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Effects of Long-Chain Fatty Acids on Lipid-Metabolizing Genes and High-Density Lipoprotein Cholesterol Production in Cul...

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PAGE 1

EFFECTS OF LONG-CHAIN FATTY AC IDS ON LIPID METABOLIZING GENES AND HIGH-DENSITY LIPOPROTEIN CHOLESTEROL PRODUCTION IN CULTURED HUMAN AND RAT HEPATOCYTES By ELIZABETH SARAH GREENE A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2006

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Copyright 2006 by Elizabeth Sarah Greene

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To my parents, Dave and Hilary Johnson, fo r instilling in me a love of learning and supporting me through my seemingly never-ending quest for knowledge.

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iv ACKNOWLEDGMENTS I would like to thank my supervisory co mmittee chair, Dr. Lokenga Badinga, for allowing me the opportunity to be a part of hi s laboratory. I am grat eful for his support and advice in helping me become a better res earcher. I would also like to thank my committee members, Dr. Joel Brendem uhl, Dr. Bobbi Langkamp-Henken and Dr. Charles Staples for their guidance and dedication to my education. I would also like to than k the other members of the laboratory, Carlos Rodriguez-Sallaberry and Cristina CaldariTorres for many important problem-solving and stress-relieving conversati ons during morning coffee breaks. Special thanks go to Teri Woodham for being one of the best frie nds anyone could ever ask for. Additionally, I could not have succeeded wit hout all of the friends I made during the last four years. Finally, I thank my husband, Nic, for his love patience, strength, and never-ending faith in my abilities. His belief in me made everything possible.

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v TABLE OF CONTENTS page ACKNOWLEDGMENTS.................................................................................................iv LIST OF TABLES...........................................................................................................viii LIST OF FIGURES.............................................................................................................x ABBREVIATIONS KEY................................................................................................xiv ABSTRACT....................................................................................................................xvi i CHAPTER 1 INTRODUCTION........................................................................................................1 2 LITERATURE REVIEW.............................................................................................5 Structure and Metabolism of Lipids.............................................................................5 Structure and Nomenclature of Lipids..................................................................5 Biosynthesis of Fatty Acids...................................................................................7 Degradation of Fatty Acids.................................................................................10 Nutritional and Biological Properties of the Polyuns aturated Fatty Acids.................13 Dietary Requirements of th e Essential Fatty Acids.............................................13 Long-Chain Polyunsaturated Fatt y Acids of the n-6 Family...............................16 Long-Chain Polyunsaturated Fatt y Acids of the n-3 Family...............................17 Digestion and Assimila tion of Dietary Fats................................................................17 Dietary Fats in Relation to Health..............................................................................24 Dietary Fats in Relati on to Weight Control.........................................................24 Dietary Fats and Blood Cholesterol....................................................................26 Dietary Fats and Cardiovascular Disease............................................................28 Conjugated Linoleic Acid...........................................................................................32 Roles of the Peroxisome Proliferator-Ac tivated Receptors in Lipid Metabolism......38 PPAR .................................................................................................................40 PPAR / ..............................................................................................................42 PPAR .................................................................................................................44

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vi 3 EFFECTS OF N-3 AND N-6 FATT Y ACIDS ON LIPI D METABOLIZING GENES AND HIGH-DENSITY LIPOPROTEIN CHOLESTEROL PRODUCTION IN CULTURED HU MAN AND RAT HEPATOCYTES...............47 Introduction.................................................................................................................47 Materials and Methods...............................................................................................48 Materials..............................................................................................................48 Cell Culture and Treatment.................................................................................49 RNA Isolation and Analysis................................................................................50 Lipid Extraction...................................................................................................51 HDL Cholesterol Assay.......................................................................................51 Statistical Analysis..............................................................................................52 Results........................................................................................................................ .52 Effects of Fatty Acids on HepG2 Cells...............................................................52 Effects of Fatty Acids on H-4-II-E Cells.............................................................53 Role of PPAR in Stearic Acid-Induced Effects on Gene Expression...............53 Discussion...................................................................................................................54 Summary.....................................................................................................................58 4 EFFECTS OF ISOMERS OF CONJ UGATED LINOLEIC ACID ON LIPID METABOLIZING GENES AND HIGH-DENSITY LIPOPROTEIN CHOLESTEROL PRODUCTION IN CULTURED HUMAN AND RAT HEPATOCYTES........................................................................................................80 Introduction.................................................................................................................80 Materials and Methods...............................................................................................81 Materials..............................................................................................................81 Cell Culture and Treatment.................................................................................82 RNA Isolation and Analysis................................................................................83 Lipid Extraction...................................................................................................84 HDL Cholesterol Assay.......................................................................................84 Statistical Analysis..............................................................................................84 Results........................................................................................................................ .85 Effects of Conjugated Linoleic Acid on HepG2 Cells........................................85 Effects of Conjugated Linolei c Acid on H-4-II-E Cells......................................86 Role of PPAR in trans -10, cis -12 CLA-Induced Effects on Gene Expression.86 Discussion...................................................................................................................87 Summary.....................................................................................................................91 5 EFFECTS OF CIS AND TRANS ISOMERS OF OCTADECENOIC ACID ON LIPID METABOLIZING GENES AND HIGH-DENSITY LIPOPROTEIN CHOLESTEROL PRODUCTION IN CULTURED HUMAN AND RAT HEPATOCYTES......................................................................................................114 Introduction...............................................................................................................114 Materials and Methods.............................................................................................115 Materials............................................................................................................115

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vii Cell Culture and Treatment...............................................................................115 RNA Isolation and Analysis..............................................................................116 Lipid Extraction.................................................................................................117 HDL Cholesterol Assay.....................................................................................118 Statistical Analysis............................................................................................118 Results.......................................................................................................................1 19 Effects of Octadecenoic Acids on HepG2 Cells................................................119 Effects of Octadecenoic Acids on H-4-II-E Cells.............................................119 Role of PPAR in Vaccenic Acid-Induced Effects on Gene Expression..........120 Discussion.................................................................................................................121 Summary...................................................................................................................124 6 GENERAL DISCUSSION.......................................................................................147 APPENDIX LS MEANS AND P-VALUES FOR ANALYSIS OF FATTY ACID EFFECTS ON LIPID-M ETABOLIZING GENES AND HDL CHOLESTEROL PRODUCTION IN HEPG2 and H-4-II-E CELLS...................................................156 LIST OF REFERENCES.................................................................................................165 BIOGRAPHICAL SKETCH...........................................................................................194

