Defect in Peroxisome Proliferator-activated Receptor α-inducible Fatty Acid Oxidation Determines the Severity of Hepatic Steatosis in Response to Fasting
2000; Elsevier BV; Volume: 275; Issue: 37 Linguagem: Inglês
10.1074/jbc.m910350199
ISSN1083-351X
AutoresTakashi Hashimoto, William S. Cook, Chao Qi, Anjana V. Yeldandi, Janardan K. Reddy, Mahendra S. Rao,
Tópico(s)Metabolism and Genetic Disorders
ResumoFasting causes lipolysis in adipose tissue leading to the release of large quantities of free fatty acids into circulation that reach the liver where they are metabolized to generate ketone bodies to serve as fuels for other tissues. Since fatty acid-metabolizing enzymes in the liver are transcriptionally regulated by peroxisome proliferator-activated receptor α (PPARα), we investigated the role of PPARα in the induction of these enzymes in response to fasting and their relationship to the development of hepatic steatosis in mice deficient in PPARα (PPARα−/−), peroxisomal fatty acyl-CoA oxidase (AOX−/−), and in both PPARα and AOX (double knock-out (DKO)). Fasting for 48–72 h caused profound impairment of fatty acid oxidation in both PPARα−/− and DKO mice, and DKO mice revealed a greater degree of hepatic steatosis when compared with PPARα−/− mice. The absence of PPARα in both PPARα−/− and DKO mice impairs the induction of mitochondrial β-oxidation in liver following fasting which contributes to hypoketonemia and hepatic steatosis. Pronounced steatosis in DKO mouse livers is due to the added deficiency of peroxisomal β-oxidation system in these animals due to the absence of AOX. In mice deficient in AOX alone, the sustained hyperactivation of PPARα and up-regulation of mitochondrial β-oxidation and microsomal ω-oxidation systems as well as the regenerative nature of a majority of hepatocytes containing numerous spontaneously proliferated peroxisomes, which appear refractory to store triglycerides, blunt the steatotic response to fasting. Starvation for 72 h caused a decrease in PPARα hepatic mRNA levels in wild type mice, with no perceptible compensatory increases in PPARγ and PPARδ mRNA levels. PPARγ and PPARδ hepatic mRNA levels were lower in fed PPARα−/− and DKO mice when compared with wild type mice, and fasting caused a slight increase only in PPARγ levels and a decrease in PPARδ levels. Fasting did not change the PPAR isoform levels in AOX−/− mouse liver. These observations point to the critical importance of PPARα in the transcriptional regulatory responses to fasting and in determining the severity of hepatic steatosis. Fasting causes lipolysis in adipose tissue leading to the release of large quantities of free fatty acids into circulation that reach the liver where they are metabolized to generate ketone bodies to serve as fuels for other tissues. Since fatty acid-metabolizing enzymes in the liver are transcriptionally regulated by peroxisome proliferator-activated receptor α (PPARα), we investigated the role of PPARα in the induction of these enzymes in response to fasting and their relationship to the development of hepatic steatosis in mice deficient in PPARα (PPARα−/−), peroxisomal fatty acyl-CoA oxidase (AOX−/−), and in both PPARα and AOX (double knock-out (DKO)). Fasting for 48–72 h caused profound impairment of fatty acid oxidation in both PPARα−/− and DKO mice, and DKO mice revealed a greater degree of hepatic steatosis when compared with PPARα−/− mice. The absence of PPARα in both PPARα−/− and DKO mice impairs the induction of mitochondrial β-oxidation in liver following fasting which contributes to hypoketonemia and hepatic steatosis. Pronounced steatosis in DKO mouse livers is due to the added deficiency of peroxisomal β-oxidation system in these animals due to the absence of AOX. In mice deficient in AOX alone, the sustained hyperactivation of PPARα and up-regulation of mitochondrial β-oxidation and microsomal ω-oxidation systems as well as the regenerative