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Abstract

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. References
  7. Supporting Information

Peroxisome proliferator-activated receptor γ (PPARγ) has been implicated in the differentiation and growth inhibition of cancer cells. We examined the effects of PPARγ activation by troglitazone on hepatocellular carcinoma (HCC) cell growth, proliferation, and apoptosis in vitro and in vivo. We also studied relationships between PPARγ activation and cyclooxygenase-2 (COX-2) expression. Human HCC cell lines Huh7 and Hep3B were cultured in the presence or absence of troglitazone. Cell growth was determined via WST-1 assay, proliferation by cell cycle analysis and proliferating cell nuclear antigen (PCNA) Western blotting, and apoptosis by flow cytometry and TUNEL. Tumor growth after subcutaneous implantation of Huh7 cells in nude mice was monitored, and the effects of treatment with troglitazone were determined. In resected HCCs, PPARγ expression was less compared with the histologically normal surrounding liver. In cultures of Hep3B and Huh7 cells, basal expression of PPARγ was relatively low, but troglitazone caused dose-dependent induction of PPARγ expression. Cell cycle analysis revealed a decreased proportion of cells in S phase, with arrest at G0/G1. Concomitant downregulation of PCNA and an increase in TUNEL staining, cells were consistent with decreased proliferation and induction of apoptosis by troglitazaone. Troglitazone-mediated PPARγ activation also suppressed COX-2 expression and induced p27 in HCC cells. Administration of troglitazone to Huh7 tumor-bearing mice significantly reduced tumor growth and caused tumor regression. In conclusion, collectively, these results indicate that PPARγ could be a regulator of cell survival and growth in HCC. PPARγ therefore represents a putative molecular target for chemopreventive therapy or inhibition of liver cancer growth.. (HEPATOLOGY 2006;43:134–143.)

On a global scale, hepatocellular carcinoma (HCC) remains the third leading cause of cancer death due to both high incidence and poor survival. Although prevention of hepatitis B and C virus infections (primary prevention), cleaner water supplies, and more adequate antiviral treatment of advanced-stage hepatitis B and C (secondary prevention) can reduce the risk of HCC, there has been growing interest in the chemoprevention of cancer among high-risk individuals. At least some HCCs develop in a stepwise fashion from dysplastic lesions, through adenomatous proliferation to carcinoma in an analogous manner to colorectal carcinoma. The liver is therefore an attractive target for the identification and use of chemopreventive agents. In addition to DNA mutations, the biological processes relevant to hepatocarcinogenesis include disordered hepatocyte proliferation and operation of cell death pathways, particularly by apoptosis; these may be targets for chemopreventive agents.

Peroxisome proliferator-activated receptors (PPARs) are ligand-activated nuclear receptors that mediate transcriptional regulation of genes involved in the oxidation, transport, and storage of lipids.1 In addition to this pivotal role in lipid metabolism, PPARs influence such biological processes as inflammation, cell survival, differentiation, cell proliferation, and tumorigenesis.2 Among the three PPAR isoforms, PPARγ is of particular interest in this respect. PPARγ encodes two isoforms, PPARγ1 and PPARγ2; the human liver expresses mostly PPARγ1.3In vitro, ligands that activate PPARγ inhibit growth and induce differentiation in breast, prostate, colon, gastric, and liver cancer cells.3, 4 It is less clear whether these findings in cell lines can be translated to in vivo biological effects. Thus, there are conflicting data on whether PPARγ ligands promote or oppose tumorigenesis when applied in different animal models of colon cancer.5–9 The reasons for these discrepant findings is unclear, but it is salient that colorectal cancer and HCC (especially associated with chronic hepatitis C) share the similarity of multistage tumor development. Furthermore, even though PPARγ agonists inhibit cell growth of some other cancers, the molecular mechanisms of their action is not clear.

In this study, we first determined PPARγ expression in clinical samples of human HCC and nontumorous surrounding liver. We used a prototypic thiazolidinedione, troglitazone, to examine the effects of PPARγ activation on PPARγ expression in two human HCC cell lines: Hep3B and Huh7. Having noted PPARγ activation, we then studied the effects on expression of cyclooxygenase-2 (COX-2), another putative epigenetic pathway of tumorigenesis. Troglitazone exerted impressive biological effects in blocking the growth of HCC cells in vitro, as well as their growth as tumors in a nude mouse model. We therefore explored the locus of biological actions of PPARγ activation in HCC cells in terms of cell proliferation and apoptosis.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. References
  7. Supporting Information

Patients and Tissues.

