kept at 4°C for 16 hours, with gentle agitation, to precipitate cPLA2α-antibody complex. The protein A/G agarose pellet was collected by centrifuge and washed four times with cold homogenization buffer at 4°C. SDS sample loading buffer (20 μL) was then added to the pellet, and the mixture was boiled for 5 minutes before SDS-PAGE using 4% to 20% Tris-glycine gels. After blocking nonspecific binding, the blot was incubated overnight with rabbit anti-phosphorylated cPLA2α (Ser505) antibody (1:1,000 dilution) in 5% milk PBST at 4°C. The HRP-conjugated donkey anti-rabbit antibody (1: 10,000 dilution) was used as the second antibody. Specific cPLA2α band was visualized by ECL Western blotting detection system.

Measurement of PGE2 production. HuH7 cells cultured in serum-free medium in six-well plates were treated as indicated in the text. The supernatant was collected and centrifuged to remove floating cells. Each sample (100 μL) was used to measure PGE2 level using the PGE2 enzyme immunoassay system as described previously (9, 42).

Purification of nuclear extract. HuH7 cells cultured in 100-mm dishes at 80% to 90% confluence were treated as described in the text. Following treatment, the cells were washed twice with ice-cold PBS and scraped with a rubber policeman. The cell pellet was then swelled in 5-fold volume of hypotonic buffer for 20 minutes on ice. Following homogenization using 27-gauge sterile needle on ice, the nuclei were pelleted by centrifugation at 600 × g for 10 minutes. The nuclei were then washed twice in the isotonic buffer and resuspended in HKMG buffer [10 mmol/L HEPES, (pH 7.9), 100 mmol/L KCL, 5 mmol/L MgCl2, 10% glycerol, 1 mmol/L DTT, 0.5% NP40] containing protease inhibitors and phosphatase inhibitors. The nuclei suspension was then subjected to sonication, and the cellular debris was removed by centrifugation at 14,000 rpm for 20 minutes at 4°C. The supernatant was collected as nuclear extract and frozen at −80°C until use. Aliquots of the nuclear extracts were used to quantitate the protein concentration using the BCA reagent.

ELISA-based PPARδ binding to its DNA response element. The experiments were carried out using the 96-well ELISA kit purchases from Cayman Chemical. Briefly, the oligonucleotide containing the PPARδ binding consensus sequence was immobilized onto the bottom of wells. Nuclear extract (50 μg) from treated cells or control cell was added to the dsDNA-coated well and incubated at 4°C overnight. After complete washing, PPARδ antibody was added and the samples were incubated at room temperature for 1 hour. The HRP-conjugated secondary antibody and developing solution were sequentially added, and the A655 nm value was determined.

Biotinylated DRE oligonucleotide precipitation assay. The assay was done as previously reported with modification (43). The nucleotide sequences of biotinylated DRE were 5′-GCGTGAGCGCTCACAGGTCAATTCG-3′ and 5′-CCGAATTGACCTGTGAGCGCTCACG-3′ (33). These two complementary strands were annealed in TEN buffer. After treatment, the cells were lysed by sonication in 200 μL HKMG buffer containing protease and phosphatase inhibitors. The cellular debris was removed by centrifugation. The cell extracts (50 μg) were precleared with 30 μL immobilized streptavidin-agarose beads for 1 hour at 4°C with gentle agitation. The cleared nuclear extracts were then incubated with 1 μg of biotinylated double-strand DRE and 10 μg of poly(dI-dC).poly(dI-dC) for 16 hours. DRE-bound protein was pulled down by incubating the samples with 35 μL of streptavidin-agarose beads for 1 hour at 4°C with gentle agitation. The agarose mixture was collected by centrifugation and washed four times with cold HKMG buffer. SDS sample buffer was then added to the pellet. The samples were boiled for 5 minutes and subjected to SDS-PAGE and Western blotting for PPARδ.

Statistical analysis. Statistical analysis was done using Microsoft Excel 2003 software. Comparisons were done using two-tailed unpaired Student's t test. Values of P < 0.05 were considered statistically significant.

The role of PPARδ in human HCC cell growth was first examined by using GW501516, a synthetic pharmacologic ligand that is selective for PPARδ with no effect on PPARα or PPARγ (even at dose as high as 10 μmol/L; refs. 22, 35, 44). As shown in Fig. 1A, GW501516 treatment increased the growth of three human HCC cell lines (HuH7, HepG2, and Hep3B). This effect was dose dependent (0.5–100 nmol/L) and was observed at different treatment periods (24–72 hours). The dose range is lower than that used in a previous study (38), possibly due to the difference in compound source and quality. Western blot analysis reveals that PPARδ protein is expressed in all these cells (Fig. 1A). The transcriptional function of PPARδ in these cells was verified by the observation that GW501516 treatment significantly increased the reporter activity of a luciferase promoter construct containing the DRE (33). Accordingly, inhibition of PPARδ by siRNA blocked the DRE activity in cells with or without GW501516 treatment (Fig. 1B). The direct effect of PPARδ on HCC growth was further supported by the observation that siRNA inhibition of PPARδ significantly reduced the growth of human HCC cells, both under spontaneous culture and in response to GW501516 treatment (Fig. 1B). These results document an important role of PPARδ in human HCC cell growth.

Figure 1.