nature of a majority of hepatocytes containing numerous spontaneously proliferated peroxisomes, which appear refractory to store triglycerides, blunt the steatotic response to fasting. Starvation for 72 h caused a decrease in PPARα hepatic mRNA levels in wild type mice, with no perceptible compensatory increases in PPARγ and PPARδ mRNA levels. PPARγ and PPARδ hepatic mRNA levels were lower in fed PPARα−/− and DKO mice when compared with wild type mice, and fasting caused a slight increase only in PPARγ levels and a decrease in PPARδ levels. Fasting did not change the PPAR isoform levels in AOX−/− mouse liver. These observations point to the critical importance of PPARα in the transcriptional regulatory responses to fasting and in determining the severity of hepatic steatosis. triacylglycerols straight chain fatty acyl-CoA oxidase carnitine acetyltransferase carnitine octanoyltransferase encode microsomal cytochrome P450 fatty acid ω-hydroxylases double knock-out nullizygous for both PPARα and AOX free fatty acids 3-hydroxyacyl-CoA dehydrogenase 3-hydroxybutyrate 3-hydroxy-3 methylglutaryl-CoA synthase medium chain acyl-CoA dehydrogenase mitochondrial 3-ketoacyl-CoA thiolase mitochondrial trifunctional protein mitochondrial acetoacetyl-CoA-specific thiolase peroxisomal d-3-hydroxyacyl-CoA dehydratase/d-3-hydroxyacyl-CoA dehydrogenase bifunctional protein peroxisomal enoyl-CoA hydratase/3-hydroxyacyl-CoA dehydrogenase bifunctional protein peroxisome proliferator-activated receptor α, β/δ or γ peroxisomal 3-ketoacyl-CoA thiolase short chain acyl-CoA dehydrogenase succinyl-CoA: 3-oxoacid transferase very long chain acyl-CoA synthetase very long chain acyl-CoA dehydrogenase Higher animals, under fed conditions, preferentially burn carbohydrate to generate ATP, and surplus carbohydrate is converted into fatty acids, which are then stored as triacylglycerols (TG)1 in adipose tissue. When glucose availability is low during periods of starvation, the TG stored in adipose tissue are hydrolyzed to free fatty acids (FFA) and mobilized into plasma to reach liver where they play a major role in energy production (1Seitz H.J. Mullerm M.J. Krone E. Tarnowski W. Arch. Biochem. Biophys. 1977; 183: 647-663Crossref PubMed Scopus (68) Google Scholar, 2Eaton S. Bartlett K. Pourfarazam M. Biochem. J. 1996; 320: 345-357Crossref PubMed Scopus (351) Google Scholar, 3Roe C.R. Coates P.M. Scriver C.R. Beaudet A.I. Sly W.S. Valle D. Metabolic and Molecular Absence of Inherited Disease. McGraw-Hill Inc., New York1995: 1501-1533Google Scholar). In liver, the influxed fatty acids are oxidized predominantly by the mitochondrial β-oxidation system and to a lesser extent by the peroxisomal β-oxidation, as well as by CYP4A-catalyzed microsomal ω-oxidation pathways (4Hashimoto T. Neurochem. Res. 1999; 24: 551-563Crossref PubMed Scopus (61) Google Scholar, 5Aoyama T. Hardwick J.P. Imaoka S. 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Neurol. 1996; 9: 477-485Crossref PubMed Scopus (28) Google Scholar). However, during conditions that lead to short term fasting, they manifest severe hypoketotic hypoglycemia, increased plasma FFA, variable degree of hepatic steatosis, and sudden death in early life because of their inability to oxidize FFAs in liver due to enzymatic defects (8Taroni F. Uziel G. Curr. Opin. Neurol. 1996; 9: 477-485Crossref PubMed Scopus (28) Google Scholar). These metabolic diseases have provided valuable insights pertaining to the role of individual enzymes in fatty acid oxidation and energy utilization. It is now well recognized that fasting causes a rapid transcriptional activation of genes encoding mitochondrial, peroxisomal, and microsomal fatty acid oxidation in liver in healthy individuals (9Nagao M. Parimoo B. Tanaka K. J. Biol. Chem. 1993; 268: 24114-24124Abstract Full Text PDF PubMed Google Scholar, 10Hillgartner F.B. Salati L.M. Goodridge A.G. Physiol. 