Abstract

Peroxisome proliferators, such as lipid-lowering fibrate drugs, are agonists for the peroxisome proliferator-activated receptor α (PPARα). Sustained activation of PPARα leads to the development of liver tumors in rodents. Paradoxically, humans appear to be resistant to the induction of peroxisome proliferation and development of liver tumors by peroxisome proliferators. To examine the species differences in response to peroxisome proliferators, a PPARα humanized mouse (hPPARα) was generated, in which the human PPARα was expressed in liver under control of the Tet-OFF system. To evaluate the susceptibility of hPPARα mice to peroxisome proliferator-induced hepatocarcinogenesis, a long-term feeding study of Wy-14,643 was carried out. hPPARα and wild-type (mPPARα) mice were fed either a control diet or one containing 0.1% Wy-14,643 for 44 and 38 weeks, respectively. Gene expression analysis for peroxisomal and mitochondrial fatty acid metabolizing enzymes revealed that both hPPARα and mPPARα were functional. However, the incidence of liver tumors including hepatocellular carcinoma was 71% in Wy-14,643-treated mPPARα mice, and 5% in Wy-14,643-treated hPPARα mice. Upregulation of cell cycle regulated genes such as cd1 and Cdks were observed in non-tumorous liver tissue of Wy-14,643-treated mPPARα mice, whereas p53 gene expression was increased only in the livers of Wy-14,643-treated hPPARα mice. These findings suggest that structural differences between human and mouse PPARα are responsible for the differential susceptibility to the peroxisome proliferator-induced hepatocarcinogenesis. This mouse model will be useful for human cancer risk assessment of PPARα ligands.

Introduction

A variety of chemicals such as hypolipidemic drugs, phthalate ester plasticizers and industrial solvents have been shown to increase the size and number of peroxisomes in rats and mice ( 1 ). Because of their unique property of inducing peroxisome proliferation, they are grouped as peroxisome proliferators ( 2 ). The hepatic changes caused by treatment with peroxisome proliferators include liver hyperplasia and hypertrophy, proliferation of peroxisomes, and increases in the oxidation of fatty acids through induction of genes encoding mitochondrial, peroxisomal and microsomal enzymes involved in fatty acid oxidation ( 3 , 4 ). Long-term treatment of mice and rats with peroxisome proliferators eventually results in the formation of hepatocellular adenomas and/or carcinomas; these chemicals are regarded as non-genotoxic carcinogens since they do not directly cause genetic damage ( 59 ).

Peroxisome proliferators exert their effects by activating peroxisome proliferator-activated receptor α (PPARα), a member of the superfamily of ligand-activated, nuclear receptors. In response to ligand activation of PPARα, there is an increase in expression of target genes encoding enzymes and proteins involved in fatty acid transport and catabolism ( 3 ). In rodent model systems, PPARα has also been implicated in regulating a number of other biological processes including cell proliferation, apoptosis, inflammation and oxidative stress. Targeted disruption of the mouse Ppar α gene confirmed that this receptor is responsible for the peroxisome proliferator-induced pleiotropic responses in mice including the process of hepatocarcinogenesis ( 5 , 10 ).

Notable species differences in response to peroxisome proliferators have been demonstrated ( 11 , 12 ). In spite of their carcinogenic activity in mice and rats, primate and humans appear to be resistant to the induction of peroxisome proliferation and the development of hepatocarcinomas after chronic exposure to the peroxisome proliferator fibrate drugs ( 1315 ). However, humans treated with gemfibrozil, a hypolipidemic drugs, for many years have shown no evidence for increased peroxisomes or other pathological alterations found in rodents ( 14 ). In addition, epidemiological studies on hypolipidemic drugs showed no increase in cancer incidence ( 16 ). Although human PPARα was shown to be functional in transactivation assays ( 1719 ), the mechanism underlying the species difference in response to peroxisome proliferators is still unknown.

Recently, a novel transgenic mouse line humanized for PPARα (hPPARα mice) was generated in order to elucidate the mechanism of species difference in response to peroxisome proliferators ( 20 ). These mice express human PPARα specifically in the liver under the control of the doxycycline (Tet-OFF) system in a PPARα null background. When treated for up to 8 weeks with the potent peroxisome proliferator, [4-chloro-6-(2,3-xylidino)-pyrimidynylthio]acetic acid (Wy-14,643), hPPARα mice exhibit decreased serum triglycerides and increased expression of genes encoding peroxisomal, mitochondrial and microsomal fatty acid oxidation enzymes, albeit to a lesser extent than wild-type mice. However, unlike wild-type mice, the hPPARα mice did not display increases in ligand-induced hepatomegally, cell proliferation and expression of cell cycle control genes. These data suggest that the hPPARα mouse may be resistant to peroxisome proliferator-induced hepatocarcinogenesis.

In the present study, to determine the susceptibility of the hPPARα mice to peroxisome proliferators-induced hepatocarcinogenesis, a long-term feeding study of Wy-14,643 was carried out. Expression of some direct PPARα target genes involved in the regulation of the cell cycle was also analyzed.