PPARδ promotes human HCC cell growth and induces COX-2 gene expression. A, activation of PPARδ by GW501516 promotes human HCC cell growth. HuH7, HepG2, and Hep3B cells were seeded onto 96-well plate (3,000 per well) in 10% FBS medium and incubated overnight. After 24 hours of serum starvation, GW501516 (0.5–50 nmol/L) was added into the medium and the cells were cultured for 24, 48, and 72 hours. Cell proliferation was measured using WST-1 reagent as described in Materials and Methods. Columns, mean of six identical treatments; bars, SD. The level of PPARδ protein in human HCC cells (HuH7, HepG2, and Hep3B) was determined by SDS-PAGE and Western blot analysis (with the cholangiocarcinoma cell line CCLP1 as positive control). B, inhibition of PPARδ by siRNA reduces human HCC cell growth. Top left, GW501516 activates PPARδ. HuH7 cells (70% confluence in six-well plate) were cotransfected with 1 μg of human DRE reporter plasmid and 100 pmol siRNA using LipofectAMINE 2000 reagent. After 42 hours of transfection, the cells were then treated with GW501516 (10 nmol/L) or vehicle DMSO (1:10,000 dilution) in serum-free medium for 6 hours. At the end of treatment, the cells were washed twice with cold PBS and the cell lysates were obtained for luciferase activity assay. GW501516 treatment significantly increased the DRE reporter activity, and this effect was blocked by siRNA inhibition of PPARδ (2.5-fold versus 0.61-fold). **, P < 0.01, compared with control siRNA with vehicle treatment; ***, P < 0.01, compared with control siRNA with GW501516 treatment (n = 3). Top right, siRNA inhibition of PPARδ prevents HCC cell growth. HuH7 cells (3,000 per well) were seeded in 96-well plate and cultured overnight. The cells were transfected overnight with 5 pmol per well of siRNA-PPARδ or siRNA-control using LipofectAMINE 2000 reagent in 0.5% FBS medium. Following transfection, the cells were treated with GW501516 (10 nmol/L) or vehicle DMSO (1:10,000 dilution) for 24 hours. WST-1 reagent was used to determine cell growth. Columns, mean of three individual experiments; bars, SD. The cells transfected with the PPARδ siRNA exhibited significant reduction of growth either under basal culture conditions or in response to GW501516 treatment. *, P < 0.05, compared with control siRNA cells without GW501516 treatment; **, P < 0.01, compared with control siRNA cells without GW501516 treatment; ***, P < 0.01, compared with control siRNA cells with GW501516 treatment. Bottom, PPARδ protein levels in HuH7 cells with or without PPARδ siRNA transfection. β-Actin was used as the loading control. C, activation of PPARδ by GW501516 enhances COX-2 protein expression. Inhibition of PPARδ by siRNA reduces COX-2 protein level. Top, time course effect of GW501516 on COX-2 protein expression. HuH7 cells were pre-serum starved for 24 hours and treated with GW501516 (10 nmol/L) in serum-free medium for increasing times (2–8 hours). The cell lysates were obtained for SDS-PAGE and Western blot analysis to determine the level of COX-2 protein. The same blot was stripped and reprobed with β-actin antibody as loading control. Increased COX-2 protein was observed after GW501516 treatment for 4 to 8 hours. Middle, dose-dependent effect of GW501516 on COX-2 protein expression. HuH7 cells were incubated with GW501516 (1–50 nmol/L) in serum-free medium for 6 hours, and the whole-cell lysates were obtained for SDS-PAGE and Western blot analysis to determine the level of COX-2. Bottom, siRNA inhibition of PPARδ reduces COX-2 protein level. HuH7 cells (70% confluence) were transfected with PPARδ siRNA or nontargeted control siRNA for 48 hours. Cellular protein (30 μg) was subjected to SDS-PAGE and Western blotting to determine COX-2 protein, with β-actin as loading control. D, activation of PPARδ by GW501516 enhances COX-2 gene promoter activity and PGE2 production. Top, GW501516 treatment enhanced COX-2 gene promoter reporter activity. HuH7 cells (70% confluence in six-well plate) were cotransfected with the COX-2 promoter reporter plasmid (1 μg) plus control siRNA or PPARδ siRNA using LipofectAMINE 2000 reagent. After 48 hours, the cells were incubated with GW501516 (10 nmol/L) or vehicle DMSO (1:10,000 dilution) for 6 hours. Following washing twice with cold PBS, the cells were lysed and the cell lysates were obtained for luciferase activity assay. GW501516 treatment (10 nmol/L) induced a 2.2-fold increase of the COX-2 promoter reporter activity. **, P < 0.01 (n = 3). This effect was completely blocked by siRNA inhibition of PPARδ. ***, P < 0.01, compared with control siRNA plus GW501516 treatment (n = 3). PPARδ siRNA also inhibited the COX-2 promoter activity in cells without GW501516 treatment. *, P < 0.05, compared with control siRNA with vehicle treatment. Bottom left, GW501516 treatment increases PGE2 production. HuH7 cells (80% confluence in six-well plate, with overnight serum starvation) were treated with GW501516 (10–50 nmol/L) or vehicle for 6 hours in serum-free medium. At the end of treatment, the culture medium was collected and cell debris was removed by centrifugation for 25 minutes at 4°C. Medium (100 μL) was used for the measurement of PGE2. Columns, mean of three experiments (0.22 ng/mL for control versus 0.28, 0.32, and 0.45 ng/mL for 10, 20, and 50 nmol/L GW501516, respectively); bars, SD. Bottom right, siRNA inhibition of PPARδ prevents GW501516-induced PGE2 production. HuH7 cells were transfected with PPARδ siRNA or control siRNA for 48 hours followed by vehicle DMSO or GW501516 treatment for 6 hours. **, P < 0.01, compared with control siRNA plus vehicle treatment; ***, P < 0.01, compared with control siRNA plus GW501516 treatment (n = 3).

Figure 1.

PPARδ promotes human HCC cell growth and induces COX-2 gene expression. A, activation of PPARδ by GW501516 promotes human HCC cell growth. HuH7, HepG2, and Hep3B cells were seeded onto 96-well plate (3,000 per well) in 10% FBS medium and incubated overnight. After 24 hours of serum starvation, GW501516 (0.5–50 nmol/L) was added into the medium and the cells were cultured for 24, 48, and 72 hours. Cell proliferation was measured using WST-1 reagent as described in Materials and Methods. Columns, mean of six identical treatments; bars, SD. The level of PPARδ protein in human HCC cells (HuH7, HepG2, and Hep3B) was determined by SDS-PAGE and Western blot analysis (with the cholangiocarcinoma cell line CCLP1 as positive control). B, inhibition of PPARδ by siRNA reduces human HCC cell growth. Top left, GW501516 activates PPARδ. HuH7 cells (70% confluence in six-well plate) were cotransfected with 1 μg of human DRE reporter plasmid and 100 pmol siRNA using LipofectAMINE 2000 reagent. After 42 hours of transfection, the cells were then treated with GW501516 (10 nmol/L) or vehicle DMSO (1:10,000 dilution) in serum-free medium for 6 hours. At the end of treatment, the cells were washed twice with cold PBS and the cell lysates were obtained for luciferase activity assay. GW501516 treatment significantly increased the DRE reporter activity, and this effect was blocked by siRNA inhibition of PPARδ (2.5-fold versus 0.61-fold). **, P < 0.01, compared with control siRNA with vehicle treatment; ***, P < 0.01, compared with control siRNA with GW501516 treatment (n = 3). Top right, siRNA inhibition of PPARδ prevents HCC cell growth. HuH7 cells (3,000 per well) were seeded in 96-well plate and cultured overnight. The cells were transfected overnight with 5 pmol per well of siRNA-PPARδ or siRNA-control using LipofectAMINE 2000 reagent in 0.5% FBS medium. Following transfection, the cells were treated with GW501516 (10 nmol/L) or vehicle DMSO (1:10,000 dilution) for 24 hours. WST-1 reagent was used to determine cell growth. Columns, mean of three individual experiments; bars, SD. The cells transfected with the PPARδ siRNA exhibited significant reduction of growth either under basal culture conditions or in response to GW501516 treatment. *, P < 0.05, compared with control siRNA cells without GW501516 treatment; **, P < 0.01, compared with control siRNA cells without GW501516 treatment; ***, P < 0.01, compared with control siRNA cells with GW501516 treatment. Bottom, PPARδ protein levels in HuH7 cells with or without PPARδ siRNA transfection. β-Actin was used as the loading control. C, activation of PPARδ by GW501516 enhances COX-2 protein expression. Inhibition of PPARδ by siRNA reduces COX-2 protein level. Top, time course effect of GW501516 on COX-2 protein expression. HuH7 cells were pre-serum starved for 24 hours and treated with GW501516 (10 nmol/L) in serum-free medium for increasing times (2–8 hours). The cell lysates were obtained for SDS-PAGE and Western blot analysis to determine the level of COX-2 protein. The same blot was stripped and reprobed with β-actin antibody as loading control. Increased COX-2 protein was observed after GW501516 treatment for 4 to 8 hours. Middle, dose-dependent effect of GW501516 on COX-2 protein expression. HuH7 cells were incubated with GW501516 (1–50 nmol/L) in serum-free medium for 6 hours, and the whole-cell lysates were obtained for SDS-PAGE and Western blot analysis to determine the level of COX-2. Bottom, siRNA inhibition of PPARδ reduces COX-2 protein level. HuH7 cells (70% confluence) were transfected with PPARδ siRNA or nontargeted control siRNA for 48 hours. Cellular protein (30 μg) was subjected to SDS-PAGE and Western blotting to determine COX-2 protein, with β-actin as loading control. D, activation of PPARδ by GW501516 enhances COX-2 gene promoter activity and PGE2 production. Top, GW501516 treatment enhanced COX-2 gene promoter reporter activity. HuH7 cells (70% confluence in six-well plate) were cotransfected with the COX-2 promoter reporter plasmid (1 μg) plus control siRNA or PPARδ siRNA using LipofectAMINE 2000 reagent. After 48 hours, the cells were incubated with GW501516 (10 nmol/L) or vehicle DMSO (1:10,000 dilution) for 6 hours. Following washing twice with cold PBS, the cells were lysed and the cell lysates were obtained for luciferase activity assay. GW501516 treatment (10 nmol/L) induced a 2.2-fold increase of the COX-2 promoter reporter activity. **, P < 0.01 (n = 3). This effect was completely blocked by siRNA inhibition of PPARδ. ***, P < 0.01, compared with control siRNA plus GW501516 treatment (n = 3). PPARδ siRNA also inhibited the COX-2 promoter activity in cells without GW501516 treatment. *, P < 0.05, compared with control siRNA with vehicle treatment. Bottom left, GW501516 treatment increases PGE2 production. HuH7 cells (80% confluence in six-well plate, with overnight serum starvation) were treated with GW501516 (10–50 nmol/L) or vehicle for 6 hours in serum-free medium. At the end of treatment, the culture medium was collected and cell debris was removed by centrifugation for 25 minutes at 4°C. Medium (100 μL) was used for the measurement of PGE2. Columns, mean of three experiments (0.22 ng/mL for control versus 0.28, 0.32, and 0.45 ng/mL for 10, 20, and 50 nmol/L GW501516, respectively); bars, SD. Bottom right, siRNA inhibition of PPARδ prevents GW501516-induced PGE2 production. HuH7 cells were transfected with PPARδ siRNA or control siRNA for 48 hours followed by vehicle DMSO or GW501516 treatment for 6 hours. **, P < 0.01, compared with control siRNA plus vehicle treatment; ***, P < 0.01, compared with control siRNA plus GW501516 treatment (n = 3).