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Sci. 1996; 804: 176-201Crossref PubMed Scopus (100) Google Scholar), consist of three isotypes, namely PPARα, PPARδ (also called PPARβ), and PPARγ which are products of separate genes (11Issemann I. Green S. Nature. 1990; 347: 645-650Crossref PubMed Scopus (3059) Google Scholar, 13Zhu Y. Alvares K. Huang Q. Rao M.S. Reddy J.K. J. Biol. Chem. 1993; 268: 26817-26820Abstract Full Text PDF PubMed Google Scholar, 14Dryer C. Krey G. Keller H. Givel F. Helftenbein G. Wahli W. Cell. 1992; 68: 879-887Abstract Full Text PDF PubMed Scopus (1214) Google Scholar). Peroxisome proliferators are structurally diverse agents which, when administered to rats and mice, induce not only a marked peroxisome proliferation and increase in the enzyme proteins of the peroxisomal fatty acid oxidation but also induce changes in carbohydrate and lipid metabolisms (4Hashimoto T. Neurochem. Res. 1999; 24: 551-563Crossref PubMed Scopus (61) Google Scholar, 12Reddy J.K. Chu R. Ann. N. Y. Acad. Sci. 1996; 804: 176-201Crossref PubMed Scopus (100) Google Scholar). The induction of mitochondrial, peroxisomal, and microsomalCYP4A genes involved in fatty acid oxidation requires the formation of PPARα heterodimerization with retinoid X receptor, and this PPARα·retinoid X receptor complex binds to PPAR response element, a region consisting of a degenerate direct repeat of the canonical AGGTCA sequence separated by 1 base pair (DR1), present in the 5 ′-flanking region of target genes (15Kliewer S.A. Umesono K. Noonan D.J. Heyman R.A. Evans R.M. Nature. 1992; 358: 771-774Crossref PubMed Scopus (1525) Google Scholar). The generation of PPARα−/− mice established that PPARα is critical for peroxisome proliferation and the coordinate transcriptional activation of fatty acid oxidation enzymes in liver (16Lee S.-S.-T. Pineau T. Drago J. Lee E.J. Owens J.W. Kroetz D.L. Fernandez-Salguero P.M. Westpal H. Gonzalez F.J. Mol. Cell. Biol. 1995; 15: 3012-3022Crossref PubMed Scopus (1506) Google Scholar). Furthermore, PPARα−/− mice have provided valuable information on the constitutive levels of expression of mitochondrial and peroxisomal fatty acid-metabolizing enzymes in liver (17Aoyama T. Peters J.M. Iritani N. Nakajima T. Furihata K. Hashimoto T. Gonzalez F.J. J. Biol. Chem. 1998; 273: 5678-5684Abstract Full Text Full Text PDF PubMed Scopus (754) Google Scholar) and the response of these mice to dietary overload as well as short term fasting (18Kroetz D.L. Yook P. Costet P. Bianchi P. Pineau T. J. Biol. Chem. 1998; 273: 31581-31589Abstract Full Text Full Text PDF PubMed Scopus (190) Google Scholar, 19Kersten S. Seydoux J. Peters J.M. Gonzalez F.J. Desvergne B. Wahli W. J. Clin. Invest. 1999; 103: 1489-1498Crossref PubMed Scopus (1371) Google Scholar, 20Leone T.C. Weinheimer C.J. Kelly D.P. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 7473-7478Crossref PubMed Scopus (824) Google Scholar). Mice deficient in peroxisomal fatty acyl-CoA oxidase (AOX−/−) exhibited sustained PPARα hyperfunction presumably caused by accumulation of endogenous ligand(s) due to the impairment of the peroxisomal fatty acid oxidation pathway (21Fan C.-Y. Pan J. Usuda N. Yeldandi A.V. Rao M.S. Reddy J.K. J. Biol. Chem. 1998; 273: 15639-15645Abstract Full Text Full Text PDF PubMed Scopus (312) Google Scholar). Mice nullizygous for both PPARα and AOX (PPARα−/− AOX−/−double nulls (DKO)) have also served as valuable tools to explore the role of PPARα and fatty acid oxidation in constitutive lipid metabolism and hepatic fatty liver phenotype under fed state (22Hashimoto T. Fujita T. Usuda N. Cook W. Qi C. Peters J.M. Gonzalez F.J. Yeldandi A.V. Rao M.S. Reddy J.K. J. Biol. Chem. 1999; 274: 19228-19236Abstract Full Text Full Text PDF PubMed Scopus (215) Google Scholar). The availability of these genetically altered PPARα−/−(16Lee S.-S.