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viii LIST OF TABLES Table page 1-1 Biological functions of key genes studied..................................................................4 A-1 Effects of n-3 and n-6 FA on lipid-m etabolizing genes and HDL cholesterol production in HepG2 cells......................................................................................156 A-2 Effects of n-3 and n-6 FA on lipid-m etabolizing genes and HDL cholesterol production in H-4-II-E cells...................................................................................156 A-3 Effects of WY 14,643 on mRNA res ponses to ST in HepG2 cells........................157 A-4 Effects of MK886 on mRNA respons es to ST in HepG2 cells..............................157 A-5 Effects of WY 14,643 on mRNA respons es to ST in H-4-II-E cells.....................158 A-6 Effects of MK886 on mRNA responses to ST in H-4-II-E cells...........................158 A-7 Effects of CLA on lipid-metabolizing genes and HDL cholesterol production in HepG2 cells............................................................................................................159 A-8 Effects of CLA on lipid-metabolizing genes and HDL cholesterol production in H-4-II-E cells..........................................................................................................159 A-9 Effects of WY 14,643 on mRNA responses to trans -10, cis -12 CLA in HepG2 cells.........................................................................................................................1 60 A-10 Effects of MK886 on mRNA responses to trans -10, cis -12 CLA in HepG2 cells.160 A-11 Effects of WY 14,643 on mRNA responses to trans -10, cis -12 CLA in H-4-II-E cells.........................................................................................................................1 61 A-12 Effects of MK886 on mRNA responses to trans -10, cis -12 CLA in H-4-II-E cells.........................................................................................................................1 61 A-13 Effects of cis and trans isomers of octadecenoic acid on lipid-metabolizing genes and HDL cholesterol production in HepG2 cells.........................................162 A-14 Effects of cis and trans isomers of octadecenoic acid on lipid-metabolizing genes and HDL cholesterol production in H-4-II-E cells......................................162

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ix A-15 Effects of WY 14,643 on mRNA responses to cis -vaccenic acid in HepG2 cells.163 A-16 Effects of MK886 on mRNA responses to cis -vaccenic acid in HepG2 cells.......163 A-17 Effects of WY 14,643 on mRNA responses to trans -vaccenic acid in H-4-II-E cells.........................................................................................................................1 64 A-18 Effects of MK886 on mRNA responses to trans -vaccenic acid in H-4-II-E cells.164

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x LIST OF FIGURES Figure page 3-1 Effect of long-chain FA on AC O mRNA expression in HepG2 cells......................59 3-2 Effect of long-chain FA on HM G-R mRNA expression in HepG2 cells.................60 3-3 Effects of long-chain FA on Apo A-I mRNA expression in HepG2 cells...............61 3-4 Effects of long-chain FA on HDL ch olesterol production in HepG2 cells..............62 3-5 Effects of long-chain FA on ACO mRNA expression in H-4-II-E cells..................63 3-6 Effects of long-chain FA on HMG-R mRNA expression in H-4-II-E cells.............64 3-7 Effects of long-chain FA on Apo A -I mRNA expression in H-4-II-E cells.............65 3-8 Effects of long-chain FA on HDL chol esterol production in H-4-II-E cells............66 3-9 Effect of WY 14,643 on ACO mRNA response to ST in HepG2 cells...................67 3-10 Effect of WY 14,643 on HMG-R mRNA response to ST in HepG2 cells...............68 3-11 Effect of WY 14,643 on Apo A-I mR NA response to ST in HepG2 cells..............69 3-12 Effect of MK886 on ACO mRNA response to ST in HepG2 cells..........................70 3-13 Effect of MK886 on HMG-R mRNA response to ST in HepG2 cells.....................71 3-14 Effect of MK886 on Apo A-I mRNA response to ST in HepG2 cells.....................72 3-15 Effect of WY14,643 on ACO mRNA re sponse to ST in H-4-II-E cells..................73 3-16 Effect of WY 14,643 on HMG-R mRNA re sponse to ST in H-4-II-E cells............74 3-17 Effect of WY 14,643 on A po A-I mRNA response to ST in H-4-II-E cells............75 3-18 Effect of MK886 on ACO mRNA response to ST in H-4-II-E cells.......................76 3-19 Effect of MK886 on HMG-R mRNA re sponse to ST in H-4-II-E cells..................77 3-20 Effect of MK886 on Apo A-I mRNA response to ST in H-4-II-E cells..................78

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xi 3-21 Regulation of lipid metabolizing ge nes and HDL cholesterol production by long-chain fatty acids...............................................................................................79 4-1 Effect of CLA on ACO mRNA expression in HepG2 cells.....................................93 4-2 Effect of CLA on HMG-R mR NA expression in HepG2 cells................................94 4-3 Effect of CLA on Apo A-I mRNA expression in HepG2 cells................................95 4-4 Effect of CLA on HDL chol esterol production by HepG2 cells..............................96 4-5 Effect of CLA acid on ACO mRNA expression in H-4-II-E cells...........................97 4-6 Effect of CLA on HMG-R mRNA expression in H-4-II-E cells.............................98 4-7 Effect of CLA on Apo A-I mR NA expression in H-4-II-E cells.............................99 4-8 Effect of CLA on HDL choles terol production by H-4-II-E cells.........................100 4-9 Effect of WY 14,643 on ACO mRNA response to trans -10, cis -12 CLA in HepG2 cells............................................................................................................101 4-10 Effect of WY 14,643 on HMG-R mRNA response to trans -10, cis -12 CLA in HepG2 cells............................................................................................................102 4-11 Effect of WY 14,643 on Apo A-I mRNA response to trans -10, cis -12 CLA in HepG2 cells............................................................................................................103 4-12 Effect of MK886 on ACO mRNA response to trans -10, cis -12 CLA in HepG2 cells.........................................................................................................................1 04 4-13 Effect of MK886 on HMG-R mRNA response to trans -10, cis -12 CLA in HepG2 cells............................................................................................................105 4-14 Effect of MK886 on Apo A-I mRNA response to trans -10, cis -12 CLA in HepG2 cells............................................................................................................106 4-15 Effect of WY 14,643 on ACO mRNA response to trans -10, cis -12 CLA in H-4-II-E cells..........................................................................................................107 4-16 Effect of WY 14,643 on HMG-R mRNA response to trans -10, cis -12 CLA in H-4-II-E cells..........................................................................................................108 4-17 Effect of WY 14,643 on Apo A-I mRNA response to trans -10, cis -12 CLA in H-4-II-E cells..........................................................................................................109 4-18 Effect of MK886 on ACO mRNA response to trans -10, cis -12 CLA in H-4-II-E cells.........................................................................................................................110