Close modal

Because COX-2–derived PGE2 signaling is implicated in hepatocarcinogenesis, we postulated that COX-2 and PGE2 signaling might play a role in PPARδ-induced HCC cell growth. To evaluate this hypothesis, we examined the effect of PPARδ activation on COX-2 gene expression in HCC cells. As shown in Fig. 1C, activation of PPARδ by GW501516 enhanced the level of COX-2 protein in HuH7 cells, whereas siRNA inhibition of PPARδ reduced it (a similar effect was also observed in HepG2 cells). This effect is likely mediated through up-regulation of COX-2 gene transcription because GW501516 treatment enhanced the COX-2 promoter reporter activity (Fig. 1D). Accordingly, GW501516 treatment also induced a dose-dependent increase of PGE2 production, which was blocked by siRNA inhibition of PPARδ (Fig. 1D). These findings show a stimulatory effect of PPARδ on COX-2 expression and PGE2 production in human HCC cells. Our data are consistent with a recent study showing that PPARδ agonist GW501516 enhances COX-2 gene reporter activity in HepG2 cells (38). The exact mechanism for PPARδ-induced COX-2 expression is not fully understood at the present time. Because the DRE is not identified in human COX-2 promoter, it is possible that PPARδ may regulate COX-2 gene transcription indirectly through control of other transcriptional factors, although the possibility of PPARδ binding to other unidentified DNA element in the COX-2 gene cannot be entirely excluded.

Given that COX-2–derived PGE2 has been show to promote HCC cell growth through activation of EGFR and Akt (9, 12), we next determined the potential effect of PPARδ activation on EGFR and Akt phosphorylation. As shown in Fig. 2A and B, activation of PPARδ by GW501516 (10 nmol/L) enhanced EGFR and Akt phosphorylation, whereas inhibition of PPARδ decreased their phosphorylation. Furthermore, siRNA inhibition of COX-2 prevented both endogenous and GW501516-induced EGFR and Akt phosphorylation (Fig. 2C). Consistent with these observations, COX-2 siRNA also inhibits cell growth both under basal culture condition and with GW501516 treatment (Fig. 2D). The above results further support the involvement of COX-2 in PPARδ-mediated EGFR/Akt phosphorylation and cell growth.

Figure 2.

Activation of PPARδ by GW501516 induces the phosphorylation of EGFR and Akt through COX-2 in HCC cells. A, GW501516 induces EGFR and Akt phosphorylation. HuH7 cells (80% confluence in six-well plate with overnight serum starvation) were treated with GW501516 (10 nmol/L) for different times (4–8 hours). Whole-cell lysates were collected, and 30 μg protein was subjected to Western blotting for detection of phosphorylated EGFR (p-EGFR), EGFR, phosphorylated Akt (p-Akt), Akt, COX-2, and β-actin. GW501516 treatment increased the level of COX-2 expression as well as the phosphorylation of EGFR and Akt. B, siRNA inhibition of PPARδ reduces phosphorylated EGFR and phosphorylated Akt levels. HuH7 cells were transfected with control or PPARδ siRNA for 48 hours. Cellular protein (30 μg) was subjected to SDS-PAGE and Western blotting. C, siRNA inhibition of COX-2 prevents GW501516-induced EGFR and Akt phosphorylation. HuH7 cells (90% confluence in six-well plate) were transfected with COX-2 siRNA or nontargeted control siRNA using LipofectAMINE 2000 reagent for 48 hours followed by treatment with GW501516 (10 nmol/L) or vehicle DMSO (1:10,000) for 6 hours. Whole-cell lysate (30 μg) was subjected to Western blotting for detection of COX-2, phosphorylated EGFR, EGFR, phosphorylated Akt, Akt, and β-actin. All the experiments were repeated thrice. D, inhibition of COX-2 by siRNA prevents GW501516-induced cell growth. HuH7 cells transfected with control siRNA or COX-2 siRNA for 24 hours were treated with GW501516 or vehicle for additional 24 hours. Cell proliferation assay was carried out using WST-1 assay. *, P < 0.05, compared with control siRNA plus vehicle treatment; **, P < 0.01, compared with control siRNA plus vehicle treatment; ***, P < 0.01, compared with control siRNA plus GW501516 treatment.

Figure 2.

Activation of PPARδ by GW501516 induces the phosphorylation of EGFR and Akt through COX-2 in HCC cells. A, GW501516 induces EGFR and Akt phosphorylation. HuH7 cells (80% confluence in six-well plate with overnight serum starvation) were treated with GW501516 (10 nmol/L) for different times (4–8 hours). Whole-cell lysates were collected, and 30 μg protein was subjected to Western blotting for detection of phosphorylated EGFR (p-EGFR), EGFR, phosphorylated Akt (p-Akt), Akt, COX-2, and β-actin. GW501516 treatment increased the level of COX-2 expression as well as the phosphorylation of EGFR and Akt. B, siRNA inhibition of PPARδ reduces phosphorylated EGFR and phosphorylated Akt levels. HuH7 cells were transfected with control or PPARδ siRNA for 48 hours. Cellular protein (30 μg) was subjected to SDS-PAGE and Western blotting. C, siRNA inhibition of COX-2 prevents GW501516-induced EGFR and Akt phosphorylation. HuH7 cells (90% confluence in six-well plate) were transfected with COX-2 siRNA or nontargeted control siRNA using LipofectAMINE 2000 reagent for 48 hours followed by treatment with GW501516 (10 nmol/L) or vehicle DMSO (1:10,000) for 6 hours. Whole-cell lysate (30 μg) was subjected to Western blotting for detection of COX-2, phosphorylated EGFR, EGFR, phosphorylated Akt, Akt, and β-actin. All the experiments were repeated thrice. D, inhibition of COX-2 by siRNA prevents GW501516-induced cell growth. HuH7 cells transfected with control siRNA or COX-2 siRNA for 24 hours were treated with GW501516 or vehicle for additional 24 hours. Cell proliferation assay was carried out using WST-1 assay. *, P < 0.05, compared with control siRNA plus vehicle treatment; **, P < 0.01, compared with control siRNA plus vehicle treatment; ***, P < 0.01, compared with control siRNA plus GW501516 treatment.

Close modal

cPLA2α is the rate-limiting enzyme that releases arachidonic acid from membrane phospholipids and thus provides substrate for COX enzymes. The cPLA2α- and COX-2-controlled PG synthesis has been implicated in hepatocarcinogenesis (3). Whereas coupled activation of cPLA2α and COX-2 plays an important role for PG production (4547), there is also evidence indicating that PGE2 can further activate cPLA2α in other cancer cells (48). Therefore, we sought to further determine whether PPARδ-induced PGE2 synthesis might affect cPLA2α activation in human HCC cells. As shown in Fig. 3A, treatment of HCC cells with 10 μmol/L PGE2 for 30 minutes increased the phosphorylation of cPLA2α. This effect was completely blocked by pretreatment with the p44/42 MAPK inhibitor (PD98059, 20 μmol/L) or the p38 MAPK (SB203580, 10 μmol/L) but not by the PKC inhibitor (bisindolylmaleimide I, 20 μmol/L) or the PI3K/Akt inhibitor (LY294002, 20 μmol/L; Fig. 3B). These observations suggest the involvement of p38 and p42/44 MAPKs in PGE2-induced cPLA2α phosphorylation in HCC cells.

Figure 3.