-T. Pineau T. Drago J. Lee E.J. Owens J.W. Kroetz D.L. Fernandez-Salguero P.M. Westpal H. Gonzalez F.J. Mol. Cell. Biol. 1995; 15: 3012-3022Crossref PubMed Scopus (1506) Google Scholar), PPARα−/− AOX−/− (DKO) (22Hashimoto T. Fujita T. Usuda N. Cook W. Qi C. Peters J.M. Gonzalez F.J. Yeldandi A.V. Rao M.S. Reddy J.K. J. Biol. Chem. 1999; 274: 19228-19236Abstract Full Text Full Text PDF PubMed Scopus (215) Google Scholar), and AOX−/− (21Fan C.-Y. Pan J. Usuda N. Yeldandi A.V. Rao M.S. Reddy J.K. J. Biol. Chem. 1998; 273: 15639-15645Abstract Full Text Full Text PDF PubMed Scopus (312) Google Scholar) mice provides an opportunity to examine the comparative responses to changes in energy metabolism imposed by fasting. We demonstrate the critical importance of PPARα-dependent induction of fatty acid oxidation in determining the degree of hepatic steatosis. Wild type (C57BL/6J), AOX-null (AOX−/−) (21Fan C.-Y. Pan J. Usuda N. Yeldandi A.V. Rao M.S. Reddy J.K. J. Biol. Chem. 1998; 273: 15639-15645Abstract Full Text Full Text PDF PubMed Scopus (312) Google Scholar), PPARα-null (PPARα−/−) (16Lee S.-S.-T. Pineau T. Drago J. Lee E.J. Owens J.W. Kroetz D.L. Fernandez-Salguero P.M. Westpal H. Gonzalez F.J. Mol. Cell. Biol. 1995; 15: 3012-3022Crossref PubMed Scopus (1506) Google Scholar), and AOX−/− PPARα−/− double knock-out (DKO) (22Hashimoto T. Fujita T. Usuda N. Cook W. Qi C. Peters J.M. Gonzalez F.J. Yeldandi A.V. Rao M.S. Reddy J.K. J. Biol. Chem. 1999; 274: 19228-19236Abstract Full Text Full Text PDF PubMed Scopus (215) Google Scholar) mice were housed in a controlled environment with a 12-h light/dark cycle with free access to water and standard laboratory chow as described (22Hashimoto T. Fujita T. Usuda N. Cook W. Qi C. Peters J.M. Gonzalez F.J. Yeldandi A.V. Rao M.S. Reddy J.K. J. Biol. Chem. 1999; 274: 19228-19236Abstract Full Text Full Text PDF PubMed Scopus (215) Google Scholar). All experiments were performed using mice ranging in age from 16 to 20 weeks. Starvation was commenced by removing food at 8:00 a.m., and groups of mice were fasted up to 96 h. Control mice were fed ad libitum. After mice were anesthetized, blood was collected in heparinized tubes and centrifuged, and the plasma was frozen until use. Organs were removed and frozen in liquid nitrogen and stored at −80 °C. All animal procedures used in this study were reviewed and preapproved by the Institutional Review Boards for Animal Research of the Northwestern University. For light microscopy, pieces of liver were fixed in 10% neutral buffered formalin, embedded in paraffin, and 4-μm-thick sections stained with hematoxylin and eosin. Frozen sections of formalin-fixed liver (5 μm) were stained with Oil Red O and counterstained with Giemsa. For cell proliferation analysis, mice were given bromodeoxyuridine (0.5 mg/ml) in drinking water, and their livers were processed for immunohistochemical localization as described previously (23Qi C. Zhu Y. Pan J. Yeldandi A.V. Rao M.S. Maeda N. Subbarao V. Pulikuri S. Hashimoto T. Reddy J.K. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 1585-1590Crossref PubMed Scopus (70) Google Scholar), using antibodies raised against bromodeoxyuridine (Becton Dickinson). Histological analysis and image processing were carried out using Leica DMRE microscope equipped with a Spot digital camera. Images were taken at × 20 and 40 magnification and captured at 1315 × 1033 pixels. Montages of images were prepared with the use of Photoshop 5.0 (Adobe, Mountain View, CA). Plasma glucose (24Bergmeyer H.U. Bernt E. Bergmeyer H.U. Methods of Enzymatic Analysis. 2nd English Ed. Academic Press, Inc., New York1974: 1205-1215Google Scholar), lactate (25Gutmann I. Wahlefeld A.W. Bergmeyer H.U. Methods of Enzymatic Analysis. 2nd English Ed. Academic Press, Inc., New York1974: 1464-1468Google Scholar), and 3-hydroxybutyrate (3-HB) (26Williamson D.H. Mellanby J. Bergmeyer H.U. Methods of Enzymatic Analysis. 