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xii 4-19 Effect of MK886 on HMG-R mRNA response to trans -10, cis -12 CLA in H-4-II-E cells..........................................................................................................111 4-20 Effect of MK886 on Apo A-I mRNA response to trans -10, cis -12 CLA in H-4-II-E cells..........................................................................................................112 4-21 Regulation of lipid metabolizing ge nes and HDL cholesterol production by CLA........................................................................................................................113 5-1 Effect of cis and trans isomers of octadecenoic ac id on ACO mRNA expression in HepG2 cells........................................................................................................126 5-2 Effect of cis and trans isomers of octadecenoic acid on HMG-R mRNA expression in HepG2 cells......................................................................................127 5-3 Effect of cis and trans isomers of octadecenoic acid on Apo A-I mRNA expression in HepG2 cells......................................................................................128 5-4 Effects of cis and trans isomers of octadecenoic acid on HDL cholesterol production by HepG2 cells.....................................................................................129 5-5 Effect of cis and trans isomers of octadecenoic ac id on ACO mRNA expression in H-4-II-E cells.....................................................................................................130 5-6 Effect of cis and trans isomers of octadecenoic acid on HMG-R mRNA expression in H-4-II-E cells...................................................................................131 5-7 Effect of cis and trans isomers of octadecenoic acid on Apo A-I mRNA expression in H-4-II-E cells...................................................................................132 5-8 Effects of cis and trans isomers of octadecenoic acid on HDL cholesterol production in H-4-II-E cells...................................................................................133 5-9 Effect of WY 14,643 on ACO mRNA response to cis -vaccenic acid in HepG2 cells.........................................................................................................................1 34 5-10 Effect of WY 14,643 on HMG-R mRNA response to cis -vaccenic acid in HepG2 cells............................................................................................................135 5-11 Effect of WY 14,643 on Apo A-I mRNA response to cis -vaccenic acid in HepG2 cells............................................................................................................136 5-12 Effect of MK886 on ACO mRNA response to cis -vaccenic acid in HepG2 cells.........................................................................................................................1 37 5-13 Effect of MK886 on HMG-R mRNA response to cis -vaccenic acid in HepG2 cells.........................................................................................................................1 38

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xiii 5-14 Effect of MK886 on Apo A-I mRNA response to cis -vaccenic acid in HepG2 cells.........................................................................................................................1 39 5-15 Effect of WY 14,643 on ACO mRNA response to transvaccenic acid in H-4-II-E cells..........................................................................................................140 5-16 Effect of WY 14,643 on HMG-R mRNA response to trans -vaccenic acid in H-4-II-E cells..........................................................................................................141 5-17 Effect of WY 14,643 on Apo A-I mRNA response to trans -vaccenic acid in H-4-II-E cells..........................................................................................................142 5-18 Effect of MK886 on ACO mRNA response to transvaccenic acid in H-4-II-E cells.........................................................................................................................1 43 5-19 Effect of MK886 on HMG-R mRNA response to trans -vaccenic acid in H-4-II-E cells..........................................................................................................144 5-20 Effect of MK886 on Apo A-I mRNA response to transvaccenic acid in H-4-II-E cells..........................................................................................................145 5-21 Regulation of lipid metabolizing ge nes and HDL cholesterol production by cis and trans octadecenoic fatty acids.........................................................................146

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xiv ABBREVIATIONS KEY AA arachidonic acid ABCA1 adenosine triphosphate-binding cassette transporter-A1 ACC acetyl-CoA carboxylase ACO acyl-CoA oxidase ACP acyl carrier protein AI adequate intake Apo apolipoprotein BMI body mass index CoA coenzyme A CHD coronary heart disease CLA conjugated linoleic acid CM chylomicron CPT-I or -II carnitine-pa lmitoyl transferase I or -II CVD cardiovascular disease DGAT diacylglygerol acyltransferase DHA docosahexaenoic acid EPA eicosapentaenoic acid ER endoplasmic reticulum ETF electron transfer flavoprotein

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xv FA fatty acid FABPc cytosolic fatty acid binding protein FAS fatty acid synthase GLA gamma-linolenic acid HDL high-density lipoprotein HMG-R 3-hydroxy, 3-methylglutaryl CoA reductase HODE hydroxyoctadecadienoic acid IUPAC International Union of Pure and Applied Chemistry LA linoleic acid LCAT lecithin:cholesterol acyltransferase LNA linolenic acid LPL lipoprotein lipase MGAT monoacylglycerol acyltransferase MTP microsomal transfer protein MUFA monounsaturated fatty acid NCEP National Cholesterol Education Program NEFA non-esterified fatty acid PPAR peroxisome proliferator-activated receptor PPRE peroxisome proliferator response element PUFA polyunsaturated fatty acid RXR retinoid X receptor SCD stearyl-CoA desaturase ST stearic acid

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xvi TAG triacylglycerol TPN total parenteral nutrition TZD thiazolidinediones VLDL very low-density lipoprotein