Materials and methods

Animals and treatments

A mouse line humanized for the hPPARα was generated as previously described ( 20 ). Briefly, a tetracycline response element (TRE) driving human PPARα cDNA transgene (TRE-hPPARA) was microinjected into fertilized FVB/N mouse eggs, and transgene-positive mice were mated to mice expressing the tetracycline-controlled transactivator (tTA) transgene, under the control of the liver-specific Cebp /β promoter (tTALap5Bjd), ( 21 ). Mice expressing both transgenes were bred into a mouse PPARα-null background (129/S4/SvJae-Ppara tm1Gonz/J ) to generate CEBP/β-tTA; TRE-hPPARα; PPARα-null transgenic mice (129S4/SvJae.Cg-Ppara tm1Gonz/J Tg(TRE-hPPARA)1Gonz Tg(tTALap)5Bjd). This mouse line expresses human PPARα cDNA in the liver hepatocytes on a mouse PPARα-null background in the absence of doxycycline, a tetracycline derivative (Tet-OFF system). The genotypes for all animals used in this study were verified using PCR detection of tTA (tTA forward, 5′-CTCGCCCAGAAGCTAGGTGT-3′; tTA reverse, 5′-CCATCGCGATGACTTAGT-3′, recognizing at 200 bp), mouse PPARα (mαF1, 5′-GAGAAGTTGCAAGGAGGGGATTGTG-3′; mαR1, 5′-CCCATTTCGGTAGCAGGTAGTCTT-3′; and mαNEOR1, 5′-GCAATCCATCTTGTTCAATGGC-3′, recognizing wild-type allele at ∼400 bp and the null allele at ∼650 bp), and human PPARα (hPPAR2 GENO F3, 5′-CGATTTCACAAGTGCCTTTCTGTC-3′; hPPAR2 GENO R3, 5′-AAGTTTGCGAAGCCTGGGATGG-3′, recognizing at 430 bp).

A total of 30, 6-week-old male hPPARα mice were divided into two groups and 10 and 20 mice were given a pelleted diet with or without 0.1% (w/w) Wy-14,643 (ChemSyn Science Laboratories, Lenexa, KS, USA) up to week 44 of the experimental period, respectively. Similarly, a total of 19, 6-week-old male wild-type mice (mPPARα mice) with the same background were divided into two groups, and 9 and 10 animals were given diet with or without Wy-14,643 until the final killing at week 38 of the experimental period, respectively. Animals were housed 4–5 in plastic cages with hard wood chips for bedding in an animal room with a 12 h light/dark cycle, carefully monitored general health status, and the body weights were measured weekly until week 5, after which weights were measured once every two weeks. Those becoming moribund during the course of the study were killed by CO 2 asphyxiation and examined histologically before the termination of the experiment. All animal experimental procedures were carried out in accordance with animal study protocols approved by the National Cancer Institute Animal Care and Use Committee.

Histological analysis

Mice were killed at the termination of the experiment and liver tissues were carefully investigated for the existence of tumors after being weighed, and small portions of liver tissues without any macroscopic nodules were snap frozen in liquid nitrogen and stored at −80°C until further analysis. The remaining liver tissues were fixed in 10% phosphate-buffered formalin, embedded in paraffin, sectioned at 3–4 µM, and prepared for routine hematoxylin and eosin staining for histological examination. The lesions and tumors found were classified according to the criteria of the International Agency for Research on Cancer (WHO) ( 22 ).

Western blot analysis

Nuclear extracts from liver samples were prepared using NE–PER nuclear extraction kit (Pierce Chemical, Rockford, IL, USA), separated on 10% SDS–PAGE and blotted onto nitrocellulose membrane (Whatman, Stanford, ME, USA). The membrane was incubated overnight at 4°C with a 1:750 dilution of a primary antibody against PPARα (Geneka Biotechnology, Montreal, Canada) and GAPDH (Chemicon International, Temecula, CA, USA). After washing, the membrane was incubated for 30 min with 1:10 000-diluted horseradish peroxidase-conjugated secondary antibodies (Sigma, St Louis, MO, USA), and the reaction product was visualized using SuperSignal West Pico an enhanced Chemiluminescent Substrate (Pierce).

Northern blot analysis

Total RNA was extracted from liver using TRIzol reagent (Invitrogen, Carlsbad, CA, USA). Northern blot analysis was carried out as described previously ( 23 ) and hybridized with random primer 32 P-labeled cDNA probes ( 23 , 24 ), and exposed to a PhosphorImager screen cassette followed by visualization using a Storm 860 PhosphorImager system (Amersham Biosciences, NJ, USA). Signals were quantified using ImageQuant TL software (Amersham Biosciences). The expression levels of target genes were normalized with those of reference gene (acidic ribosomal phosphoprotein; 36B4). Five mice each per group were subjected to the analysis.

Statistical analysis

Survival curves were created using the Kaplan–Meier method, and the statistical significance of differences was calculated by the log-rank test. Variations in liver/body weight ratio, incidences of tumors, gene expression analysis between the different treatments or animal strains were evaluated with one-way fractorial ANOVA and multiple comparison tests. All the calculations for statistical analysis were performed using the Statview SE+ Graphics, version 5.0 (Abacus Concepts, Berkeley, CA, USA).