GW501516 and PGE2 increase the phosphorylation of cPLA2α in human HCC cells. Evidence for involvement of p42/44 and p38 MAPKs. A, PGE2 treatment enhances cPLA2α phosphorylation. HuH7 cells (80% confluence in six-well plate with overnight serum starvation) were incubated with PGE2 (10 μmol/L) for 5 to 90 minutes. The whole-cell lysates were then collected and precleared with protein A/G agarose beads for 1 hour followed by overnight incubation with cPLA2α antibody at 4°C. The protein-antibody complex was pulled down by incubation with protein A/G agarose beads for 1 hour and subjected to Western blotting for detection of phosphorylated cPLA2α (cPLA2-p) and total cPLA2α (cPLA2-total). A similar effect was also seen in HepG2 cells. B, inhibition of p42/44 and p38 MAPKs prevents PGE2-induced cPLA2α phosphorylation. HuH7 cells (80% confluence in six-well plate) were serum starved overnight and preincubated for 30 minutes with PKC inhibitor [bisindolylmaleimide I (Bisin I), 20 μmol/L], PI3K inhibitor (LY294002, 20 μmol/L), p44/42 inhibitor (PD98059, 20 μmol/L), p38 inhibitor (SB203580, 10 μmol/L), or vehicle DMSO (1:10,000 dilution) followed by coincubation with PGE2 (10 μmol/L) for additional 30 minutes. The whole-cell lysates were collected and subjected to immunoprecipitation and Western blot analysis to determine cPLA2α phosphorylation. The PGE2-induced cPLA2α phosphorylation was blocked by the p44/42 MAPK inhibitor PD98059 or the p38 MAPK inhibitor SB203580 but not by the PKC inhibitor bisindolylmaleimide I or the PI3K inhibitor LY294002. C, GW501516 treatment increases cPLA2α phosphorylation. This effect is inhibited by the MAPK inhibitors. Top, effect of GW501516 on cPLA2α phosphorylation. HuH7 cells (80% confluence in six-well plate) were serum starved overnight and then incubated with GW501516 (10 nmol/L) for 4 to 8 hours. Whole-cell lysates were collected and subjected to immunoprecipitation and Western blot to detect cPLA2α phosphorylation. GW501516 treatment increased cPLA2α phosphorylation, with peak effect at 6 hours. All the experiments were repeated thrice. Bottom, MAPK inhibitors prevent GW501516-induced cPLA2α phosphorylation. HuH7 cells were treated with GW501516 (10 nmol/L) for 6 hours in the presence or absence of PD98059 (20 μmol/L) or SB203580 (10 μmol/L). D, inhibitors of MAPKs and cPLA2α prevent GW501516-induced cell growth. HuH7 cells were cotreated overnight with GW501516 plus vehicle (DMSO, 1:10,000), PD98059 (20 μmol/L), SB203580 (10 μmol/L), or AACOCF3 (25 μmol/L). WST-1 reagents were used for cell proliferation assay. **, P < 0.01, compared with vehicle; ***, P < 0.05, compared with GW501516 treatment (n = 6).

Figure 3.

GW501516 and PGE2 increase the phosphorylation of cPLA2α in human HCC cells. Evidence for involvement of p42/44 and p38 MAPKs. A, PGE2 treatment enhances cPLA2α phosphorylation. HuH7 cells (80% confluence in six-well plate with overnight serum starvation) were incubated with PGE2 (10 μmol/L) for 5 to 90 minutes. The whole-cell lysates were then collected and precleared with protein A/G agarose beads for 1 hour followed by overnight incubation with cPLA2α antibody at 4°C. The protein-antibody complex was pulled down by incubation with protein A/G agarose beads for 1 hour and subjected to Western blotting for detection of phosphorylated cPLA2α (cPLA2-p) and total cPLA2α (cPLA2-total). A similar effect was also seen in HepG2 cells. B, inhibition of p42/44 and p38 MAPKs prevents PGE2-induced cPLA2α phosphorylation. HuH7 cells (80% confluence in six-well plate) were serum starved overnight and preincubated for 30 minutes with PKC inhibitor [bisindolylmaleimide I (Bisin I), 20 μmol/L], PI3K inhibitor (LY294002, 20 μmol/L), p44/42 inhibitor (PD98059, 20 μmol/L), p38 inhibitor (SB203580, 10 μmol/L), or vehicle DMSO (1:10,000 dilution) followed by coincubation with PGE2 (10 μmol/L) for additional 30 minutes. The whole-cell lysates were collected and subjected to immunoprecipitation and Western blot analysis to determine cPLA2α phosphorylation. The PGE2-induced cPLA2α phosphorylation was blocked by the p44/42 MAPK inhibitor PD98059 or the p38 MAPK inhibitor SB203580 but not by the PKC inhibitor bisindolylmaleimide I or the PI3K inhibitor LY294002. C, GW501516 treatment increases cPLA2α phosphorylation. This effect is inhibited by the MAPK inhibitors. Top, effect of GW501516 on cPLA2α phosphorylation. HuH7 cells (80% confluence in six-well plate) were serum starved overnight and then incubated with GW501516 (10 nmol/L) for 4 to 8 hours. Whole-cell lysates were collected and subjected to immunoprecipitation and Western blot to detect cPLA2α phosphorylation. GW501516 treatment increased cPLA2α phosphorylation, with peak effect at 6 hours. All the experiments were repeated thrice. Bottom, MAPK inhibitors prevent GW501516-induced cPLA2α phosphorylation. HuH7 cells were treated with GW501516 (10 nmol/L) for 6 hours in the presence or absence of PD98059 (20 μmol/L) or SB203580 (10 μmol/L). D, inhibitors of MAPKs and cPLA2α prevent GW501516-induced cell growth. HuH7 cells were cotreated overnight with GW501516 plus vehicle (DMSO, 1:10,000), PD98059 (20 μmol/L), SB203580 (10 μmol/L), or AACOCF3 (25 μmol/L). WST-1 reagents were used for cell proliferation assay. **, P < 0.01, compared with vehicle; ***, P < 0.05, compared with GW501516 treatment (n = 6).

Close modal

Consistent with the effect of PPARδ on COX-2 expression and PGE2 synthesis, activation of PPARδ by GW501516 also increased cPLA2α phosphorylation in HCC cells, which was blocked by the p38 and p42/44 MAPK inhibitors (Fig. 3C). The GW501516-induced cPLA2α phosphorylation was observed at 6 hours, whereas the GW501516-induced COX-2 protein increase was observed ∼4 hours after treatment, and the effect peaks at 8 hours (Fig. 1C). The reason for lacking cPLA2α phosphorylation at 8-hour time point is unclear and remains speculative, although the possibility of phosphatase involvement cannot be excluded.

These results presented in the above sections suggest that PPARδ induces COX-2 expression and PGE2 production that in turn enhances cPLA2α phosphorylation, thus further amplifying PGE2 signaling. The involvement of COX-2 and cPLA2α in PPARδ-mediated cell growth is further supported by the observation that inhibiting COX-2 or cPLA2α activation prevents PPARδ agonist-induced HCC cell growth (Figs. 2D and 3D).

Although recent evidence suggests the involvement of cPLA2 in the activation of PPARα and PPARγ in primary and transformed hepatocytes and lung epithelial cells (49, 50), the potential role of cPLA2 in PPARδ activation has not been investigated. In this study, the effect of cPLA2 on PPARδ activation was examined in human HCC cells. To this end, HuH7 cells transfected with the DRE reporter plasmid (a luciferase reporter construct under the control of DRE) were treated with the calcium ionophore A23187. The use of A23187 was based on the fact that calcium is required for the nuclear translocation and activation of cPLA2α. As shown in Fig. 4A, activation of cPLA2α by A23187 (1 μmol/L) significantly increased the PPARδ transcription activity in HuH7 cells (3.5-fold of control; P < 0.01); this effect was inhibited by the cPLA2 inhibitor AACOCF3 (25 μmol/L). In addition, inhibition of cPLA2 by AACOCF3 also decreased basal level of PPARδ transcription activity. These findings suggest the involvement of cPLA2 in PPARδ activation.