Surgically excised HCC tissues and surrounding nontumor liver tissues were obtained from 20 HCC patients (mean age, 63.1 years; 13 males and 8 females). Written consent forms were obtained before tissue collection, and the study was approved by the Human Ethics Committees of the University of Magdeburg, Germany and Western Sydney Area Health Service, Australia. Among 20 HCC patients, 2 were infected with hepatitis C virus, 3 with hepatitis B virus, and in 15 there was no known cause of HCC; 12 HCC cases were concurrent with cirrhosis and 8 with chronic hepatitis. Each HCC was graded histologically into well differentiated (n = 8), moderately differentiated (n = 8), or poorly differentiated (n = 4) according to Liver Cancer Study Group criteria.10 Tissue aliquots were snap-frozen in liquid nitrogen for later messenger RNA (mRNA) and protein analysis. Other tissue aliquots were fixed in 4% paraformaldehyde and processed for paraffin-embedding.

Cell Culture.

Liver cancer cell lines were obtained from the American Tissue Culture Collection (Manassas, VA) and maintained in Dulbecco's modified Eagle medium with 10% fetal bovine serum (Trace, Victoria, Australia) and penicillin (200 U/mL), and were maintained at 37°C in a humidified atmosphere with 5% CO2. After 24 hours, cultures of approximately 60% cell confluence were treated with various concentrations of troglitazone (Calbiochem, Darmstadt, Germany) and incubated for a further 48 hours. All tissue culture media and media supplements were purchased from Invitrogen (Gaithersberg, MD).

Cell Cycle Analysis.

Cells were seeded (1 × 106 cells/well) in 6-well plates, treated with or without troglitazone for 48 hours and then trypsinized, washed in phosphate-buffered saline and fixed in ice-cold 70% ethanol–phosphate-buffered saline. DNA was labeled with propidium iodide. Cells were sorted by FACScan analysis, and cell cycle profiles were determined using ModFitLT software (Becton Dickinson, San Diego, CA).

Cell Growth Assay (WST-1).

To assay growth rates, cells (1 × 106/well) were seeded in 6-well plates with or without troglitazone. After drug treatment, WST-1 reagent (10:1) (Roche Diagnostics, Sydney, Australia) was added before incubation at 37°C for 2 hours. Absorbance was measured with an ELISA reader (Opsys MR; Dynex Technologies, Worthing West Sussex, UK) at 450 nm. Cell viability was expressed as a percentage of absorbance in treated wells relative to that of untreated (control) wells.

Apoptosis Assay.

Apoptosis was analyzed via two methods. First, sub-G1 DNA analysis was conducted by growing cells in 6-well plates (1 × 106cells/well) with troglitazone for 48 hours. Free nuclei stained with propidium iodide were obtained from hypotonic lysis of cells in a buffer containing sodium citrate (0.1%), Triton X (0.1%), and propidium iodide (50 g/mL) and analyzed on a Becton Dickinson FACScan. Cells undergoing apoptosis were detected as sub-G1 population due to loss of fragmented DNA. Second, Terminal deoxynucleotidyl transferase–mediated nick end labeling (TUNEL) was performed following the manufacturer's protocol (ssDNA apoptosis TUNEL kit, Roche Diagnostics, Indianapolis, IN).

Complementary DNA Synthesis and Reverse-Transcriptase Polymerase Chain Reaction.

Total RNA was extracted by using RNA Trizol reagent (Invitrogen, Carlsbad, CA) according to the manufacturer's protocol. Five micrograms of total RNA was reverse-transcribed into complementary DNA. GAPDH served as an internal control for total complementary DNA content. RNA levels of PPARγ and COX-2 were quantified by real-time reverse-transcriptase polymerase chain reaction (RT-PCR) using SYBRGreen Master Mix (Applied Biosystems, Foster City, CA). Samples were amplified using the ABI Prism 7700 Sequence Detection System (Applied Biosystems).

Western Blotting.