Results

Survival, body and liver weights

From the onset of the experiment, Wy-14,643-treated mPPARα mice exhibited suppressed growth compared with all other groups and the percentage of animals of this group that survived decreased to 50% at the termination of the study period ( Figures 1 and 2 ). Of the 10 mice in the Wy-14,643-treated mPPARα group, 3 died of toxicity and were not evaluated further whereas 2 mice were killed due to morbidity before the termination of the experiment. However, the Wy-14,643-treated hPPARα mice showed a similar rate of growth as those of the non-treated control mice of both strains, and only one animal in this group was killed due to morbidity before termination of the experiment ( Figures 1 and 2 ).

Fig. 1.

Time-course changes in mean body weights. Mice were weighed every 2 weeks throughout the course of the study. The mPPARα mice + Wy-14,643 group showed a lower rate of growth from the onset of the experiment.

Fig. 1.

Time-course changes in mean body weights. Mice were weighed every 2 weeks throughout the course of the study. The mPPARα mice + Wy-14,643 group showed a lower rate of growth from the onset of the experiment.

Fig. 2.

Time-course changes in survival rates. Mice were examined twice a week and those exhibiting distress were killed. Surviving curves were created using the Kaplan–Meier method. *P < 0.05 versus all other groups (log-rank test).

Fig. 2.

Time-course changes in survival rates. Mice were examined twice a week and those exhibiting distress were killed. Surviving curves were created using the Kaplan–Meier method. *P < 0.05 versus all other groups (log-rank test).

The final body weight of Wy-14,643-treated mPPARα mice was markedly decreased and both absolute and relative liver weights increased as compared with those of all other groups ( Table I ). Wy-14,643-treated hPPARα mice showed an increased absolute and relative liver weights compared with the corresponding control diet group; however, the extent of increase was considerably less that found in the Wy-14,643-treated mPPARα mice ( Table I ).

Table I.

Final body and absolute/relative liver weights

Genotype Treatment Effective no. of mice Final body weight (g)  Liver weight
 
 

 

 

 

 
Absolute (g)
 
Relative (%) a
 
hPPARα Control diet 10  39.2 ± 7.9 b 1.57 ± 0.4 3.96 ± 0.4 
hPPARα Wy-14,643 20 40.3 ± 6.1  2.37 ± 0.5 *  5.87 ± 0.7 * 
mPPARα Control diet 33.9 ± 6.4 1.10 ± 0.1 3.29 ± 0.4 
MPPARα Wy-14,643  16.4 ± 2.9 ***  3.03 ± 1.4 **  17.86 ± 5.6 *** 
Genotype Treatment Effective no. of mice Final body weight (g)  Liver weight
 
 

 

 

 

 
Absolute (g)
 
Relative (%) a
 
hPPARα Control diet 10  39.2 ± 7.9 b 1.57 ± 0.4 3.96 ± 0.4 
hPPARα Wy-14,643 20 40.3 ± 6.1  2.37 ± 0.5 *  5.87 ± 0.7 * 
mPPARα Control diet 33.9 ± 6.4 1.10 ± 0.1 3.29 ± 0.4 
MPPARα Wy-14,643  16.4 ± 2.9 ***  3.03 ± 1.4 **  17.86 ± 5.6 *** 
*

P < 0.05 versus hPPARα + Control; **P < 0.05 versus all other groups; ***P < 0.0001 versus all other groups.

a

Liver wt/body wt (%).

b

Mean ± SD.

Hepatic lesions

Most of the mPPARα mice fed the Wy-14,643 diet had multiple, grossly visible nodules that were randomly distributed among the liver lobes ( Figure 3A ). The hepatocellular adenomas were well-circumscribed lesions compressing adjacent parenchyma without normal lobular architecture, and were composed of well-differentiated hepatocytes ( Figure 3E ). The hepatocellular carcinomas were characterized by a trabecular or pseudoglandular histologic pattern ( Figure 3F ).

Fig. 3.

Representative photomicrographs of liver lesions. ( A ) A grossly visible hepatic nodular mass is seen in the liver of a wild-type mouse after 38 weeks of Wy-14,643 feeding. ( B ) Photomicrograph of the eosinophilic altered cell foci in a wild-type mouse after 38 weeks of Wy-14,643 feeding. ( C ) High magnification of (B) shows severe toxic lesions including large eosinophilic hepatocytes with eosinophilic granular cytoplasm, pigment accumulation within Kupffer cells (arrows) and small basophilic cells of proliferating oval cells. ( D ) A photomicrograph of livers from a humanized PPARα mouse after 44 weeks of Wy-14,643 feeding. Mild fatty change and glycogen deposition are seen, but the toxic lesions are not evident compared with liver from wild-type mice after 38 weeks of Wy-14,643 feeding. ( E ) and ( F ) A photomicrograph of a representative hepatocellular adenoma and carcinoma in wild-type mouse after 38 weeks of Wy-14,643 feeding, respectively.

Fig. 3.