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xvii Abstract of Dissertation Pres ented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy EFFECTS OF LONG-CHAIN FATTY AC IDS ON LIPID METABOLIZING GENES AND HIGH-DENSITY LIPOPROTEIN CHOLESTEROL PRODUCTION IN CULTURED HUMAN AND RAT HEPATOCYTES By Elizabeth Sarah Greene May 2006 Chair: Lokenga Badinga Major Department: Animal Sciences A series of experiments were conducted to examine the short-term effects of long-chain fatty acids (FA) on acyl CoA oxi dase (ACO), 3-hydroxy, 3-methylglutaryl CoA reductase (HMG-R) and apolipoprotein A-I (Apo A-I) gene expression, and high-density lipoprotein c holesterol (HDL-C) producti on in HepG2 (human) and H-4-II-E (rat) hepatocytes. In the three e xperiments, the FA studied were 1) FA of differing saturation and chain length, 2) conjugated linol eic acid (CLA), and 3) cis (c9, c11) and trans (t9, t11) isomers of octadecenoic acid. In HepG2 cells, ACO mRNA was up-regulated by trans -10, cis -12 CLA and cis -vaccenic acid (c11). HMG-R gene expressi on was increased by stearic acid (ST) and trans -10, cis -12 CLA. Steady-state levels of Apo A-I mRNA were increased by all FA in the first experiment, trans -10, cis -12 CLA, and c11. HDL-C was decreased only by cis -9, trans -11 CLA. In H-4-II-E cells, ACO mR NA was up-regulated by LA, CLA, ST,

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xviii oleic acid, and trans -vaccenic acid (t11). HMG-R gene expression was increased by ST, CLA isomers, and t11. Apo A-I was increa sed by ST and EPA, but decreased by CLA and cis and trans monounsaturated FA. HDL-C wa s increased by LNA in the first experiment. Based on these findings, we investigated th e possibility that the FA effects are mediated by peroxisome pro liferator-activ ated receptor (PPAR ). In HepG2 cells, activation or inhibition of PPAR had minimal effects on basal or FA-effects on gene expression, consistent with the low-levels of endogenous PPAR in this cell line. In H-4-II-E cells, activation of PPAR increased the abundance of basal ACO mRNA, enhanced the effect of ST on ACO and Apo A-I mRNA, and enhanced the effects of t11 on ACO, HMG-R, and Apo A-I gene expression. Inhibition of PPAR decreased basal expression of ACO and attenuated the effect s of ST and t11 on ACO and effects of trans -10, cis -12 CLA on Apo A-I gene expression. Th ese results indicate that specific FAs may regulate lipid-metabolizing genes in the liver through a PPAR -dependent mechanism. Because of different responses to FA in human and rat cell lines, however, net effects are likely species specific.

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1 CHAPTER 1 INTRODUCTION Dietary fat is an important nutrient for the function and survival of all organisms. Historically, body lipids have been considered primarily to serve as an energy source, as constituents of cell membranes, and as pr ecursors for molecules involved in signal transduction, such as steroids and prostagl andins. More recently however, fatty acids (FA) have been shown to affect gene expression, leading to changes in cell differentiation, growth, and metabolism (Clark e and Jump, 1994; Jump et al., 1996). Additionally, dietary fat has been implicat ed in the progression of several chronic diseases, including type II diabetes, cardiova scular disease, and some types of cancer (Sanders, 2003), though the effects may de pend on the composition of dietary fat consumed. Therefore, unde rstanding the molecular basis for FA effects on gene regulation is necessary for furt her elucidation of the role of fats in human health. To address this issue, our studies focused on the effects of three genera l classes of FA that may play a significant role in health and metabolism: n-3 and n-6 long-chain polyunsaturated fatty acids, conjug ated linoleic acids (CLA), and cis and trans isomers of fatty acids. Dietary polyunsaturated fatty acids (PUFA) have been reported to lower blood triglycerides, alter the blood lipid profile, decrease intram uscular lipid droplet size, improve insulin sensitivity, and enhance gluc ose utilization (Jump and Clarke, 1999). Since the seminal observation that PUFAs c ould inhibit hepatic lipogenesis in mice (Allmann and Gibson, 1965), numerous studies ha ve demonstrated that diets rich in

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2 PUFAs influence metabolic changes by coordi nately suppressing lipid synthesis in the liver and enhancing fatty acid oxidation in bo th liver and skeletal muscle (Jump and Clarke, 1999). The PUFA induction of gene s encoding proteins involved in lipid oxidation include 3-hydroxy, 3-methylglutaryl -CoA synthase (Rodr iguez et al., 1994), carnitine palmitoyltransferase, fatty acid bi nding proteins, and pe roxisomal acyl-CoA oxidase (ACO; Reddy and Hashimoto, 2001). Conjugated linoleic acid (C LA) is a collective term for positional and geometric isomers of linoleic acid (LA). Though over 16 individual isomers ha ve been identified (Rickert et al 1999), only cis -9, trans -11 CLA and trans -10, cis -12 CLA are known to possess biological activity (Pariza et al., 2000). Cis -9, trans -11 CLA is the predominant CLA produced as an intermediate in the rumen during biohydrogenation of dietary LA and is commonly found in dairy products a nd ruminant meat. Dietary sources of trans -10, cis -12 CLA derive predominantly from s ynthetic partial hydrogenation and are found in margarines, shortenings, and supplements (Gaullier et al., 2002 ). First identified in grilled beef as a poten tial anti-carcinogen (Pariza a nd Hargraves, 1985), numerous health benefits have been attributed to CLA mixtures, including actions as an antiadipogenic (Park et al, 1997), antidiabe togenic (Houseknecht et al., 1998), and antiatherosclerotic (Kritchevsky et al., 2004) agent. More rece ntly, studies involving individual isomers have shown that the two ma in isoforms can have different effects on metabolism and cell function and may act throu gh different signaling pathways (Wahle et al., 2004). Metabolic responses to cis -9, trans -11 and trans -10, cis -12 CLA may differ, but both isomers have implicati ons for human healt h. Most studies have been performed in animal models, with species differences observed. In particular, only some of the