Total protein was extracted in Tris-HCl (pH 7.4) buffer containing 1% Triton X-100 and a protease inhibitor cocktail (Roche Diagnostics). Protein concentration was determined by a method described by Bradford (DC protein assay, Bio-Rad, Hercules, CA). Twenty-five micrograms of protein were separated via 12% SDS-PAGE and transferred onto an equilibrated polyvinylidene difluoride membrane (Amersham Biosciences, Buckinghamshire, UK) via electroblotting. Membranes were blocked using 5% skim milk, then incubated with primary antibodies (Santa Cruz Biotechnology, Santa Cruz, CA) against PPARγ (1:1,000), proliferating cell nuclear antigen (PCNA) (1:1,000), p53 (1:2,000), p21 (1:1,000), p27 (1:1,000), or GAPDH (1:3,000) overnight at 4°C. After incubation with the secondary antibody, proteins were detected by enhanced chemiluminescence (ECL, Amersham Biosciences), and bands quantified by scanning densitometry.

Tumor Xenografts in Balb/c Nude Mice.

Care of animals and all experimental procedures were approved by the Animal Care and Ethics Committee of the Western Sydney Area Health Service in compliance with the highest international standards of humane care in animal experimentation. Four-week-old male athymic nude mice (Balb/c nu/nu) were obtained from ARC (Perth, Australia). All mice had free access to sterilized food and autoclaved water. Experiments were started after 1 week of acclimatization. On day 1, a suspension of Huh7 cells (1 × 106 cells in 0.1 mL phosphate-buffered saline) was injected subcutaneously into the dorsal flank of each mouse. Animals were randomized to receive either troglitazone in chow (200 ppm) (n = 5) or vehicle (n = 4) on day 1, and experiments continued for 25 days. In separate studies designed to assess possible tumor regression, mice were randomly allocated to be treated with troglitazone (200 ppm) or receive vehicle; these treatments were started when tumors reached 2-4 mm diameter (day 12 after Huh7 injection) or 8-10 mm diameter (usually day 15 after Huh7 injection). Tumor diameter was measured with a caliber ruler every other day. Tumor volume (mm3) was estimated by measuring the longest and shortest diameter of the tumor and calculating volume as: volume = (shortest diameter)2 × (longest diameter) × 0.5. Tumor weight was measured on the day of harvest, after excision of the tumor from the euthanized mouse. A portion of the tumor tissue was fixed in 10% formalin for subsequent histological examination, and the remaining tissue was flash-frozen in liquid nitrogen and stored at −70°C for molecular studies.

Statistical Analysis.

Data are expressed as the mean ± SD. For comparisons between groups, we used ANOVA after Bonferroni's correction for multiple comparisons. The difference in tumor volume between the animals fed with troglitazone and vehicle control was determined via Student t test. The correlation between PPARγ and COX-2 expression was analyzed by Pearson correlation coefficient. A P value of less than .05 was considered statistically significant.

Results

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. References
  7. Supporting Information

Expression of PPARγ in Hepatocellular Carcinomas, and in HCC Cell Lines.

As determined by real-time RT-PCR, levels of PPARγ mRNA were detectable but significantly lower in tumor samples than in the surrounding liver tissue (Fig. 1A). Expression levels of PPARγ mRNA appeared related to the state of histological differentiation; poorly differentiated tumors exhibited lower levels of PPARγ mRNA than well-differentiated tumors (Fig. 1B). The results of Western immunoblot analysis confirmed that PPARγ protein was expressed at lower levels in the HCCs compared with surrounding nontumerous liver tissue (Fig. 1C). There was no significant association between PPARγ expression and other clinical-pathological parameters of HCC, including hepatitis B or C infection or presence of cirrhosis.

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Figure 1. PPARγ expression in human hepatocellular carcinomas, and in surrounding liver. (A) Relative levels of PPARγ mRNA (normalized to GAPDH) in primary liver tumors and in surrounding nontumorous liver. Bar graphs indicate group values (n = 20) as the mean ± SD. *P < .05, tumor compared with nontumorous liver. (B) Relative levels of PPARγ mRNA (normalized to GAPDH) in liver tumors according to state of histological differentiation. #P < .05, poorly differentiated (n = 4) compared with well-differentiated tumors (n = 8). The remaining 8 were moderately differentiated tumors. (C) Western blot analysis showing PPARγ protein expression (normalized to GAPDH) in 20 primary liver tumors and in corresponding nontumorous liver. **P < .01, tumor compared with nontumorous liver. mRNA, messenger RNA; PPARγ, peroxisome proliferator-activated receptor.