Representative photomicrographs of liver lesions. ( A ) A grossly visible hepatic nodular mass is seen in the liver of a wild-type mouse after 38 weeks of Wy-14,643 feeding. ( B ) Photomicrograph of the eosinophilic altered cell foci in a wild-type mouse after 38 weeks of Wy-14,643 feeding. ( C ) High magnification of (B) shows severe toxic lesions including large eosinophilic hepatocytes with eosinophilic granular cytoplasm, pigment accumulation within Kupffer cells (arrows) and small basophilic cells of proliferating oval cells. ( D ) A photomicrograph of livers from a humanized PPARα mouse after 44 weeks of Wy-14,643 feeding. Mild fatty change and glycogen deposition are seen, but the toxic lesions are not evident compared with liver from wild-type mice after 38 weeks of Wy-14,643 feeding. ( E ) and ( F ) A photomicrograph of a representative hepatocellular adenoma and carcinoma in wild-type mouse after 38 weeks of Wy-14,643 feeding, respectively.

Table II shows incidence of the hepatocellular adenoma/carcinoma and pre-neoplastic altered cell foci in the liver of the various groups. The incidence of hepatocellular adenomas in the Wy-14,643-treated mPPARα mice was 71% and two animals had hepatocellular carcinomas. Most animals in this group with tumors had single or multiple altered cell foci ( Figure 3B and C ). In contrast, only 1 out of 20 hPPARα mice treated with Wy-14,643 harbored a hepatocellular adenoma, and significant differences in the incidence was detected as compared with Wy-14,643-treated mPPARα mice. This tumor was also morphologically similar to spontaneous mouse liver tumor with basophilic and clear hepatocytes, whereas the tumors in mPPARα mice were more diffusely basophilic. Altered cell foci were detected in one hPPARα animal in each of the mice with and without Wy-14,643 treatment. Histologically, toxic lesions induced by Wy-14,643 were seen in mPPARα mice and were much less severe in hPPARα mice ( Figure 3B–D ).

Table II.

Incidences of liver lesions

Genotype Treatment Effective no. of mice Altered foci (%)  Tumor (%)
 
  

 

 

 

 
Adenoma
 
Carcinoma
 
Total
 
hPPARα Control diet 10 1 (10) 0 (0) 0 (0) 0 (0) 
hPPARα Wy-14,643  20 a 1 (5) 1 (5) 0 (0) 1 (5) 
mPPARα Control diet 0 (0) 0 (0) 0 (0) 0 (0) 
mPPARα Wy-14,643  7 a 5 (71) 5 (71) 2 (29)  5 (71) * 
Genotype Treatment Effective no. of mice Altered foci (%)  Tumor (%)
 
  

 

 

 

 
Adenoma
 
Carcinoma
 
Total
 
hPPARα Control diet 10 1 (10) 0 (0) 0 (0) 0 (0) 
hPPARα Wy-14,643  20 a 1 (5) 1 (5) 0 (0) 1 (5) 
mPPARα Control diet 0 (0) 0 (0) 0 (0) 0 (0) 
mPPARα Wy-14,643  7 a 5 (71) 5 (71) 2 (29)  5 (71) * 
*

P < 0.01 versus values of hPPARα + Wy-14,643.

a

Includes liver from mice that were euthanized due to morbidity before the termination of experiment.

PPARα expression

At the termination of the carcinogenicity study, both gene and protein expression levels of PPARα were evaluated using the non-tumorous portion of liver tissue ( Figure 4 ). Total RNAs were hybridized with specific probes for human and mouse PPARα genes independently for northern blot analysis. Both hPPARα and mPPARα mice showed expression of the respective human and mouse PPARα mRNA regardless the treatment of Wy-14,643 ( Figure 4A ). Moreover, PPARα proteins were also expressed similarly in hPPARα and mPPARα mice ( Figure 4B ).

Fig. 4.

Gene expression ( A ) and protein expression ( B ) of PPARα. (A) Total RNAs from livers of non-cancerous tissue were hybridized with specific probes for human and mouse PPARα genes. (B) Nuclear extracts proteins were subjected to western blots using polyclonal PPARα antibody. Nuclear extracts from liver tissue of PPARα-null mice and from PPARα expressing HepG2 cells were used as negative and positive controls, respectively. Cont., control diet; Wy-14,643, control diet containing 0.1% Wy-14,643.

Fig. 4.

Gene expression ( A ) and protein expression ( B ) of PPARα. (A) Total RNAs from livers of non-cancerous tissue were hybridized with specific probes for human and mouse PPARα genes. (B) Nuclear extracts proteins were subjected to western blots using polyclonal PPARα antibody. Nuclear extracts from liver tissue of PPARα-null mice and from PPARα expressing HepG2 cells were used as negative and positive controls, respectively. Cont., control diet; Wy-14,643, control diet containing 0.1% Wy-14,643.