Figure 4.

cPLA2α enhances PPARδ activation in human HCC cells. A, effect of cPLA2α on DRE reporter activity. HuH7 cells (90% confluence in six-well plate) transfected with 1 μg of DRE-driven luciferase reporter plasmid were pretreated with AACOCF3 (25 μmol/L) or vehicle DMSO (1:10,000 dilution) for 8 hours. The cells were then exposed to A23187 (1 μmol/L) for 20 minutes, and the cultures were continued for additional 2 hours. At the end of incubation, the cells were washed twice with cold PBS and the cell lysates were obtained to measure luciferase reporter activity. A23187 treatment induced a 3.5-fold increase of the DRE reporter activity. *, P < 0.01, compared with control (n = 3). This effect was completely blocked by pretreatment with AACOCF3. **, P < 0.01, compared with A23187 treatment (n = 3). Treatment with AACOCF3 also significantly inhibited basal level of DRE reporter activity. *, P < 0.01, compared with control (n = 3). B, overexpression of cPLA2α enhances PPARδ binding to its response element (ELISA-based PPARδ transcription factor assay). HuH7 cells were transfected with the cPLA2α expression plasmid or the control vector pMT-2, and the nuclear extracts were obtained. Nuclear extract (50 μg) was added into PPAR response element-coated 96-well plate and incubated overnight to allow protein-DNA binding. After complete wash, anti-PPARδ antibody was added into sample wells and the plates were incubated for 1 hour. For color development, HRP-conjugated secondary antibody and color development solution were added and A655 nm value was measured. Columns, mean of three experiments; bars, SD. *, P < 0.05. C, activation of cPLA2α enhances PPARδ binding to its response element (ELISA-based PPARδ transcription factor assay). HuH7 cells were serum starved overnight and then treated with GW501516 (10 nmol/L, 8 hours) or AACOCF3 (25 μmol/L, 8 hours) before the addition of A23187 (1 μmol/L, 20 minutes). Nuclear extract (50 μg) from these cells was used for PPARδ transcription factor assay as described above. Columns, mean of three individual experiments; bars, SD. *, P < 0.05, compared with vehicle; **, P < 0.01, compared with vehicle; ***, P < 0.01, compared with A23187. D, activation of cPLA2α promotes PPARδ binding to DRE (biotinylated DRE oligonucleotide precipitation assay). HuH7 cells (with overnight serum starvation) were treated with the cPLA2α inhibitors AACOCF3 (25 μmol/L, 8 hours) and the trisubstituted pyrrolidine derivative (2 μmol/L, 20 minutes) or vehicle DMSO (1:10,000 dilution) before the addition of the calcium ionophore A23187 (1 μmol/L, 20 minutes). At the end of treatment, the cell lysates were obtained and processed for biotinylated DRE oligonucleotide precipitation assay. The cell lysates (∼50 μg per sample) were precleared with immobilized streptavidin-agarose beads for 1 hour at 4°C followed by incubation overnight with biotin-labeled DRE oligonucleotide. The protein-DRE complex was pulled down by incubation with immobilized streptavidin beads for 1 hour at 4°C, and the samples were subjected to Western blotting to detect bound PPARδ as described in Materials and Methods. Activation of cPLA2 by A23187 enhanced the binding of PPARδ to DRE. Both cPLA2α inhibitors AACOCF3 and the trisubstituted pyrrolidine derivative inhibited the spontaneous or A23187-indued binding of PPARδ to DRE. The binding specificity was verified by the absence of binding in cold competitive control (biotinylated DRE oligonucleotide admixed with 100-fold of unlabeled DRE oligonucleotide) or with omission of cell lysate. All the experiments were repeated thrice.

Figure 4.

cPLA2α enhances PPARδ activation in human HCC cells. A, effect of cPLA2α on DRE reporter activity. HuH7 cells (90% confluence in six-well plate) transfected with 1 μg of DRE-driven luciferase reporter plasmid were pretreated with AACOCF3 (25 μmol/L) or vehicle DMSO (1:10,000 dilution) for 8 hours. The cells were then exposed to A23187 (1 μmol/L) for 20 minutes, and the cultures were continued for additional 2 hours. At the end of incubation, the cells were washed twice with cold PBS and the cell lysates were obtained to measure luciferase reporter activity. A23187 treatment induced a 3.5-fold increase of the DRE reporter activity. *, P < 0.01, compared with control (n = 3). This effect was completely blocked by pretreatment with AACOCF3. **, P < 0.01, compared with A23187 treatment (n = 3). Treatment with AACOCF3 also significantly inhibited basal level of DRE reporter activity. *, P < 0.01, compared with control (n = 3). B, overexpression of cPLA2α enhances PPARδ binding to its response element (ELISA-based PPARδ transcription factor assay). HuH7 cells were transfected with the cPLA2α expression plasmid or the control vector pMT-2, and the nuclear extracts were obtained. Nuclear extract (50 μg) was added into PPAR response element-coated 96-well plate and incubated overnight to allow protein-DNA binding. After complete wash, anti-PPARδ antibody was added into sample wells and the plates were incubated for 1 hour. For color development, HRP-conjugated secondary antibody and color development solution were added and A655 nm value was measured. Columns, mean of three experiments; bars, SD. *, P < 0.05. C, activation of cPLA2α enhances PPARδ binding to its response element (ELISA-based PPARδ transcription factor assay). HuH7 cells were serum starved overnight and then treated with GW501516 (10 nmol/L, 8 hours) or AACOCF3 (25 μmol/L, 8 hours) before the addition of A23187 (1 μmol/L, 20 minutes). Nuclear extract (50 μg) from these cells was used for PPARδ transcription factor assay as described above. Columns, mean of three individual experiments; bars, SD. *, P < 0.05, compared with vehicle; **, P < 0.01, compared with vehicle; ***, P < 0.01, compared with A23187. D, activation of cPLA2α promotes PPARδ binding to DRE (biotinylated DRE oligonucleotide precipitation assay). HuH7 cells (with overnight serum starvation) were treated with the cPLA2α inhibitors AACOCF3 (25 μmol/L, 8 hours) and the trisubstituted pyrrolidine derivative (2 μmol/L, 20 minutes) or vehicle DMSO (1:10,000 dilution) before the addition of the calcium ionophore A23187 (1 μmol/L, 20 minutes). At the end of treatment, the cell lysates were obtained and processed for biotinylated DRE oligonucleotide precipitation assay. The cell lysates (∼50 μg per sample) were precleared with immobilized streptavidin-agarose beads for 1 hour at 4°C followed by incubation overnight with biotin-labeled DRE oligonucleotide. The protein-DRE complex was pulled down by incubation with immobilized streptavidin beads for 1 hour at 4°C, and the samples were subjected to Western blotting to detect bound PPARδ as described in Materials and Methods. Activation of cPLA2 by A23187 enhanced the binding of PPARδ to DRE. Both cPLA2α inhibitors AACOCF3 and the trisubstituted pyrrolidine derivative inhibited the spontaneous or A23187-indued binding of PPARδ to DRE. The binding specificity was verified by the absence of binding in cold competitive control (biotinylated DRE oligonucleotide admixed with 100-fold of unlabeled DRE oligonucleotide) or with omission of cell lysate. All the experiments were repeated thrice.

Close modal

The role of cPLA2α in PPARδ activation was further examined by assessing the binding of PPARδ to DRE in vitro. For this purpose, two complementary approaches were used, including the biotinylated oligonucleotide precipitation assay to characterize the specific binding phenomenon and the ELISA-based nuclear transcription factor assay to quantitate the amount of PPARδ bound to its response element. As shown in Fig. 4B, overexpression of cPLA2α significantly increased the binding of PPARδ to its response element as determined by the ELISA-based nuclear transcription factor assay. Furthermore, activation of cPLA2 by A23187 also significantly increased the binding of PPARδ to its response element and this effect was completely blocked by the selective cPLA2 inhibitor AACOCF3 (Fig. 4C). These observations further support the role of cPLA2α in PPARδ activation. The fact that AACOCF3 also inhibited PPARδ activation in cells without A23187 treatment (Fig. 4A and C) suggests the presence of endogenous cPLA2α for PPARδ activation.