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Among the five HCC lines (Hep3B, HepG2, Huh7, HTC, and ARL6) tested, all expressed PPARγ mRNA. Based on their apparently diverse genetic basis for hepatocarcinogenesis, we selected two cell lines derived from human HCCs for further study: Hep3B, which exhibits p53 deletion apparently associated with an integrated hepatitis B virus genome, and Huh7, which is free of the hepatitis B virus genome and has a known p53 mutation.

Ligand-Dependent Induction of PPARγ Expression, and the Effect of PPARγ Activation on Viability of Cultured HCC Cells.

To evalute the effects of PPARγ-regulated genes on cell viability, HCC cells were cultured with varying concentrations of troglitazone, a relatively selective PPARγ agonist. Exposure to troglitazone for 48 hours produced a dose-dependent increase of PPARγ mRNA (Fig. 2A) and protein (Fig. 2B) in both Hep3B and Huh7 cells. Concomitant with this upregulation and continued activation of PPARγ by troglitazone, this agonist caused a dose-dependent suppression of cell viability (Suppl Fig. available at the HEPATOLOGY website (http://interscience.wiley.com/jpages/0270-9139/suppmat/index.html).).

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Figure 2. Expression of PPARγ mRNA and protein in Hep3B and Huh7 cells. (A) PPARγ mRNA levels were determined using real-time PCR. Both HCC cell lines expressed PPARγ, and after exposure to troglitazone for 48 hours at the concentrations indicated, PPARγ mRNA levels increased. GAPDH was used as the internal standard for RNA quantity and integrity in real-time RT-PCR. The ratio of PPARγ mRNA to GAPDH mRNA at each time point is expressed as the mean ± SD (n = 3). *P < .05, **P < .01, ***P < .001 compared with control. (B) PPARγ protein was detected via Western blotting. Inconsistent with mRNA expression, protein levels were increased in a dose-dependent manner after treatment with troglitazone for 48 hours. GAPDH was used as the internal standard. mRNA, messenger RNA; PPARγ, peroxisome proliferator-activated receptor.

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PPARγ Agonist Induces G0/G1 Cell Cycle Arrest in Cultured HCC Cells.

To determine whether troglitazone decreased viability of cultured HCC cells by inhibiting cell growth, we first investigated the effect of this PPARγ agonist on cell cycle distribution. After propidium iodide staining, FACs analysis of troglitazone-treated HCC cells revealed a dose-dependent decrease in the number of cells in S phase at 48 hours (Fig. 3A). Western blot analysis showed a dose-dependent decrease PCNA expression (Fig. 3B), confirming the inhibitory effect of PPARγ activation on cell cycle progression in HCC cells. Concomitant with this inhibition, there was a dose-dependent increase in the number of cells accumulating in the G0/G1 phase (Fig. 3C), consistent with the proposal that PPARγ activation blocks the cell cycle at the G0/G1 checkpoint.

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Figure 3. Effect of troglitazone on proliferation and cell cycle regulation of Hep3B and Huh7 cells. (A) Cell proliferation was calculated as the fraction of cells in S phase determined via flow cytometry for the indicated concentration of troglitazone. (B) Expression of PCNA as determined via Western immunoblotting after treatment with troglitazone for 48 hours. Concomitant with the reduced number of cells in S phase, a dose-dependent decrease in expression of PCNA is evident in both human HCC cell lines after exposure to troglitazone. GAPDH was used as the internal standard. (C) The number of cells in G0/G1 phases was also determined via flow cytometry. Values are expressed as the mean ± SD from five replicate experiments. *P < .05; **P < .001, troglitazone-treated versus untreated control. PCNA, proliferating cell nuclear antigen.

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Growth Arrest by Troglitazone Is Associated With Induction of p27 but Not of p21.