Gene expression analysis

To examine the expression of PPARα target genes and genes involved in cell cycle/apoptosis, northern blot analyses were carried out ( Figure 5A ), using non-tumorous portions of liver collected at the termination of the animal experiment, and quantification of expression normalized with a reference gene ( Figure 5B–D ). Expression levels of acyl-CoA oxidase (ACOX), cytochrome P450 A (CYP4A) and medium chain acyl-CoA dehydrogenase (MCAD) mRNAs in both Wy-14,643-treated hPPARα and mPPARα mice were significantly increased, compared with those of corresponding groups fed the control diet ( Figure 5B ). mRNA encoding malic enzyme (ME) was significantly increased in the Wy-14,643-treated mPPARα group, compared with that of the control diet group, but no difference was detected between Wy-14,643- and control-diet-treated hPPARα mice groups. Figure 5C shows the expression level of genes regulating cell cycles. mRNAs encoding CD1, cyclin-dependent kinases (CDKs) 1 and 4 were highly expressed in the livers of the Wy-14,643-treated mPPARα mouse group and were statistically different from those in all other groups. The cMYC mRNA was also significantly overexpressed in the Wy-14,643-treated mPPARα group compared with those of control-diet-treated mPPARα group, but no difference was detected compared with those of other groups. The mRNAs encoding apoptosis associated proteins, p53, p21, BAX and BCL2 were also examined ( Figure 5D ). Expression of the p53 gene was increased in the Wy-14,643-treated hPPARα group as compared with those of all other groups, although the P -value between Wy-14,643-treated mPPARα was marginal ( P = 0.416). The p21 mRNA showed a slight but not statistically significant increase in the Wy-14,643-treated hPPARα. No difference was found in expression of mRNAs encoding BAX and BCL2 among all groups.

Fig. 5.

Northern blot analysis of PPARα target genes and cell cycle/apoptosis regulating genes using probes as indicated ( A ). Total RNA (10 µg/lane) was isolated from livers of non-cancerous tissue ( n = 5 animals) from each animal. The signal for acidic ribosomal phosphoprotein (36B4) was used as a control for loading and RNA integrity. Cont., control diet; Wy-14,643, control diet containing 0.1% Wy-14,643; ACOX, acyl-CoA oxidase; CYP4A, cytochrome P450 4A family; MCAD, medium chain acyl-CoA dehydrogenase; ME, malic enzyme; C MYC, c-myc; CD1, cyclin D1; CDK, cyclin-dependent kinase. Quantification analysis of gene expression (B, C and D). All values were normalized to the signal for 36B4. Results represent mean ± SD. ( B ) Results for PPARα target genes. *P < 0.05, **P < 0.01 versus corresponding to mice fed with control diet. ( C ) Results for cell cycle regulating genes. *P < 0.05, **P < 0.01 versus mPPARα fed with diet containing 0.1% Wy-14,643. ( D ) Results for apoptosis regulating genes. *P < 0.05, **P < 0.01 versus hPPARα fed with diet containing 0.1% Wy-14,643.

Fig. 5.

Northern blot analysis of PPARα target genes and cell cycle/apoptosis regulating genes using probes as indicated ( A ). Total RNA (10 µg/lane) was isolated from livers of non-cancerous tissue ( n = 5 animals) from each animal. The signal for acidic ribosomal phosphoprotein (36B4) was used as a control for loading and RNA integrity. Cont., control diet; Wy-14,643, control diet containing 0.1% Wy-14,643; ACOX, acyl-CoA oxidase; CYP4A, cytochrome P450 4A family; MCAD, medium chain acyl-CoA dehydrogenase; ME, malic enzyme; C MYC, c-myc; CD1, cyclin D1; CDK, cyclin-dependent kinase. Quantification analysis of gene expression (B, C and D). All values were normalized to the signal for 36B4. Results represent mean ± SD. ( B ) Results for PPARα target genes. *P < 0.05, **P < 0.01 versus corresponding to mice fed with control diet. ( C ) Results for cell cycle regulating genes. *P < 0.05, **P < 0.01 versus mPPARα fed with diet containing 0.1% Wy-14,643. ( D ) Results for apoptosis regulating genes. *P < 0.05, **P < 0.01 versus hPPARα fed with diet containing 0.1% Wy-14,643.

Discussion

Chronic dietary exposure of mice and rats to Wy-14,643 and other peroxisome proliferators typically results in hepatocellular neoplasia ( 5 , 79 ). In the present study, a high incidence of hepatocellular tumors (71%) including hepatocellular carcinomas was detected in mPPARα, fed with 0.1% Wy-14,643 for up to 38 weeks. Owing to the moribund status of mice in this group, Wy-14,643 treatment was prematurely terminated at 38 weeks from the beginning of the experiment. In sharp contrast, hPPARα mice given the same diet for up to 44 weeks, at which the wild-type mice exhibited a 100% incidence of liver tumors under the same treatment ( 5 ), demonstrated a very low incidence of hepatocellular tumors (5%). This observation represents conclusive evidence for low-susceptibility of this humanized mouse model to peroxisome proliferator-induced hepatocarcinogenesis.

In the presence of ligand, PPARα heterodimerizes with RXRα and binds to peroxisome proliferator response elements (PPREs) in the promoter region of target genes, resulting in increased transcription and expression of proteins and enzymes necessary for the transport and catabolism of fatty acids. A large number of genes including peroxisomal ( Acox ), microsomal ( Cyp4a ) and mitochondrial ( Mcad ) fatty acid oxidation genes, as well as fatty acid synthase ( Fas ) gene are well documented as PPARα target genes ( 25 ). In this study, the expression of most of these target genes were elevated in the Wy-14,643-treated hPPARα mouse, revealing that hPPARα was functional in control of fatty acid homeostasis similar to the wild-type mouse and that transcription was continuously stimulated throughout the course of ligand treatment.