The effect of cPLA2α on PPARδ binding to its response element was further confirmed by the biotinylated DRE oligonucleotide immunoprecipitation assay. Under this assay system, activation of cPLA2α by A23187 enhanced the binding of PPARδ to DRE and that two structurally unrelated cPLA2α inhibitors, AACOCF3 and the 1,2,4-trisubstituted pyrrolidine derivative, prevented PPARδ-DRE binding (Fig. 4D). The specificity of the assay was confirmed by the complete elimination of binding with the unlabeled DRE oligonucleotides.

Given that PGE2 can phosphorylate and activate cPLA2α and that cPLA2α is implicated in PPARδ activity, we next examined the effect of PGE2 on PPARδ activation in HuH7 cells and evaluated the role of cPLA2 in this process. Figure 5A shows that PGE2 treatment significantly increased the PPARδ transcription activity as determined by transient transfection and reporter activity assays, and this effect was completely blocked by the cPLA2 inhibitor AACOCF3. These observations further support the role of cPLA2 in PGE2-induced PPARδ activation in human HCC cells.

Figure 5.

The effect of arachidonic acid versus PGE2 on PPARδ activation in human HCC cells. A, cPLA2 activity is required for PGE2-induced PPARδ activation in HCC cells. HuH7 cells (90% confluence in six-well plate) were transfected overnight with 1 μg of DRE reporter plasmid using LipofectAMINE Plus reagent. Following transfection, the cells were incubated with AACOCF3 (25 μmol/L) for 4 hours followed by treatment with PGE2 (10 μmol/L) for additional 4 hours. The cells were then washed twice with cold PBS, and the cell lysates were obtained for luciferase activity assay. PGE2 treatment induced a 2.5-fold increase of the DRE reporter activity, which was completely blocked by pretreatment with AACOCF3. *, P < 0.01, compared with vehicle control; **, P < 0.01, compared with PGE2 treatment (n = 3). B, direct effect of arachidonic acid on PPARδ binding to DRE. Top, addition of arachidonic acid (AA) to isolated nuclear extract induces PPARδ binding to DRE in vitro. The nuclear extract isolated from serum-starved HuH7 cells was precleared for 1 hour by incubation with immobilized streptavidin-agarose beads. Precleared nuclear extract (10 μg) was incubated with PGE2 (10 μmol/L), GW501516 (10 nmol/L), arachidonic acid (10 μmol/L), or vehicle DMSO (1:10,000 dilution) for 30 minutes at 4°C followed by overnight incubation with biotinylated DRE oligonucleotide at 4°C. The PPARδ-DRE complex was pulled down with streptavidin-agarose beads and subjected to Western blotting to detect bound PPARδ. Addition of arachidonic acid to the nuclear extract promotes PPARδ binding to DRE, whereas PGE2 has no effect in the same system. The level of arachidonic acid–induced PPARδ binding is comparable with that induced by GW501516. The binding specificity was verified by the absence of binding in cold competitive control (biotinylated DRE oligonucleotide admixed with 100-fold of unlabeled DRE oligonucleotide) or with omission of cell lysate. Bottom, inhibition of COX has no effect on A23187-induced PPARδ binding to DRE in human HCC cells. Serum-starved HuH7 cells were incubated overnight with the COX inhibitors indomethacin (30 μmol/L) or NS-398 (30 μmol/L) or vehicle DMSO before incubation with A23187 (1 μmol/L, 20 minutes). At the end of treatment, the whole-cell lysates were obtained as described in Materials and Methods. Cell lysate (50 μg) was precleared and then incubated overnight with the biotinylated DRE. The PPARδ-DRE complex was pulled down and subjected to Western blotting for detection of bound PPARδ. Indomethacin and NS-398 had no influence on A23187-induced PPARδ binding to DRE. All the experiments were repeated thrice. C, inhibition of COX has no effect on cPLA2α overexpression-induced DRE reporter activity. HuH7 cells were cotransfected with DRE and cPLA2α expression plasmid or control vector pMT-2 for 3 hours followed by treatment with indomethacin (30 μmol/L) or vehicle DMSO overnight. Although overexpression of cPLA2α enhances the DRE reporter activity (**, P < 0.01, compared with control vector), this effect is not inhibited by indomethacin (P > 0.05). Columns, mean of three separate experiments; bars, SD. D, the PGE2-induced DRE reporter activity is blocked by the p38 MAPK inhibitor (SB203580) and the p42/44 MAPK inhibitor (PD98059) but not by the PI3K/Akt inhibitor (LY294002). HuH7 cells were transfected with DRE reporter plasmid for 3 hours followed by treatment with indomethacin (30 μmol/L) or PGE2 (10 μmol/L) plus SB203580 (10 μmol/L), PD98059 (25 μmol/L), or LY294002 (20 μmol/L) overnight. *, P < 0.05; **, P < 0.01, compared with vehicle control; ***, P < 0.01, compared with PGE2 treatment (n = 3).

Figure 5.

The effect of arachidonic acid versus PGE2 on PPARδ activation in human HCC cells. A, cPLA2 activity is required for PGE2-induced PPARδ activation in HCC cells. HuH7 cells (90% confluence in six-well plate) were transfected overnight with 1 μg of DRE reporter plasmid using LipofectAMINE Plus reagent. Following transfection, the cells were incubated with AACOCF3 (25 μmol/L) for 4 hours followed by treatment with PGE2 (10 μmol/L) for additional 4 hours. The cells were then washed twice with cold PBS, and the cell lysates were obtained for luciferase activity assay. PGE2 treatment induced a 2.5-fold increase of the DRE reporter activity, which was completely blocked by pretreatment with AACOCF3. *, P < 0.01, compared with vehicle control; **, P < 0.01, compared with PGE2 treatment (n = 3). B, direct effect of arachidonic acid on PPARδ binding to DRE. Top, addition of arachidonic acid (AA) to isolated nuclear extract induces PPARδ binding to DRE in vitro. The nuclear extract isolated from serum-starved HuH7 cells was precleared for 1 hour by incubation with immobilized streptavidin-agarose beads. Precleared nuclear extract (10 μg) was incubated with PGE2 (10 μmol/L), GW501516 (10 nmol/L), arachidonic acid (10 μmol/L), or vehicle DMSO (1:10,000 dilution) for 30 minutes at 4°C followed by overnight incubation with biotinylated DRE oligonucleotide at 4°C. The PPARδ-DRE complex was pulled down with streptavidin-agarose beads and subjected to Western blotting to detect bound PPARδ. Addition of arachidonic acid to the nuclear extract promotes PPARδ binding to DRE, whereas PGE2 has no effect in the same system. The level of arachidonic acid–induced PPARδ binding is comparable with that induced by GW501516. The binding specificity was verified by the absence of binding in cold competitive control (biotinylated DRE oligonucleotide admixed with 100-fold of unlabeled DRE oligonucleotide) or with omission of cell lysate. Bottom, inhibition of COX has no effect on A23187-induced PPARδ binding to DRE in human HCC cells. Serum-starved HuH7 cells were incubated overnight with the COX inhibitors indomethacin (30 μmol/L) or NS-398 (30 μmol/L) or vehicle DMSO before incubation with A23187 (1 μmol/L, 20 minutes). At the end of treatment, the whole-cell lysates were obtained as described in Materials and Methods. Cell lysate (50 μg) was precleared and then incubated overnight with the biotinylated DRE. The PPARδ-DRE complex was pulled down and subjected to Western blotting for detection of bound PPARδ. Indomethacin and NS-398 had no influence on A23187-induced PPARδ binding to DRE. All the experiments were repeated thrice. C, inhibition of COX has no effect on cPLA2α overexpression-induced DRE reporter activity. HuH7 cells were cotransfected with DRE and cPLA2α expression plasmid or control vector pMT-2 for 3 hours followed by treatment with indomethacin (30 μmol/L) or vehicle DMSO overnight. Although overexpression of cPLA2α enhances the DRE reporter activity (**, P < 0.01, compared with control vector), this effect is not inhibited by indomethacin (P > 0.05). Columns, mean of three separate experiments; bars, SD. D, the PGE2-induced DRE reporter activity is blocked by the p38 MAPK inhibitor (SB203580) and the p42/44 MAPK inhibitor (PD98059) but not by the PI3K/Akt inhibitor (LY294002). HuH7 cells were transfected with DRE reporter plasmid for 3 hours followed by treatment with indomethacin (30 μmol/L) or PGE2 (10 μmol/L) plus SB203580 (10 μmol/L), PD98059 (25 μmol/L), or LY294002 (20 μmol/L) overnight. *, P < 0.05; **, P < 0.01, compared with vehicle control; ***, P < 0.01, compared with PGE2 treatment (n = 3).