2nd English Ed. Academic Press, Inc., New York1974: 1837-1839Google Scholar) were determined by the cited procedures. Plasma FFA and TG were determined by the use of reagent kits (NEFA C-Test Wako and Triglyceride E-Test Wako, respectively, from Wako Pure Chemical Industries, Ltd. Osaka, Japan). Liver glycogen was determined by the use of coupling reactions of amyloglucosidase and glucose oxidase (27Roehrig K.L. Allred J.B. Anal. Biochem. 1974; 58: 414-421Crossref PubMed Scopus (221) Google Scholar). Total carnitines were determined using carnitine acetyltransferase (CAT) (28Pearson D.J. Tubbs P.K. Chase J.F.A. Bergmeyer H.U. Methods of Enzymatic Analysis. 2nd English Ed. Academic Press, Inc., New York.1974: 1758-1771Crossref Google Scholar). Protein concentrations were determined using a protein assay kit (Bio-Rad) using bovine serum albumin as standard. Liver, kidney, and heart extracts were subjected to 10% SDS-polyacrylamide gel electrophoresis and transferred to nitrocellulose membranes. The membranes were incubated with the primary antibody (see Refs. 17Aoyama T. Peters J.M. Iritani N. Nakajima T. Furihata K. Hashimoto T. Gonzalez F.J. J. Biol. Chem. 1998; 273: 5678-5684Abstract Full Text Full Text PDF PubMed Scopus (754) Google Scholar and 22Hashimoto T. Fujita T. Usuda N. Cook W. Qi C. Peters J.M. Gonzalez F.J. Yeldandi A.V. Rao M.S. Reddy J.K. J. Biol. Chem. 1999; 274: 19228-19236Abstract Full Text Full Text PDF PubMed Scopus (215) Google Scholarfor the source of various primary antibodies used in this study) followed by alkaline phosphatase-conjugated goat anti-rabbit IgG. Antibodies against 3-hydroxy-3-methylglutaryl-CoA synthase (HS), 3-hydroxy-3-methylglutaryl-CoA lyase (29Boukaftane Y. Mitchell G.A. Gene (Amst.). 1997; 195: 121-126Crossref PubMed Scopus (13) Google Scholar), and succinyl-CoA:oxoacid transferase (SCOT) (30Song X.Q. Fukao T. Mitchell G.A. Kassouska-Bratinova S. Ugarte M. Wanders R.J. Hirayama M.K. Shintaku H. Churchill P. Watanabe H. Orii T. Kondo N. Biochim. Biophys. Acta. 1997; 12: 151-156Crossref Scopus (21) Google Scholar) were provided by Dr. G. A. Mitchell and Dr. T. Fukao, respectively. The Western blot signals were quantified by scanning densitometry, and the values from mice fed control diets were assigned the number 1.0. Results are expressed as the means ± S.D. of three determinations. Total RNA was isolated from liver using the acid guanidinium thiocyanate/phenol/chloroform extraction method. RNA was glyoxylated, electrophoresed, transferred to a nylon membrane, and then hybridized at 42o in 50% formamide hybridization solution using32P-labeled cDNA probes as described previously (21Fan C.-Y. Pan J. Usuda N. Yeldandi A.V. Rao M.S. Reddy J.K. J. Biol. Chem. 1998; 273: 15639-15645Abstract Full Text Full Text PDF PubMed Scopus (312) Google Scholar,22Hashimoto T. Fujita T. Usuda N. Cook W. Qi C. Peters J.M. Gonzalez F.J. Yeldandi A.V. Rao M.S. Reddy J.K. J. Biol. Chem. 1999; 274: 19228-19236Abstract Full Text Full Text PDF PubMed Scopus (215) Google Scholar). Equal loading was verified by the intensity of methylene blue-stained 18 S and 28 S RNA or by probing the blots with 18 S RNA probe. Changes in mRNA levels were estimated by densitometric scanning of autoradiograms. RNase protection assay was performed using the following gene-specific probes: PPARα, nucleotides 1186–1565 (GenBankTMaccession number X57638) (16Lee S.-S.-T. Pineau T. Drago J. Lee E.J. Owens J.W. Kroetz D.L. Fernandez-Salguero P.M. Westpal H. Gonzalez F.J. Mol. Cell. Biol. 1995; 15: 3012-3022Crossref PubMed Scopus (1506) Google Scholar); PPARγ, nucleotides 1597–1914 (GenBankTM accession number U01841) (39Martin G Schoonjans K. Lefebvre A.M. Staels B. Auwerx J. J. Biol. Chem. 1997; 272: 28210-28217Abstract Full Text Full Text PDF PubMed Scopus (479) Google Scholar); PPARδ, nucleotides 1004–1268 (GenBankTM accession number U10375) (22Hashimoto T. Fujita T. Usuda N. Cook W. Qi C. Peters J.M. Gonzalez F.J. Yeldandi A.V. Rao M.S. Reddy J.K. J. Biol. Chem. 