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3 findings attributed to animal models pe rtain to human subjects, and even when comparing studies in humans, results are often inconclusive (Terpstra, 2004). Trans -fatty acids are geometrica l isomers of unsaturated FA that assume a saturated fat-like configuration that di ffers from that of their cis counterparts. The predominant source of trans fats in the human diet is hydroge nated oils (such as margarine and partially hydrogenated soybean oil) commonly found in bake d goods and deep fat-fried fast foods (Hu et al., 2001). Metabolic st udies in several species have shown that trans -FA can negatively alter the lipid profile to a greater extent than saturated fats, because trans -FA not only increase small, dense LD L cholesterol (Mauger et al., 2003), but also decrease HDL cholesterol in some studies (Judd et al., 1994; de Roos et al., 2003). Additionally, epidemiological evidence associates trans -FA intake with increased risk for cardiovascular disease (Ascherio et al., 1999). Few studies, however, have examined the role of individual trans -FA in modulating lipid metabolism. As with other FA, it is possible that cis and trans isomers of octadecenoic acid may also have differential effects on lipid metabolism. Based on both dietary and in vitro studies of lipid metabolism, we hypothesized that various FA of differing degree of saturation and double-bond position will have differing effects on ACO, 3-hydroxy, 3-methyl glutaryl CoA reductase (HMG-R), and apolipoprotein A-I (Apo A-I) gene expressi on, as well as HDL cholesterol production in HepG2 and H-4-II-E hepatoma cells (Table 1-1). Also, because several FA and their derivatives are known ligands for peroxisome proliferator-activated receptors (PPAR; Schoonjans et al., 1996), we hypothesized that these FA may act on lipid-metabolizing genes through activation of PPAR the predominant recepto r subtype in the liver

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4 (Braissant et al., 1996). If this hypot hesis is correct, activation of PPAR should mimic the effects of FA, wher eas inhibition of PPAR would be expected to block FA effects in HepG2 and H-4-II-E hepatoma cell lines. The overall aim of our studies was to examine the differential roles of fatty acids on lipid metabolizing genes involved in peroxisomal -oxidation and cholesterol synthesis in human and rat hepatoma cell lines. Table 1-1. Biological functi ons of key genes studied Gene Function ACO Rate limiting in peroxisomal -oxidation HMG-R Rate limiting in cholesterol synthesis; converts HMG-CoA to mevlonate Apo A-I Necessary for proper packaging of HDL cholesterol

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5 CHAPTER 2 LITERATURE REVIEW Structure and Metabolism of Lipids Structure and Nomenclature of Lipids Based on physical properties, the term lip id denotes a heterogeneous group of substances that are insoluble in water, but are soluble in non-polar solvents such as chloroform and alcohols (Smith, 2000). Th is definition covers a wide range of molecules, including FA, phospholipids, sterol s, sphingolipids, terpenes, and others (Christie, 2003). Fatty acid s consist of a chain of two or more carbon atoms, with a methyl group at one end, and a carboxyl group at the other end of the chain. The main structural features are thei r chain length, degree of unsaturation (number of double bonds), and presence of substituent groups. Additionally, the pr esence of double bonds allows for positional and geometric isomerism. Positional isomers occur when double bonds are located at different positions along the carbon chain. The position of unsaturation is numbered in reference to th e first of the pair of carbon atoms between which the double bond occurs. Geometric isomer ism refers to the configuration of the hydrogen atoms in respect to the double bond. If the hydrogen atoms are on the same side of the molecule opposite the double bond, it is said to be in the cis configuration. Alternately, if the hydrogen atoms are on opposite sides, the configuration is trans Most naturally occurring unsatu rated FA are in the cis configuration, but na tural and synthetic trans isomers do exist.

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6 The naming scheme for FA must be able to clearly define a lipid structure in a manner that is amenable to scientists and re searchers of all fields. Several systems are currently used, though to different degrees. Fi rst, there are the triv ial names, such as stearic acid and linoleic acid, which were assign ed as the individual FA were discovered. Although these may be used for the mo st-commonly occurring FA, naming and remembering unusual unsaturated, branch ed, or hydroxyl-FA becomes unwieldy. Because of this difficulty, two different syst ems have been developed. The older system used Greek letters to identify carbon atoms, beginning at the carboxyl end. Considering the carboxyl carbon as C1, C2 is called the -carbon, C3 the -carbon, and so on, ending with the -carbon at the methyl end. Though this syst em is no longer preferred, it is used to name the -3 and -6 FAs, in which the last double bond in the chain occurs three and six carbons from the -carbon, respectively. In much of the newer literature, the is often replaced by an n but the meaning remains the same. Currently, the preferred system for sp ecifying individual FA is the numbering system standardized by the International Un ion of Pure and Applied Chemistry (IUPAC) (IUPAC-IUB, 1977). For linoleic ac id, an 18-carbon FA with two cis double bonds in positions 9 and 12 from the carboxyl end, the systematic name is cis -9, cis -12 octadecadienoic acid. In the shorthand system, FA are identified by two numbers separated by a colon; the first number indi cates the number of carbon atoms, the second indicates the number of double bonds in the stru cture. For example, a saturated fat such as stearic acid would be 18:0, whereas a polyunsat urated fat, such as linoleic acid would be represented by cis -9, cis -12 18:2.