Close modal

To further delineate the effect of arachidonic acid and PGE2 on PPARδ activation, a cell-free system was used, in which the nuclear extracts from HuH7 cells were incubated with PGE2 or arachidonic acid in the presence of biotinylated DRE oligonucleotide to determine the binding of PPARδ to DRE in vitro. Addition of arachidonic acid to the nuclear extract induced the binding of PPARδ to DRE, which is comparable with the effect induced by the synthetic PPARδ ligand GW501516 (Fig. 5B). In contrast, PGE2 failed to induce PPARδ binding when directly added to the nuclear extract (Fig. 5B). These findings suggest that PGE2 lacks the ability to directly activate PPARδ, although arachidonic acid itself can bind PPARδ and alter PPARδ transcription activity. The latter assertion is further supported by the observation that the COX-2 inhibitors indomethacin and NS-398 had no apparent influence on A23187-induced PPARδ binding to DRE (Fig. 5B). Thus, given that PGE2 activates PPARδ only in intact cell, its effect is most likely mediated through cPLA2α phosphorylation-induced arachidonic acid release rather than direct PPARδ binding. In agreement with this assertion, forced expression of cPLA2α enhanced DRE reporter activity and this effect was not blocked by the COX inhibitor indomethacin (Fig. 5C), although indomethacin alone decreased DRE reporter activity (Fig. 5D). In addition, the PGE2-induced PPARδ transcriptional activity was inhibited by the p44/42 MAPK inhibitor (PD98059) and the p38 MAPK (SB203580; Fig. 5D). These findings are consistent with the effect of these protein kinase inhibitors on PGE2-induced cPLA2α phosphorylation (Fig. 3B) and further support the role of cPLA2α in PGE2-induced PPARδ activation. In contrast to the reported involvement of the PI3K/Akt pathway in PGE2-induced PPARδ activation in colorectal adenoma cells (40), in our system blocking PI3K/Akt by LY294002 did not affect PGE2-induced PPARδ activation in HCC cells. Taken together, our results suggest that the MAPK-mediated cPLA2α phosphorylation is an important mechanism for PGE2-induced PPARδ activation in HCC cells.

In summary, this study shows an important role of PPARδ in human HCC cell growth. The most novel mechanistic aspect of this study is the identification of cPLA2α-controlled arachidonic acid metabolism for endogenous PPARδ activation in the nucleus. The importance of cPLA2α in PPAR activation can be explained by its unique characteristic of nuclear localization mediated by its NH2-terminal Ca2+-dependent lipid binding domain (CaLB or C2 domain). Our findings provide the first evidence for the activation of PPARδ by cPLA2α in human HCC cells. The observations that PPARδ enhances COX-2 gene expression and that the COX-2–derived PGE2 further activates PPARδ through phosphorylation of cPLA2α depict a novel cross-talk between PPARδ and PG signaling pathways that coordinately regulate HCC cell growth (Fig. 6). It is conceivable that disruption of this feed-forward loop may represent an important future therapeutic strategy for the chemoprevention and treatment of human HCC.

Figure 6.

Mechanisms for PPARδ-mediated human HCC cell growth. PPARδ promotes human HCC cell growth through induction of COX-2 expression and PGE2 synthesis. The produced PGE2 phosphorylates and activates cPLA2α, thus releasing arachidonic acid for further PPARδ activation and PGE2 synthesis via COX-2. This positive feed-forward loop between PPARδ and PG pathways likely plays an important role in the regulation of human HCC cell growth.

Figure 6.

Mechanisms for PPARδ-mediated human HCC cell growth. PPARδ promotes human HCC cell growth through induction of COX-2 expression and PGE2 synthesis. The produced PGE2 phosphorylates and activates cPLA2α, thus releasing arachidonic acid for further PPARδ activation and PGE2 synthesis via COX-2. This positive feed-forward loop between PPARδ and PG pathways likely plays an important role in the regulation of human HCC cell growth.

Close modal

Grant support: NIH R01 grants CA102325 and CA106280 (T. Wu).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank Drs. Vogelstein and Kinzler (Johns Hopkins Oncology Center, Baltimore, MD) for providing the DRE reporter plasmid and Drs. J.D. Clark and J.L. Knopf (Genetics Institute, Boston, MA) for the cPLA2α expression plasmid.