Cell cycle arrest caused by the overexpression of p53 has been associated with induction of p21 and p27.11–13 To establish which of these pathways were responsible for the troglitazone effects, we examined their expression via Western Blot analysis in Huh7 cells (Hep3B cells lack p53). As shown in Fig. 4A, p53 protein expression increased progressively in the Huh7 cell line subjected to troglitazone treatment. However, p21, a downstream target protein of p53, was not expressed (data not shown), indicating that the increased level of p53 protein as a transcriptional activator did not mediate p21 expression. This result encouraged us to analyze the protein expression level of p27, a regulator involved in G0/G1 arrest. Exposure to troglitazone for 48 hours (Fig. 4B) caused an obvious induction of p27 in Huh7 cells. This finding indicated that induction of p27 could mediate PPARγ-stimulated cell cycle arrest in this cell line.

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Figure 4. Effect of troglitazone on p53 and p27. Hep3B and Huh7 cells were treated with increasing concentrations of troglitazone for 48 hours. (A) p53 and (B) p27 were determined via Western blotting. GAPDH was used as the internal control.

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Induction of Apoptosis via PPARγ.

Arrest of cell cycle progression in tumor cells is usually associated with concomitant activation of cell death pathways, and this can be mediated by p53. We therefore examined the contribution of apoptosis to the observed loss of viability of troglitazone-treated HCC cell cultures. By FACs analysis, the number of cells with sub-G1 DNA content after 48 hours of troglitazone treatment was substantially increased (Fig. 5A). Consistent with this finding, there was an increased number of cells that stained TUNEL-positive after troglitazone treatment compared with untreated HCC cells (Fig. 5B).

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Figure 5. Effect of troglitazone on apoptosis of human HCC cells. (A) Troglitazone increased the rate of apoptosis, as determined by the number of cells with sub-G1 DNA content by flow cytometry. Values are expressed as the mean ± SD of five replicate experiments. *P < .001, troglitazone-treated versus untreated controls. (B) TUNEL staining of Hep3B cells incubated either alone (a1) or in the presence of troglitazone (100 μmol/L) (a2) for 48 hours; Huh7 cells were incubated either alone (b1) or in the presence of troglitazone (100 μmol/L) (b2) for 48 hours; An increase in the number of TUNEL-positive cells (arrows) is evident in the presence of troglitazone. (Original magnification ×400.)

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Relationships Between PPARγ and COX-2 Expression in Liver Tumors and HCC Cell Lines.

To investigate whether the inhibitory effects of PPARγ activation on the growth of HCC cells could be mediated via regulation of COX-2, we determined expression of COX-2 in clinical samples of HCC. Compared with the surrounding nontumorous liver, COX-2 mRNA (P < .01) and COX-2 protein (P < .05) were both increased in HCC tissues. There was a negative correlation between PPARγ and COX-2 mRNA expression in tumor samples (r = −0.48; P < .05), and a similar reverse correlation was demonstrated by parallel immunoblotting of HCC specimens (r = −0.76; P < .05). In addition, we determined the effects of troglitazone on COX-2 mRNA and protein expression in cultured HCC cells. After 48 hours, troglitazone caused a dose-dependent reduction in COX-2 mRNA (Fig. 6A) and protein (Fig. 6B) expression.

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Figure 6. Effect of troglitazone on COX-2 mRNA and protein expression in cultured human HCC cells. Expression of (A) COX-2 mRNA, and (B) COX-2 protein after exposure to the indicated concentration of troglitazone for 48 hours. COX-2 mRNA was determined via RT-PCR and COX-2 protein was determined via Western immunoblotting using GAPDH as the internal standard. A dose-dependent decrease in expression of COX-2 at both the mRNA and protein levels was observed with increasing concentrations of troglitazone. Data are expressed as the mean ± SD (n = 3), *P < .05, troglitazone-treated versus untreated control. COX-2, cyclooxygenase 2.

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Troglitazone, a PPARγ Agonist, Inhibits Liver Tumor Growth In Vivo.

In light of the observed antiproliferative and proapoptotic effects of troglitazone on HCC cell lines in vitro, we tested whether troglitazone treatment could alter growth of liver cancer cells in vivo. For these studies, we established the experimental model of Huh7 tumor–bearing Balb/c nude mice and then treated these animals with troglitazone (200 ppm mixed in the chow) or vehicle. Three experimental approaches were studied: (1) the ability of troglitazone to prevent or slow development of subcutaneous tumors (prevention study), (2) the capacity of this PPARγ agonist to slow or reverse tumor growth when introduced at an early stage of tumor development or (3) at a late stage of established tumors (regression studies).