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7 Biosynthesis of Fatty Acids Most naturally occurring FA contain an even number of carbon atoms, leading early researchers to speculat e that they were formed by the condensation of two-carbon units. This was confirmed using ra ts fed acetic acid labeled with 13C in the carboxyl group and 2H in the methyl group. When FA were isolated from the rat tissues, the labeled carbons were found in alternate positions along the chain, showing that the complete FA could be derived from acetic ac id (Rittenberg and Bloch, 1944). When the details of -oxidation were elucidated in the 1950s, it led to speculation that FA synthesis could be the simple reversal FA breakdown. However, several discoveries soon showed that the pathways were distinctly different First, NADPH (not NAD+ as in oxidation) serves as a cofactor. Second, there is a re quirement for bicarbonate (Wakil, 1962; Brady and Gurin, 1952). Fatty acid synthesis can be broken into two basic pro cesses: condensation of two carbon units to form 16 to18-carbon FA and vari ous modifications of these products. In mammals, the majority of carbon for de novo FA synthesis comes from pyruvate, the end-product of glycolysis. To be used in FA synthesis, acetyl coenzyme A (CoA; the activated form of acetic acid) must be gene rated from pyruvate. To accomplish this, the pyruvate is transported from the cytosol into the mitochondria, where the enzyme pyruvate dehydrogenase acts to produce acetyl-CoA. Acet yl-CoA and oxaloacetate combine to form citrate, which can then be transported back out of the mitochondria via a tricarboxylate anion carrier, where the cycl e is completed, and acetyl-CoA is produced by the action of ATP:citrate lyase.

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8 The first and rate-limiting reaction in de novo FA synthesis is catalyzed by acetyl-CoA carboxylase (ACC). In this enzy matic reaction, acetyl -CoA is carboxylated, leading to the formation of malonyl-CoA (Knowles, 1989). Th is reaction requires biotin as a cofactor, as shown by inhibition of carboxylation by avidin, a potent inhibitor of biotin (Wakil et al., 1958). Acetyl-CoA carboxylase is ac tivated by phosphorylation and deactivated by dephosphorylation (Shacter et al., 1986). The malonyl-CoA generated by ACC forms the source of nearly all carbons of the FA. Only the fi rst two carbons arise from the “primer molecule,” acetyl-CoA. In order for individual malonyl-CoA units to join into the FA chain, they must be attached to the acyl carrier protein (ACP). The ACP is a small molecular mass protein (8.8 kDa) th at is very stable ove r a range of pH and temperature values (D’Agnolo et al., 1975). The enzymatic steps involved in adding su ccessive malonyl-CoA units to the chain are collectively known as fatty acid synthase (FAS). In animals, FAS is a multifunctional enzyme, with discrete domains catalyzi ng the condensation, dehydration, and reduction reactions. Animal FAS complexes consist of homodimers with molecular weight of approximately 450-550 kDa (Smith, 1994). Essentially, to elongate the chain, malonyl-ACP attached to one half of the di mer interacts with th e growing acyl chain attached to the active site of the condensing enzyme on the other half of the dimer (Joshi et al., 1998). The typical end product of FAS is palmitic acid (16:0). The thioesterase actions of FAS cleave the product from the en zyme. This specificity for a 16-carbon product is likely due to stearic hindrance of the condensing domain by the large FA (Chirala and Wakil, 2004). Although the production of palmitate is most common, different organisms and tissues can produce FA of shorter chain lengths as necessary.

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9 For example, in the rat mammary gland, where large amounts of 8:0 and 10:0 are necessary for the formation of milk triacylglyce rols (TAG), a second thioesterase is present, forming medium-chain FA (Smith, 1994). Although the main product of FAS is palmita te, many tissues contain longer chain FA, particularly as a component of membrane lipids. The formation of long and very long-chain FA is catalyzed by Type III synt hases, often termed elongases due to their lengthening of pre-formed and dietary FA In mammalian tissues, two separate elongation systems are located in the mitochondr ia and endoplasmic reticulum (ER). In the mitochondria, two carbon units in the form of acetyl-CoA (not malonyl-CoA as in de novo synthesis) are added preferentially to monoenoic over saturated substrates (Moon et al., 2001). Mitochondrial elongation is essentially a reversal of -oxidation, with a requirement for NADPH and NADH (Seubert and Podack, 1973). Formation of the longer chain FA occurs at the ER. In this case, malonyl-CoA serves as the two carbon donor and NADPH is the reducing agent. Th is system can produce FA with chain lengths in excess of 20 car bons (Suneja et al., 1990). Once saturated FA have been produced by the organism, they can be used to produce unsaturated FA, mainly by the pro cess of oxidative desaturation. In this mechanism, a double bond is intr oduced directly into a preformed saturated long-chain FA, using O2 and a reducing compound (NADH) as co factors (Scheuerbrandt and Bloch, 1962). Mammalian enzymes normally introdu ce new double bonds between an existing double bond and the carboxyl group, whereas plant enzymes introduce the new bond between an existing double bond and the te rminal methyl group. There are three components to the desaturation complex: NADH-cytochrome b5 reductase, cytochrome

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10 b5, and the desaturase enzyme (Stritmatter et al., 1974). Most of what is known about desaturases is derived from early studies showing that 9 desaturase is the rate-limiting step in the conversion of stearic acid (18:0) to oleic acid (18:1, n-9) (Schroepfer and Bloch, 1965). Because of its act ions, it is also referred to as stearyl-CoA desaturase (SCD). Mammals contain desaturase s able to introduce double bonds in the 5, 6, and 9 positions. Plants additionally possess the 12 and 15 desaturases necessary for the formation of n-6 and n-3 FA. Degradation of Fatty Acids In the body, FA from dietary or stored T AG are broken down to provide a source of energy. The main forms of FA oxidation are termed alpha ( ), beta ( ), and omega ( ), depending on the carbon in the chain that is attacked. Of the three types of oxidation, -oxidation is the most prevalent. The basic mechanism for -oxidation was originally proposed by Knoop in 1904 after feeding labeled FA to dogs, and was c onfirmed by Dakin’s isolation of the proposed intermediates in 1912. Fats degraded in this manner liberate two-carbon units in the form of acetyl-CoA through the introduction of a double bond between the and -carbons, hence the name -oxidation. Until relatively recently, mitochondria were considered the only cellular site for -oxidation. Although all the necessary enzymes are present in mitochondria, the microbodies (peroxisomes in mammals and glyoxysomes in plants) can also complete the process (Lazarow a nd de Duve, 1976). The contribution of microbodies to total -oxidation depends on the organism and specific tissue considered. For example, in mammals, peroxisomal -oxidation of very long-chain FAs is