Cyclins and CDKs regulate the cell cycle and overexpression of these proteins can results in uncontrolled cell proliferation. These proteins were previously found to be markedly upregulated in wild-type mice fed Wy-14,643 ( 26 ). Consistent with these findings, a number of cell cycle control genes were also upregulated in the mPPARα mouse group fed Wy-14,643, whereas the hPPARα mouse, under the same treatment regimen, did not show increased expression of these genes. These results indicate that genes associated with cell proliferation are preferentially activated in the livers of wild-type mice as compared with the hPPARα mouse line or that the severe toxicity and/or hepatocyte damage in the wild-type mouse results in induction of cell cycle control genes associated with the hepatocyte regenerative response. However, whether an increase in the expression of these genes is a cause or an effect of carcinogenesis is not elucidated in this study, because the total RNAs used in this analysis were derived from liver tissues of mice chronically exposed to Wy-14,643.

In response to the ligand activation, a number of lipid-metabolizing enzymes are induced in rodents. Increased levels of enzymes associated with the peroxisomal fatty acid β-oxidation system including acyl-CoA oxidase and to a lesser degree, the microsomal CYP4A subfamily of enzymes involved in fatty acids ω-oxidation leads to the generation of hydrogen peroxide (H 2 O 2 ) ( 27 ). Increased H 2 O 2 could potentially react with metals and generate highly reactive hydroxyl radicals, or react with lipid resulting in lipid peroxides ultimately elevating reactive oxygen species (ROS) and the degree of oxidative stress that can cause damage to DNA, protein, lipids and other cellular components that contribute to hepatocarcinogenesis in rodents ( 28 ). However, ROS may also act as a second intermediate in intracellular signaling ( 29 ) or inhibit the mitochondria respiratory chain complex leading to apoptosis ( 30 , 31 ). Although suppression of apoptosis has been implicated as a mechanism for hepatocarcinogenicity of peroxisome proliferators, these compounds can induce cell death in the human HepG2 cell line ( 32 ). A peroxisome proliferator, BR931, was also demonstrated to induce apoptosis through a p53-dependent pathway in the rat hepatoma FaO cell line ( 33 ). Overexpression of the p53 gene in Wy-14,643-treated hPPARα mice may imply that a sustained activation of the programmed cell death system contributes to the resistance to hepatocarcinogenesis, in contrast to the Wy-14,643-treated mPPARα mice group with comparatively lower expression of the p53 gene. However, the mutated form of this protein is retained in the cell, resulting an abnormal cell cycle. Besides, it should be noted that hepatoma cells lines are markedly different from hepatocytes and thus the results obtained with a rat hepatoma FaO cell line may not reflect the activity of peroxisome proliferators in vivo . Additional studies are necessary to establish a role for p53 in the resistance of the hPPARα mice to hepatocarcinogenesis.

It must be considered that hepatic levels of PPARα and/or different ligand affinity of receptor could account for the differential response in cell proliferation between mice and humans as revealed in the altered expression of cell cycle control genes involved in hepatocyte proliferation and carcinogenesis. Indeed, transient retroviral overexpression of the human PPARα in PPARα-null mice resulted in induction of some target genes accompanied by increased cell proliferation, indicating the higher level of human PPARα may contribute liver cell proliferation ( 19 ). A different ligand affinity for Wy-14,643 between human and mouse PPARα was also reported ( 34 ). However, the level of PPARα expression does not appear to be critical in this model, because the cellular content of human PPARα protein in the hPPARα mice detected in this study was similar to the mouse PPARα protein levels in wild-type mice. Furthermore, similar expression levels of several potent PPARα target genes such as ACOX and CYP4A were found comparing the wild-type and PPARα mice, indicating that the ligand affinity differences between human and mouse PPARα may not be important under the conditions used in these experiments. It also cannot be ruled out that the hPPARα mice are resistant to the hepatotoxic effects of peroxisome proliferators due to the site of expression of the human receptor. The cDNA was placed under control of the tetracycline regulatory system and the liver-specific Cebp /β promoter that is preferentially expressed in hepatocytes. Thus, it cannot be excluded that expression of PPARα in other cells in the liver such as Kupffer or stellate cells may be involved in the hepatoproliferative response. However, PPARα is not expressed at appreciable levels in Kupffer cells ( 35 ).

The rodent model is widely accepted as a useful tool for evaluating chemical carcinogenicity and/or toxicity, in accordance with the recommendation of the International Conference on Harmonization of Technical Requirements of Pharmaceuticals for Human Use (ICH). However, for assessment of the hazards of peroxisome proliferators, including hypolipidemic drugs, to human health, an explanation for the interspecies differences to those chemicals yields some important insights. Although the epidemiological studies on long-term treatment with hypolipidemic drugs showed inconclusive evidence of carcinogenic effects in humans ( 16 ), elucidation of the mechanisms by which peroxisome proliferators induce carcinogenesis is a prerequisite to accurately assess the human risk. In this regard, the hPPARα mouse model will provide a valuable model for determining the mechanism of the species differences in liver carcinogenesis and the mechanism of action of non-genotoxic hepatocarcinogens.