In the prevention study, troglitazone (200 ppm) treatment was started on the day of Huh7 cell injection. After 25 days, all four vehicle-treated control mice developed visible subcutaneous Huh7 tumors. In contrast, only three of five troglitazone-treated animals developed subcutaneous Huh7 cell tumors (Fig. 7A1-A3). In each case, these tumors were smaller than in control mice, so that the overall inhibition of tumor growth exerted by PPARγ agonist treatment was estimated as 89% (Fig. 7A4-5).

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Figure 7. (A) Troglitazone inhibits growth of tumors derived from HCC cells in vivo. Following subcutaneous inoculation of 4-week-old male Balb/c nude mice with Huh7 cells (1 × 106 cells), animals were randomized to receive either oral troglitazone (200 ppm) or vehicle for 25 days. (A1) Effect on tumor size of troglitazone administered in chow. Tumor growth is shown in nude mice fed with (A2) vehicle (positive control) or (A3) troglitazone, which was expressed as (A4) volume (see Methods) and (A5) weight. (B) Experiments were conducted after tumors had developed to determine the effect of troglitazone on tumor growth. (B1) Twelve nude mice were injected with 1 × 106 Huh7 cells and troglitazone (200 ppm). Treatment was initiated in one group (n = 6) of mice when the tumor size was 2-4 mm (on day 12 after Huh7 injection). The experiment was terminated 13 days later. Tumor growth was expressed as (B2) volume (see Materials and Methods) and (B3) weight. (C1) Fourteen nude mice were injected with 1 × 106 Huh7 cells. Administration of troglitazone (200 ppm) to half of the mice (n = 7) was started when the tumor size had already developed to 8-10 mm (on day 15 after Huh7 injection). The experiment was terminated 9 days later. Tumor growth was expressed as (C2) volume (see Materials and Methods) and (C3) weight. Data are expressed as the mean ± SD. *P < .05, **P < .01, troglitazone-treated compared with control.

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To assess whether troglitazone could dampen liver cancer growth after creation of subcutaneous tumors, 12 nude mice were injected with 1 × 106 Huh7 cells. In one study, troglitazone was initiated when the tumor size was 2-4 mm. When the experiment was terminated 13 days later, mice treated with troglitazone had an 88% reduction in tumor growth compared with the positive controls (Fig. 7B1-B3). In a separate experiment, 14 nude mice were injected with 1 × 106 Huh7 cells, and administration of troglitazone (200 ppm) was started when the tumor size had already developed to 8-10 mm. Nine days later, the experiment was ended. Mice treated with troglitazone had a 42% reduction in tumor growth compared with controls (Fig. 7C1-C3). No drug toxicity was observed with this dose of troglitazone.

Histological analysis of Huh7 tumors from untreated mice revealed poorly differentiated, infiltrating hepatocellular carcinomas that exhibited high mitosis rates (Fig. 8A1). In contrast, tumors from mice receiving troglitazone showed few mitotic cells (Fig. 8A2). Compared with the control tumors, mitotic rates were significantly less in tumors from mice receiving troglitazone (P < .05), and PCNA expression was less than in the positive control tumors (Fig. 8C). In addition to this evidence that troglitazone decreased the proliferative rate of subcutaneously implanted HCCs, we obtained evidence that troglitazone also significantly increased their rate of apoptosis. Thus, while Huh7 tumors from untreated mice showed almost no TUNEL staining (Fig. 8B1), tumor cells from mice receiving troglitazone showed frequent condensed nuclei and cytoplasmic shrinkage, indicating the operation of apoptosis (P < .05) (Fig. 8B2). Compared with the control tumors, an increased expression of p27 and decreased expression of COX-2 were exhibited in tumors from mice receiving troglitazone (Fig. 8C).