PAGE 29

11 particularly important in the liver and ki dneys, with defects leading to devastating diseases (Fournier et al., 1994). Fatty acyl-CoAs formed within the cytoso l cannot enter the mito chondrion directly, providing a major point of control and regul ation of FA metabolism (Eaton, 2002). The observation that carnitine stimulates the -oxidation of long-chain FA gave the first clue to its function in the mitoc hondrial uptake of FA (Bremer, 1962; Fritz and Yue, 1963). Acyl residues are transferred to carnitine by carnitine-palmitoyl tran sferase (CPT-I) at the surface of the outer mitochondrial membrane. This allows the FA to transverse the membrane via porin, where they are then transported through the inner mitochondrial membrane by a carnitine:acylcarnitine translocase (Pande, 1975). The translocase causes a one-to-one exchange of carnitine for acylcarn itine, ensuring a constant level of carnitine within the mitochond ria (Ramsay and Tubbs, 1975). On ce within the mitochondrial matrix, a second carnitine-palmit oyl transferase, CPT-II, acts to transfer the acyl group from carnitine back to CoA, reforming acyl-CoAs, the substrate for -oxidation (Bieber, 1988). The reactions of -oxidation involve four enzymes in repeated sequence, resulting in the cleavage of two carbons at a time fr om the acyl chain. Acyl-CoA dehydrogenase acts to produce trans -2,3-enoyl-CoA. This step is lin ked to the respiratory chain via electron transfer flavoprotei n (ETF) and ETF-ubiquinone oxire ductase (Parker and Engel, 2000). The 2-enoyl-CoA hydratase then ac ts on the product of the first reaction, producing 3-hydroxyacyl-CoA. The next enzy me in the sequence, 3-hydroxyacyl-CoA dehydrogenase, is linked with NAD+ and produces 3-oxoacyl-CoA. Finally, 3-oxoacyl-CoA thiolase actions produce a satu rated acyl-CoA that has been shortened by

PAGE 30

12 two carbons, in the form of acyl-CoA (Eaton et al., 1996). Each of the enzymes is present in several isoforms w ith varying chain-length specificities, primarily for short, medium, long, and very long-chain acyl-CoA. This allows for improved efficiency of -oxidation and prevents buildup of intermediates that could lead to inhibition (Bartlett and Eaton, 2004). Peroxisomal -oxidation is important in almost all eukaryotic organisms (Kunau et al., 1995). The peroxisomal a nd mitochondrial enzymes of -oxidation differ in several ways. Peroxisomes do not have an elec tron transport system coupled to energy production as can be found in mitochondria. The first and rate limiting step in peroxisomal -oxidation is catalyzed by acyl-CoA oxidase (ACO), which introduces a trans -2 double bond and produces hydrogen per oxide. Next, a trifunctional enzyme produces -ketoacyl-CoA, which is acted upon by a thiolase, producing acetyl-CoA and a shortened acyl-CoA (Mannaerts et al., 2000). Due to limited substrate specificities for ACO, peroxisomes are incapable of oxidizing long-chain FA completely (Singh et al., 1984). Medium chain products of peroxisomal -oxidation are transfe rred to carnitine, allowing them to be transported into the m itochondria for complete oxidation. Defects in peroxisomal -oxidation can lead to the accumulation of very long-chain FA in various tissues, producing devastating diseases such as Zellweger syndrome and adrenoleukodystrophy (Kunau et al., 1995). The above enzymatic cycle assumes that the substrate is a straight-chain, saturated FA with an even number of carbons For FA of odd-chain lengths, -oxidation yields propionyl-CoA in addition to acetyl-CoA; theref ore, the ability of an organism or tissue to oxidize these FA depends on the ability of that organism or tissue to metabolize

PAGE 31

13 propionate. The liver is well-equipped to oxidize propionate and oxidizes odd-chain FA well, whereas the heart cannot oxidize the product, and -oxidation of odd-chain FA stops with an increase in propionate (Gr ynberg and Demaison, 1996). Beta-oxidation of unsaturated FA poses several problems. Most naturally occurring unsaturated FA contain cis double bonds and the bonds may be in the wrong position along the chain for effective -oxidation. Unsaturated FA with odd-numbered double bonds, such as the 9cis bond of LA, are shortened to 3cis -enoyl-CoA and then isomerized to 2trans -enoyl CoA that can be further degraded via -oxidation (Stoffel and Caesar, 1965). Fatty acids with even-numbered double bonds are shortened to 4cis -enoyl-CoA, which are then dehydrogenated to 2trans 4cis -dienoyl-CoA. One double bond is then removed by NADPH-dependent 2,4-dienoyl-reductase, allowing -oxidation to continue (Kunau and Dommes, 1978). The acyl-CoA produced by chain-shortening can have several different fates, depending on the tissue. In ketogenic tissues such as the liver, acetyl-CoA is used to form the ketone bodies, acetoacetate and -hydroxybutyrate, for e xport and peripheral oxidation. In most tissues, however, acetyl-C oA enters the Krebs cycle and generates energy in the form of ATP (Hiltunen and Qin, 2000). Nutritional and Biological Properties of the Polyunsaturated Fatty Acids Dietary Requirements of the Essential Fatty Acids Mammalian cells can synthesize saturate d and omega-9 (n-9) unsaturated FA de novo from acetyl-CoA, but lack the 12 and 15 desaturase enzymes necessary for the formation of double bonds in the omega-6 (n-6 ) and omega-3 (n-3) positions. Because of this enzyme deficiency, the lin oleic acid (LA; 18:2, n-6) and -linolenic acid (LNA; 18:3, n-3) are considered essentia l nutrients in the human diet (Innis, 1991). Once ingested,