1
Parkin DM, Bray F, Ferlay J, Pisani P. Estimating the world cancer burden: Globocan 2000.
Int J Cancer
2001
;
94
:
153
–6.
2
Coleman WB. Mechanisms of human hepatocarcinogenesis.
Curr Mol Med
2003
;
3
:
573
–88.
3
Wu T. Cyclooxygenase-2 in hepatocellular carcinoma.
Cancer Treat Rev
2006
;
32
:
28
–44.
4
Kondo M, Yamamoto H, Nagano H, et al. Increased expression of COX-2 in nontumor liver tissue is associated with shorter disease-free survival in patients with hepatocellular carcinoma.
Clin Cancer Res
1999
;
5
:
4005
–12.
5
Koga H, Sakisaka S, Ohishi M, et al. Expression of cyclooxygenase-2 in human hepatocellular carcinoma: relevance to tumor dedifferentiation.
Hepatology
1999
;
29
:
688
–96.
6
Shiota G, Okubo M, Noumi T, et al. Cyclooxygenase-2 expression in hepatocellular carcinoma.
Hepatogastroenterology
1999
;
46
:
407
–12.
7
Bae SH, Jung ES, Park YM, et al. Expression of cyclooxygenase-2 (COX-2) in hepatocellular carcinoma and growth inhibition of hepatoma cell lines by a COX-2 inhibitor, NS-398.
Clin Cancer Res
2001
;
7
:
1410
–8.
8
Rahman MA, Dhar DK, Yamaguchi E, et al. Coexpression of inducible nitric oxide synthase and COX-2 in hepatocellular carcinoma and surrounding liver: possible involvement of COX-2 in the angiogenesis of hepatitis C virus-positive cases.
Clin Cancer Res
2001
;
7
:
1325
–32.
9
Leng J, Han C, Demetris AJ, Michalopoulos GK, Wu T. Cyclooxygenase-2 promotes hepatocellular carcinoma cell growth through Akt activation: evidence for Akt inhibition in celecoxib-induced apoptosis.
Hepatology
2003
;
38
:
756
–68.
10
Cheng AS, Chan HL, To KF, et al. Cyclooxygenase-2 pathway correlates with vascular endothelial growth factor expression and tumor angiogenesis in hepatitis B virus-associated hepatocellular carcinoma.
Int J Oncol
2004
;
24
:
853
–60.
11
Ikeda T, Tozuka S, Hasumura Y, Takeuchi J. Prostaglandin-E-producing hepatocellular carcinoma with hypercalcemia.
Cancer
1988
;
61
:
1813
–4.
12
Han C, Michalopoulos GK, Wu T. Prostaglandin E(2) receptor EP(1) transactivates EGFR/MET receptor tyrosine kinases and enhances invasiveness in human hepatocellular carcinoma cells.
J Cell Physiol
2006
;
207
:
261
–70.
13
Balsinde J, Balboa MA, Insel PA, Dennis EA. Regulation and inhibition of phospholipase A2.
Annu Rev Pharmacol Toxicol
1999
;
39
:
175
–89.
14
Murakami M, Kudo I. Phospholipase A2.
J Biochem (Tokyo)
2002
;
131
:
285
–92.
15
Gupta RA, Dubois RN. Colorectal cancer prevention and treatment by inhibition of cyclooxygenase-2.
Nat Rev Cancer
2001
;
1
:
11
–21.
16
Smith WL, Langenbach R. Why there are two cyclooxygenase isozymes.
J Clin Invest
2001
;
107
:
1491
–5.
17
Kliewer SA, Forman BM, Blumberg B, et al. Differential expression and activation of a family of murine peroxisome proliferator-activated receptors.
Proc Natl Acad Sci U S A
1994
;
91
:
7355
–9.
18
Braissant O, Foufelle F, Scotto C, Dauca M, Wahli W. Differential expression of peroxisome proliferator-activated receptors (PPARs): tissue distribution of PPAR-α, -β, and -γ in the adult rat.
Endocrinology
1996
;
137
:
354
–66.
19
Reddy JK, Hashimoto T. Peroxisomal β-oxidation and peroxisome proliferator-activated receptor α: an adaptive metabolic system.
Annu Rev Nutr
2001
;
21
:
193
–230.
20
Moore KJ, Rosen ED, Fitzgerald ML, et al. The role of PPAR-γ in macrophage differentiation and cholesterol uptake.
Nat Med
2001
;
7
:
41
–7.
21
Fajas L, Debril MB, Auwerx J. Peroxisome proliferator-activated receptor-γ: from adipogenesis to carcinogenesis.
J Mol Endocrinol
2001
;
27
:
1
–9.
22
Oliver WR, Jr., Shenk JL, Snaith MR, et al. A selective peroxisome proliferator-activated receptor δ agonist promotes reverse cholesterol transport.
Proc Natl Acad Sci U S A
2001
;
98
:
5306
–11.
23
Evans RM, Barish GD, Wang YX. PPARs and the complex journey to obesity.
Nat Med
2004
;
10
:
355
–61.
24
Wang YX, Lee CH, Tiep S, et al. Peroxisome-proliferator-activated receptor δ activates fat metabolism to prevent obesity.
Cell
2003
;
113
:
159
–70.
25
Chawla A, Lee CH, Barak Y, et al. PPARδ is a very low-density lipoprotein sensor in macrophages.
Proc Natl Acad Sci U S A
2003
;
100
:
1268
–73.
26
Luquet S, Lopez-Soriano J, Holst D, et al. Peroxisome proliferator-activated receptor δ controls muscle development and oxidative capability.
FASEB J
2003
;
17
:
2299
–301.
27
Lee CH, Chawla A, Urbiztondo N, et al. Transcriptional repression of atherogenic inflammation: modulation by PPARδ.
Science
2003
;
302
:
453
–7.
28
Barak Y, Liao D, He W, et al. Effects of peroxisome proliferator-activated receptor δ on placentation, adiposity, and colorectal cancer.
Proc Natl Acad Sci U S A
2002
;
99
:
303
–8.
29
Burdick AD, Kim DJ, Peraza MA, Gonzalez FJ, Peters JM. The role of peroxisome proliferator-activated receptor-β/δ in epithelial cell growth and differentiation.
Cell Signal
2006
;
18
:
9
–20.
30
Lim H, Gupta RA, Ma WG, et al. Cyclo-oxygenase-2-derived prostacyclin mediates embryo implantation in the mouse via PPARδ.
Genes Dev
1999
;
13
:
1561
–74.
31
Kopelovich L, Fay JR, Glazer RI, Crowell JA. Peroxisome proliferator-activated receptor modulators as potential chemopreventive agents.
Mol Cancer Ther
2002
;
1
:
357
–63.
32
Luquet S, Gaudel C, Holst D, et al. Roles of PPARδ in lipid absorption and metabolism: a new target for the treatment of type 2 diabetes.
Biochim Biophys Acta
2005
;
1740
:
313
–7.
33
He TC, Chan TA, Vogelstein B, Kinzler KW. PPARδ is an APC-regulated target of nonsteroidal anti-inflammatory drugs.
Cell
1999
;
99
:
335
–45.
34
Gupta RA, Tan J, Krause WF, et al. Prostacyclin-mediated activation of peroxisome proliferator-activated receptor δ in colorectal cancer.
Proc Natl Acad Sci U S A
2000
;
97
:
13275
–80.
35
Gupta RA, Wang D, Katkuri S, Wang H, Dey SK, DuBois RN. Activation of nuclear hormone receptor peroxisome proliferator-activated receptor-δ accelerates intestinal adenoma growth.
Nat Med
2004
;
10
:
245
–7.
36
Park BH, Vogelstein B, Kinzler KW. Genetic disruption of PPARδ decreases the tumorigenicity of human colon cancer cells.
Proc Natl Acad Sci U S A
2001
;
98
:
2598
–603.
37
Stephen RL, Gustafsson MC, Jarvis M, et al. Activation of peroxisome proliferator-activated receptor δ stimulates the proliferation of human breast and prostate cancer cell lines.
Cancer Res
2004
;
64
:
3162
–70.
38
Glinghammar B, Skogsberg J, Hamsten A, Ehrenborg E. PPARδ activation induces COX-2 gene expression and cell proliferation in human hepatocellular carcinoma cells.
Biochem Biophys Res Commun
2003
;
308
:
361
–8.
39
Shureiqi I, Jiang W, Zuo X, et al. The 15-lipoxygenase-1 product 13-S-hydroxyoctadecadienoic acid down-regulates PPAR-δ to induce apoptosis in colorectal cancer cells.
Proc Natl Acad Sci U S A
2003
;
100
:
9968
–73.
40
Wang D, Wang H, Shi Q, et al. Prostaglandin E(2) promotes colorectal adenoma growth via transactivation of the nuclear peroxisome proliferator-activated receptor δ.
Cancer Cell
2004
;
6
:
285
–95.
41
Harman FS, Nicol CJ, Marin HE, Ward JM, Gonzalez FJ, Peters JM. Peroxisome proliferator-activated receptor-δ attenuates colon carcinogenesis.
Nat Med
2004
;
10
:
481
–3.
42
Han C, Leng J, Demetris AJ, Wu T. Cyclooxygenase-2 promotes human cholangiocarcinoma growth: evidence for cyclooxygenase-2-independent mechanism in celecoxib-mediated induction of p21waf1/cip1 and p27kip1 and cell cycle arrest.
Cancer Res
2004
;
64
:
1369
–76.
43
Hata A, Seoane J, Lagna G, Montalvo E, Hemmati-Brivanlou A, Massague J. OAZ uses distinct DNA- and protein-binding zinc fingers in separate BMP-Smad and Olf signaling pathways.
Cell
2000
;
100
:
229
–40.
44
Sznaidman ML, Haffner CD, Maloney PR, et al. Novel selective small molecule agonists for peroxisome proliferator-activated receptor δ (PPARδ)—synthesis and biological activity.
Bioorg Med Chem Lett
2003
;
13
:
1517
–21.
45
Murakami M, Naraba H, Tanioka T, et al. Regulation of prostaglandin E2 biosynthesis by inducible membrane-associated prostaglandin E2 synthase that acts in concert with cyclooxygenase-2.
J Biol Chem
2000
;
275
:
32783
–92.
46
Murakami M, Kambe T, Shimbara S, Kudo I. Functional coupling between various phospholipase A2s and cyclooxygenases in immediate and delayed prostanoid biosynthetic pathways.
J Biol Chem
1999
;
274
:
3103
–15.
47
Fitzpatrick FA, Soberman R. Regulated formation of eicosanoids.
J Clin Invest
2001
;
107
:
1347
–51.
48
Hughes-Fulford M, Tjandrawinata RR, Li CF, Sayyah S. Arachidonic acid, an ω-6 fatty acid, induces cytoplasmic phospholipase A2 in prostate carcinoma cells.
Carcinogenesis
2005
;
26
:
1520
–6.
49
Han C, Demetris AJ, Michalopoulos G, Shelhamer JH, Wu T. 85-kDa cPLA(2) plays a critical role in PPAR-mediated gene transcription in human hepatoma cells.
Am J Physiol Gastrointest Liver Physiol
2002
;
282
:
G586
–97.
50
Pawliczak R, Han C, Huang XL, Demetris AJ, Shelhamer JH, Wu T. 85-kDa cytosolic phospholipase A2 mediates peroxisome proliferator-activated receptor γ activation in human lung epithelial cells.
J Biol Chem
2002
;
277
:
33153
–63.