We thank Yatrik Shah for help and suggestions. This study was supported by the National Cancer Institute Intramural Research Program, an NFAFD contract to SoBran and by a National Institutes of Health Grant GM23750. CC was supported by a Wellcome Trust Research Fellowship 064866.

Conflict of Interest Statement : None declared.

References

1.
Peters,J.M., Cheung,C. and Gonzalez,F.J. (
2005
) Peroxisome proliferator-activated receptor-alpha and liver cancer: where do we stand?
J Mol Med.
  ,
83
,
774
–785.
2.
Reddy,J.K. and Krishnakantha,T.P. (
1975
) Hepatic peroxisome proliferation: induction by two novel compounds structurally unrelated to clofibrate.
Science
  ,
190
,
787
–789.
3.
Desvergne,B. and Wahli,W. (
1999
) Peroxisome proliferator-activated receptors: nuclear control of metabolism.
Endocr. Rev.
  ,
20
,
649
–688.
4.
Issemann,I. and Green,S. (
1990
) Activation of a member of the steroid hormone receptor superfamily by peroxisome proliferators.
Nature
  ,
347
,
645
–650.
5.
Peters,J.M., Cattley,R.C. and Gonzalez,F.J. (
1997
) Role of PPAR alpha in the mechanism of action of the nongenotoxic carcinogen and peroxisome proliferator Wy-14,643.
Carcinogenesis
  ,
18
,
2029
–2033.
6.
Reddy,J.K., Azarnoff,D.L. and Hignite,C.E. (
1980
) Hypolipidaemic hepatic peroxisome proliferators form a novel class of chemical carcinogens.
Nature
  ,
283
,
397
–398.
7.
Reddy,J.K. and Lalwani,N.D. (
1983
) Carcinogenesis by hepatic peroxisome proliferators: evaluation of the risk of hypolipidemic drugs and industrial plasticizers to humans.
Crit. Rev. Toxicol.
  ,
12
,
1
–58.
8.
Reddy,J.K., Rao,M.S., Azarnoff,D.L. and Sell,S. (
1979
) Mitogenic and carcinogenic effects of a hypolipidemic peroxisome proliferator, [4-chloro-6-(2,3-xylidino)-2-pyrimidinylthio]acetic acid (Wy-14,643), in rat and mouse liver.
Cancer Res.
  ,
39
,
152
–161.
9.
Reddy,J.K., Rao,S. and Moody,D.E. (
1976
) Hepatocellular carcinomas in acatalasemic mice treated with nafenopin, a hypolipidemic peroxisome proliferator.
Cancer Res.
  ,
36
,
1211
–1217.
10.
Lee,S.S., Pineau,T., Drago,J., Lee,E.J., Owens,J.W., Kroetz,D.L., Fernandez-Salguero,P.M., Westphal,H. and Gonzalez,F.J. (
1995
) Targeted disruption of the alpha isoform of the peroxisome proliferator-activated receptor gene in mice results in abolishment of the pleiotropic effects of peroxisome proliferators.
Mol. Cell. Biol.
  ,
15
,
3012
–3022.
11.
Bentley,P., Calder,I., Elcombe,C., Grasso,P., Stringer,D. and Wiegand,H.J. (
1993
) Hepatic peroxisome proliferation in rodents and its significance for humans.
Food Chem. Toxicol.
  ,
31
,
857
–907.
12.
Moody,D.E., Reddy,J.K., Lake,B.G., Popp,J.A. and Reese,D.H. (
1991
) Peroxisome proliferation and nongenotoxic carcinogenesis: commentary on a symposium.
Fundam. Appl. Toxicol.
  ,
16
,
233
–248.
13.
Ashby,J., Brady,A., Elcombe,C.R., Elliott,B.M., Ishmael,J., Odum,J., Tugwood,J.D., Kettle,S. and Purchase,I.F. (
1994
) Mechanistically-based human hazard assessment of peroxisome proliferator-induced hepatocarcinogenesis.
Hum. Exp. Toxicol.
  ,
13
(Suppl. 2),
S1
–S117.
14.
De La Iglesia,F.A., Lewis,J.E., Buchanan,R.A., Marcus,E.L. and McMahon,G. (
1982
) Light and electron microscopy of liver in hyperlipoproteinemic patients under long-term gemfibrozil treatment.
Atherosclerosis
  ,
43
,
19
–37.
15.
Hoivik,D.J., Qualls,C.W.,Jr, Mirabile,R.C. et al . (
2004
) Fibrates induce hepatic peroxisome and mitochondrial proliferation without overt evidence of cellular proliferation and oxidative stress in cynomolgus monkeys.
Carcinogenesis
  ,
25
,
1757
–1769.
16.
(
1984
) WHO cooperative trial on primary prevention of ischaemic heart disease with clofibrate to lower serum cholesterol: final mortality follow-up.
Lancet
  ,
2
,
600
–604.
17.
Mukherjee,R., Jow,L., Noonan,D. and McDonnell,D.P. (
1994
) Human and rat peroxisome proliferator activated receptors (PPARs) demonstrate similar tissue distribution but different responsiveness to PPAR activators.
J. Steroid Biochem. Mol. Biol.
  ,
51
,
157
–166.
18.