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Figure 8. Antitumor effect of troglitazone in vivo is attributable to suppression of cell proliferation and induction of apoptosis. (A1) Hematoxylin-eosin–stained sections of Huh7 tumors from untreated controls showed numerous cells in mitosis (arrows). (A2) Troglitazone (200 ppm) treatment suppressed cell proliferation, as indicated by the smaller number of mitotic cells. (Original magnification ×200.) A reduction in the number of mitosis cells is exhibited in troglitazone-treated tumor *P < .05, troglitazone-treated compared with control. (B1) TUNEL-stained sections of Huh7 tumors from untreated control displayed occasionally apoptotic cells (arrows) (0.58 ± 0.2). (Original magnification ×200.) (B2) Tumors from mice treated with troglitazone had a prominent population of cells with condensation or fragmentation of nuclei and cytoplasm shrinkage (arrows), indicating apoptosis (6.60 ± 2.3; P < .001, troglitazone-treated compared with control). (Original magnification ×200.) (C) An increased expression of p27, decreased expression of COX-2 and PCNA are evident in troglitazone-treated tumors. GAPDH was used as the internal standard. Data are expressed as the mean ± SD (n = 3-4/group), *P < .05, **P < .01, **P < .001 compared with control. COX-2, cyclooxygenase; PCNA, proliferating cell nuclear antigen.

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Discussion

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. References
  7. Supporting Information

In the present study, we found a significant decrease in expression of PPARγ mRNA and protein in human liver cancers compared with surrounding nontumorous liver. Similar downregulation of PPARγ expression in tumor tissue compared with adjacent nontumorous mucosa has also been observed in human esophageal,14 breast,15 and ovarian cancers.16 Several in vitro and in vivo model systems suggest that PPARγ may behave as a tumor suppressor gene.5–7, 17, 18 In the breast, inactivation of PPARγ with induction of COX-2 occurred during the development and progression of human breast carcinoma.15, 19 On the other hand, inhibition of COX-2 and activation of PPARγ prevented mammary carcinogenesis in experimental animals.20, 21 Other authors have reported that PPARγ activation can enhance tumor formation in different murine models.8, 9, 22 Given these apparently conflicting data on whether PPARγ activation could be growth-inhibitory or tumor-promoting, a better understanding of the mechanism of action of PPARγ in liver cancer is required.

We analyzed the effects of PPARγ activation in two human HCC cell lines. In both Huh7 and Hep3B cells, the PPARγ agonist, troglitazone, caused a dose-dependent increase in expression of PPARγ. Concomitant with PPARγ upregulation, there was dose-dependent growth arrest; similar results have been observed in other HCC cell lines.23, 24 FACS analysis of the effects of troglitazone on the cell cycle in treated HCC cells revealed a dose-dependent decrease in cell proliferation, a concomitant and proportionate increase of cells in G0/G1, and an increase of cells in the sub-G1 fraction. The latter finding indicates increased apoptosis, a finding confirmed by TUNEL staining.

Because downstream genes regulated by PPARγ have not been well characterized, it is difficult to speculate about the underlying mechanisms for the antiproliferative effects caused by PPARγ signaling in liver cancer cells. On the basis of the immunoblot analysis of known cell cycle inhibitors, G1 arrest by troglitazone was associated with induction of p27 but not p21. In agreement with a recent report,25 this indicates that hepatocyte cell growth arrest can be mediated by p27 when p21 is downregulated. In the present study, the induction of p27 by troglitazone in Hep3B cells, which lack p53, as well as in Huh7 cells, in which p53 is mutated, indictes that the mechanism of PPARγ–mediated p27 induction is likely to be independent of p53. p27 is a potent inhibitor of cyclin D/cyclin-dependent kinase (Cdk) 4 and cyclin E/Cdk2, which kinases govern cell cycle progression at the restriction and late transition points of G1, respectively. The role of p27 as a major player in G1 arrest has been well accepted, and p27 is generally expressed at high levels in quiescent cells.11, 26, 27 It is also known that p53, acting through p21, also plays an important role in replicative senescence.11–13 It was therefore of interest in the present study to observe that the troglitazone-induced p53 in Huh7 cells was nonfunctional in terms of p21 induction, although troglitazone induced cell arrest. The reason for the inability of enhanced p53 to induce p21 in Huh7 cells could be explained by mutation of p53. Furthermore, p53-independent replicative senescence has been demonstrated in some studies in cells that lack p53.28, 29 Other investigations have shown that PPARγ activation can cause G1 cell cycle arrest through a mechanism that involves downregulation of protein phosphatase 2A,30 cyclin D1, and cyclin E.31