1999; 274: 19228-19236Abstract Full Text Full Text PDF PubMed Scopus (215) Google Scholar); and CYP4A1, nucleotides 1421–1555 (referred to as CYP4A10, GenBankTM accession number ABO18421). Antisense RNA probes were transcribed in the presence of [32P]UTP (20 mCi/ml, 800 Ci/mmol, Amersham Pharmacia Biotech) using the MAXIscript in vitro transcription kit (Ambion, Austin, TX). After transcription, the labeled riboprobes were purified in a 5% TBE/urea polyacrylamide Ready Gel (Bio-Rad). Probes were eluted from the polyacrylamide gel fragments, and their activity was measured in a scintillation counter. Total RNA isolated from liver was hybridized with labeled probes overnight and then digested for 30 min with RNase A/RNase T1 mix at 37 °C. Protected fragments were precipitated and resuspended in 3 μl of gel loading buffer. The samples were loaded onto a 6% polyacrylamide sequencing gel 0.4 mm in thickness (Bio-Rad). After electrophoresis, the gel was dried and exposed to film or a PhosphorImager plate (Molecular Dynamics, Amersham Pharmacia Biotech) overnight at room temperature without intensification. Quantitation was with a Molecular Dynamics Storm 860 PhosphorImager. Statistical comparisons were made by using Student's t test or two-way analysis of variance. A statistically significant difference was defined asp < 0.05. Since fasting increases the capacity for fatty acid oxidation in liver under normal conditions, we subjected wild type, PPARα−/−, DKO, and AOX−/− mice to fasting for up to 96 h for a comparative analysis of liver morphology. After 48 h starvation, the livers of PPARα−/− and DKO mice were paler compared with those of fasted wild type mice, and this difference in pallor indicative of severe steatosis was grossly exaggerated with prolonged fasting. A representative example of a typical gross appearance of liver of a fed and 66-h fasted DKO mouse is illustrated in Fig.1 A. Comparative histologic appearance of liver of fed and fasted wild type, PPARα−/−, and DKO mice, as revealed by Oil Red O staining (to visualize neutral lipid) of frozen sections, is illustrated in Fig. 1. In fed wild type mice there is no detectable fatty change in hepatocytes other than the presence of Oil Red O-positive droplets in stellate cells (Fig. lB). When fasted for 48–72 h, these wild type mice exhibited subtle steatosis in centrizonal hepatocytes (Fig. 1 C,arrows). As reported elsewhere (22Hashimoto T. Fujita T. Usuda N. Cook W. Qi C. Peters J.M. Gonzalez F.J. Yeldandi A.V. Rao M.S. Reddy J.K. J. Biol. Chem. 1999; 274: 19228-19236Abstract Full Text Full Text PDF PubMed Scopus (215) Google Scholar), under fed state, only a few centrilobular hepatocytes in PPARα−/−mice and few scattered periportal hepatocytes in DKO mice revealed fatty change (Fig. 1, D and F). When these mice were fasted for 48–72 h, steatosis was extensive involving the entire liver lobule (Fig. l, E and G). At 48, 66, or 72 h of fasting, the Oil Red O staining clearly showed marked differences in the degree of hepatic steatosis between PPARα−/− and DKO mice in that fatty change appeared more prominent in DKO livers. Oil Red O-stained liver sections of 48–72 h fasted AOX−/− mice revealed fatty change in centrizonal hepatocytes, but a majority of hepatocytes with intense eosinophilic cytoplasm indicative of regeneration did not accumulate fat (Oil Red O stain not illustrated but see hematoxylin and eosin staining pattern in Fig. 2 G). Examination of hematoxylin and eosin-stained histologic sections of livers from 66-h fasted wild type, PPARα−/−, and DKO mice demonstrated clear-cut differences in steatosis (Fig. 2,A–C). In wild type animals fasted for 66 or 72 h the fatty change was minimal with few microvesicular lipid droplets (Fig.2 A). In contrast, microvesicular fatty change appeared prominent in PPARα−/− livers (Fig. 2 B), and this change was greatly exaggerated in DKO livers in which many hepatocytes also