Sher,T., Yi,H.F., McBride,O.W. and Gonzalez,F.J. (
1993
) cDNA cloning, chromosomal mapping, and functional characterization of the human peroxisome proliferator activated receptor.
Biochemistry
  ,
32
,
5598
–5604.
19.
Yu,S., Cao,W.Q., Kashireddy,P. et al . (
2001
) Human peroxisome proliferator-activated receptor alpha (PPARalpha) supports the induction of peroxisome proliferation in PPARalpha-deficient mouse liver.
J. Biol. Chem.
  ,
276
,
42485
–42491.
20.
Cheung,C., Akiyama,T.E., Ward,J.M., Nicol,C.J., Feigenbaum,L., Vinson,C. and Gonzalez,F.J. (
2004
) Diminished hepatocellular proliferation in mice humanized for the nuclear receptor peroxisome proliferator-activated receptor alpha.
Cancer Res.
  ,
64
,
3849
–3854.
21.
Kistner,A., Gossen,M., Zimmermann,F., Jerecic,J., Ullmer,C., Lubbert,H. and Bujard,H. (
1996
) Doxycycline-mediated quantitative and tissue-specific control of gene expression in transgenic mice.
Proc. Natl Acad. Sci. USA
  ,
93
,
10933
–10938.
22.
In Frith,C.H., Ward,J.M. and Turusov,V.S. (eds) (
1994
) Tumours of The Liver , IARC Scientific Publication, Lyon.
23.
Akiyama,T.E., Ward,J.M. and Gonzalez,F.J. (
2000
) Regulation of the liver fatty acid-binding protein gene by hepatocyte nuclear factor 1alpha (HNF1alpha). Alterations in fatty acid homeostasis in HNF1alpha-deficient mice.
J. Biol. Chem.
  ,
275
,
27117
–27122.
24.
Aoyama,T., Peters,J.M., Iritani,N., Nakajima,T., Furihata,K., Hashimoto,T. and Gonzalez,F.J. (
1998
) Altered constitutive expression of fatty acid-metabolizing enzymes in mice lacking the peroxisome proliferator-activated receptor alpha (PPARalpha).
J. Biol. Chem.
  ,
273
,
5678
–5684.
25.
Mandard,S., Muller,M. and Kersten,S. (
2004
) Peroxisome proliferator-activated receptor alpha target genes.
Cell Mol. Life Sci.
  ,
61
,
393
–416.
26.
Peters,J.M., Aoyama,T., Cattley,R.C., Nobumitsu,U., Hashimoto,T. and Gonzalez,F.J. (
1998
) Role of peroxisome proliferator-activated receptor alpha in altered cell cycle regulation in mouse liver.
Carcinogenesis
  ,
19
,
1989
–1994.
27.
Yeldandi,A.V., Rao,M.S. and Reddy,J.K. (
2000
) Hydrogen peroxide generation in peroxisome proliferator-induced oncogenesis.
Mutat Res.
  ,
448
,
159
–177.
28.
Rao,M.S. and Reddy,J.K. (
1996
) Hepatocarcinogenesis of peroxisome proliferators.
Ann. NY Acad. Sci.
  ,
804
,
573
–587.
29.
Dalton,T.P., Shertzer,H.G. and Puga,A. (
1999
) Regulation of gene expression by reactive oxygen.
Annu. Rev. Pharmacol. Toxicol.
  ,
39
,
67
–101.
30.
Cohen,G. and Heikkila,R.E. (
1978
) Mechanisms of action of hydroxylated phenylethylamine and indoleamine neurotoxins.
Ann. NY Acad. Sci.
  ,
305
,
74
–84.
31.
Glinka,Y., Tipton,K.F. and Youdim,M.B. (
1996
) Nature of inhibition of mitochondrial respiratory complex I by 6-Hydroxydopamine.
J. Neurochem.
  ,
66
,
2004
–2010.
32.
Jiao,H.L. and Zhao,B.L. (
2002
) Cytotoxic effect of peroxisome proliferator fenofibrate on human HepG2 hepatoma cell line and relevant mechanisms.
Toxicol. Appl. Pharmacol.
  ,
185
,
172
–179.
33.
Simbula,G., Pibiri,M., Sanna,L., Cossu,C., Molotzu,F., Columbano,A. and Ledda-Columbano,G.M. (
2004
) The peroxisome proliferator BR931 kills FaO cells by p53-dependent apoptosis.
Life Sci.
  ,
75
,
271
–286.
34.
Keller,H., Devchand,P.R., Perroud,M. and Wahli,W. (
1997
) PPAR alpha structure–function relationships derived from species-specific differences in responsiveness to hypolipidemic agents.
Biol. Chem.
  ,
378
,
651
–655.
35.
Peters,J.M., Rusyn,I., Rose,M.L., Gonzalez,F.J. and Thurman,R.G. (
2000
) Peroxisome proliferator-activated receptor alpha is restricted to hepatic parenchymal cells, not Kupffer cells: implications for the mechanism of action of peroxisome proliferators in hepatocarcinogenesis.
Carcinogenesis
  ,
21
,
823
–826.

Author notes

Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute, 1Comparative Medicine Branch, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD 20892, USA and 2Department of Pathology, Northwestern University School of Medicine, Chicago, IL 60611, USA