In the present work, we showed that the growth inhibitory effect of PPARγ activation by troglitazone was also related to induction of apoptosis. In addition to the results in cultured HCC cell lines, treatment with troglitazone increased apoptosis in xenografted tumors in nude mice. Rumi et al.23 and Koga et al.24 have also reported that troglitazone induces apoptosis of human HCC cells. However, others have indicated that apoptosis may not be the predominant mechanism through which troglitazone inhibits growth of lung cancer cells.31, 32 Others have suggested that the mechanisms through which troglitazone can induce apoptosis in breast cancer cells include reduced expression of Bcl-233 or activation of TRAIL,34 both resulting in caspase 3 activation.33

In the present study, there appeared to be a striking reciprocity between the downregulation of PPARγ we detected in liver cancer and the increased expression of COX-2, which we and others have found to be enhanced in the majority of HCCs.24, 35 A reciprocal relationship between PPARγ activity and COX-2 expression has also been observed in colon,36 cervical,37 breast,38 and tongue cancer cells.39 These findings raise the possibility that high expression of COX-2 might result from the removal of regulatory suppression by PPARγ; the inverse relationship between PPARγ expression and COX-2 expression levels in human HCC tumors noted in the present study is consistent with this proposal. We also observed that troglitazone induced a dose-dependent increase in expression of PPARγ with concomitant downregulation of COX-2 in HCC cells. The same effect was also observed in Huh7 xenograft tumors in nude mice, suggesting that COX-2 expression in HCC could be regulated through PPARγ. In other tissues, PPARγ signaling has been implicated in the control of COX-2 expression.36, 40, 41 It is therefore possible that the inhibitory properties of troglitazone against liver cancer cell growth observed in this study could be explained, at least in part, by the resultant downregulation of COX-2 expression, but this requires further study.

Having observed substantial suppression of HCC cell growth by PPARγ agonist treatment in vitro, we conducted experiments designed to test the potential for PPARγ agonists to exert chemopreventive effects against HCC in vivo. It is recognized that the model used in this study—nude mice injected with human HCC cells subcutaneously—is an artificial one and is not necessarily equivalent to the stepwise development of hepatic carcinogenesis during chronic liver disease. Nonetheless, the observation that treatment of HCC-inoculated mice with troglitazone substantially impaired tumor development and either retarded further growth or effected partial regression of established tumors (depending on the experimental design) is both novel and of potential clinical relevance. This result is supported by a recent study of carcinogen-induced liver cancer in the rat.42 The observation of decreased mitotic rates, decreased PCNA expression, increased apoptosis, and increased P27 induced by troglitazone in vivo was entirely consistent with the in vitro effects, adding further weight to the potential significance of these findings. Because only two selected human HCC cell lines were tested here, the generalizability of these results remain unclear. However, the observation from the human samples that HCC seems to be associated with the underexpression of PPARγ—and that such underexpression can be corrected or reversed through exposure to pharmacological doses of a PPARγ ligand—adds weight to the potential efficacy of phenolizidinediones against HCC. However, the precise mechanisms for the inhibition of growth by PPAR-γ ligands needs to be investigated further.

In conclusion, we report that human HCCs underexpress PPARγ in relation to surrounding liver tissue. In HCC cell lines, the PPARγ ligand troglitazone induces PPARγ expression and inhibits cell growth both in vitro and in vivo. The mechanisms appear to involve both inhibition of cell proliferation and induction of apoptosis. We also provide further evidence that ligand-mediated PPARγ activation suppressed COX-2 expression in liver cancer cells. The possibility that PPARγ ligands may be effective chemopreventive agents against liver carcinogenesis or novel chemotherapeutic agents is raised by these studies.

References

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  5. Discussion
  6. References
  7. Supporting Information
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Supporting Information

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. References
  7. Supporting Information

Supplementary material for this article can be found on the H EPATOLOGY website ( http://interscience.wiley.com/jpages/0270-9139/suppmat/index.html )

FilenameFormatSizeDescription
jws-hep.20994.tif575KThe effects of troglitazone on viability of Hep3B and Huh7 cells in culture. HCC cells (1 x 10 6cells) were incubated either alone or in the presence of troglitazone (0.50, 100 &&num;x03bc;mol/L) for 48 hours and cell viability was measured by the MST method. Treatment with troglitazone caused dose-dependent inhibition of cell viability. *P < .05, **P < .001, troglitazone treated vs. untreated control. Data are mean &&num;x00b1; SD (n = 3).

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