had macrovesicular lipid droplets (Fig.2 C). When fasted for 96-h, wild type mice exhibited no fatty change, and in fact the liver cells revealed features of cytoplasmic atrophy (Fig. 2 D). On the other hand, the fatty change persisted and progressed predominantly to macrovesicular type in the midzonal and centrilobular areas of liver lobules, while maintaining the microvesicular steatotic pattern in periportal areas of both PPARα−/− and DKO mice (Fig. 2, E andF). The degree of hepatic steatosis appeared slightly more prominent in females during the first 48 h of starvation but with prolonged starvation (66–96 h); the differences in fatty change between males and females were not apparent. We also subjected AOX−/− mice to 48- and 72-h starvation and found that regenerated hepatocytes with eosinophilic cytoplasm are resistant to lipid accumulation, whereas cells already steatotic appeared nearly the same or only slightly more steatotic (Fig. 2 G). In order to demonstrate that the eosinophilic hepatocytes that do not exhibit steatosis in response to fasting are indeed cells that have regenerated and therefore resistant, we administered bromodeoxyuridine for 4 days in drinking water and assessed its incorporation in hepatocyte nuclei by immunoperoxidase staining (Fig. 2 H). During the 4-day labeling period, several hepatocytes have incorporated this precursor indicating DNA synthesis and cell proliferation in cells that are at the interface between steatotic cells and cells with abundant eosinophilic cytoplasm that are resistant to fatty change. In contrast, an occasional cell showed bromodeoxyuridine incorporation in the livers of either fed or starved wild type, PPARα−/−, and DKO mice (Fig. 2 I) indicating minimal cell proliferation.Figure 2Comparative liver morphology as revealed in sections stained with hematoxylin and eosin. A–C (all taken at the same magnification) represent livers of wild type (A), PPARα−/−(B), and DKO (C) mice fasted for 66 h. The inset in B depicts an enlarged version of cells with microvesicular fatty change in PPARα−/− mouse liver. D–F (all taken the same magnification) represent livers of wild type (D), PPARα−/− (E), and DKO (F) mice fasted for 96 h. In D–F the insetsrepresent higher magnification of areas marked withasterisks to show the absence of fatty change in wild type mouse liver starved for 96 h (D), the presence of steatosis in all cells in PPARα−/−(E), and DKO (F) liver. G, liver of AOX−/− mouse starved for 72 h. Note the absence of fatty starvation-induced steatosis in regenerated hepatocytes that contain eosinophilic cytoplasm; hepatocytes in the centrizonal area show microvesicular fatty change similar to that seen in fed state.H, liver of AOX−/− mouse showing a leading edge of hepatocellular proliferation, at the steatotic cell interface, as evidenced by bromodeoxyuridine labeling. No significant liver cell proliferation was noted in wild type, PPAR, and DKO (I) mice. The insets in H and I show immunoperoxidase-stained nuclei representing bromodeoxyuridine labeling in proliferated cells. Eventually all steatotic cells will be replaced by regenerated liver cells in AOX−/− mouse liver.View Large Image Figure ViewerDownload (PPT) Plasma glucose levels were not much different among the four groups of mice under fed conditions, and these levels were decreased by about 50% at 48 h of starvation, and similar levels were maintained at 72 h in all groups (Fig. 3 A). Plasma lactate levels in all groups were nearly the same under non-starved conditions, and on fasting gradual decreases in lactate concentrations were observed in all four groups of animals (data not shown). Lactate, together with alanine, serves as a major precursor for gluconeogenesis in liver under starvation. Glycogen in liver, a reservoir of glucose, was largely depleted within 48 h of starvation, and this reduction was sus
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