PAGE 32

14 LA and LNA can be further elongated and desaturated into biologically important long-chain polyunsaturated fatty acids (PUFA) of 20 or more carbons and three to six double bonds. Determining the essential requirements of a nutrient generally begins with recognition of a deficiency, c ontinues with the study of intakes that can prevent or reverse the deficiency, and finally concludes wi th the definition of a range of intakes for optimal biological function. In 1929, Burr and Burr discovered that rats fed a fat-free diet developed dermatitis and grew at a slower rate than their fat-fe d counterparts. These deficiencies could only be elim inated by adding certain FAs to the diet, which were later determined to be LA and arachidonic acid (AA; 20:4). This knowledge was applied to produce essential FA deficiency in a variety of species, including man. In all species, the deficiency is characterized by skin symptoms such as dermatosis or eczema, retarded growth, impaired reproduction, and degenera tion or impairment of function in many bodily organs, including the heart and kidne ys (Sinclair, 1990). These signs are characterized by changes in the FA com position of many tissues, particularly in biological membranes and mitochondria. Well-documented essential FA deficiency in man is rare, but was first seen in the 1940s and 50s in infants receiving formula cont aining skim milk and su gar as a substitute for mother’s milk. When fed formula cont aining increasing concentrations of LA, clinical signs of defici ency disappeared when concentratio ns in the diet were above 0.1% of dietary energy (Hansen et al., 1958). Adult essential FA deficiency was most commonly seen in patients receiving total pa renteral nutrition (T PN), in which early formulas were fat-free (Holman, 1981). In some cases, patients responded to the

PAGE 33

15 application to the skin of fats with a hi gh proportion of LA, showing that the FA do not necessarily have to be absorbed through the gastrointestinal tr act to be effective. More frequently, LA deficiency may develop as a secondary condition to other disorders such as severe malnutrition and fat malabsorption. The n-3 FA can, in part, substitute for a de ficiency in n-6 FA, but also have their own distinct roles (Benatti et al., 2004). The understanding of n-3 FA essentiality lagged significantly behind that of n6 FA, partially because of th eir naturally lower amounts in the body. The first case of n-3 FA defici ency was induced by an n-3 FA-free TPN formula. Symptoms of n-3 FA deficiency in the patient included numbness, tingling, weakness, inability to walk, leg pain, psychological disturba nces and blurred vision. The patient’s plasma lipid profile showed the con centration of total n-3 FA to be at 34% of the control value. When soybean oil, a source of LNA, was added back to the TPN formula, the signs of deficiency disappeared (Holman et al., 1982). As essential FA deficiency is usually associ ated with a disease state, there is little evidence to determine dietary reference intake s for healthy populations. Therefore, based on the data that are available on health eff ects of LA and LNA, adequate intake (AI) levels have been recommended. The AI is a value based on experimentally derived intake levels or approximate mean nutrient intakes by a group of h ealthy individuals. Based on current estimates, PUFAs contribu te approximately 5-6% of energy in the Western diet (Grundy et al., 1982). For adults it is recommended that consumption of LA should be 17 g/d for men and 12 g/d for women. For LNA, recommended intakes are 1.6 g/d for men and 1.1 g/d for women (Food and Nutrition Board, 2005). Also, for cardiovascular health benef its, the long-chain desatura tion and elongation products,

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16 eicosapentaenoic acid (EPA, 20:5) and doc osahexaenoic acid (DHA, 22:6) together should represent 0.3% of dietary energy, with each FA being at le ast 0.1% of energy (Simopoulos et al., 2000). These values repres ent, in general, a decrease in n-6 FA consumption and an increase in n-3 FA cons umption for the typica l individual, altering the current n-6 to n-3 ratio from 1020:1 to 1-4:1 (Simopoulos, 1999). Long-Chain Polyunsaturated Fatty Acids of the n-6 Family Organs and tissues performing storage (adi pose tissue), chemical processing (liver), mechanical work (muscle), and excretion (ki dney) have membranes in which the n-6 FA predominate, particularly with AA as th e major component (Innis, 1991). Arachidonic acid serves an important role as a precurso r for biologically important eicosanoids and it and other n-6 FA may play a role as sec ondary messengers in the process of signal transduction. As previously stated, LA can be desaturated and elongated in mammals to produce biologically important long-chain PUFAs of the n-6 family (Klenk and Mohrhauer, 1960; Mead, 1968). These include -linolenic acid (GLA, 18:3) and AA. Normally, only a small proportion of dietary linoleate can be converted to longer-chain PUFA. Most of it is -oxidized to provide energy (C unnane and Anderson, 1997). To produce GLA, 6 desaturase acts on LA, introduc ing a double bond at carbon six in the FA chain. This product is then elongated to dihomo-linoleic acid (20:3, n-6), which is converted to AA by 5 desaturase. Arachidonic acid can be elongated to form adrenic and -6-tetracosatetraenioc acids (22:4 and 24: 4), but since there is no evidence of a functional mammalian 4 desaturase, -6-docosapentaenoic acid (22:5) must be formed via an alternate pathway. A double bond is added to tetracosatetraenioc acid by 6

PAGE 35

17 desaturase, forming tetracosapentaenoic acid, which is then oxidized in peroxisomes to form the 22 carbon product (Sprecher et al., 1995; Ferdinandusse et al., 2001). Long-Chain Polyunsaturated Fatty Acids of the n-3 Family Nervous tissue, reproductive organs, and th e retina have membranes that contain a large percentage of long-chain FA, particularly PUFAs of the n-3 series (Innis, 1991). As with LA and the n-6 PUFAs, dietary LNA can be elongated and desaturated to form long-chain PUFAs of the n-3 family. The ke y n-3 PUFAs are EPA and DHA. Within the body, the amounts of n-3 PUFAs are lo wer than that of the n-6 PU FAs. This is due to the small proportion of LNA in the diet, as we ll as the competition between FA for the 5 and 6 desaturases (Dang et al., 1989). To produce EPA, LNA is desaturated by 6 desaturase to stearidonic acid (18:4), then el ongated by 2 carbons, and further desaturated by 5 desaturase. As with PUFA of the n-6 series, no 4 desaturase is present in mammals, as in lower eukaryotes, to form DH A directly (Qiu et al., 2001). To form DHA, EPA undergoes two cycles of chain elongation to produce -3-tetracosapentaenoic acid, which is then desaturated to -3-tetracosahexaenoic acid ( 24:6) (Sprecher et al., 1995). In mammals, EPA and DHA can be derive d not only from dietary LNA, but also in the diet directly from sources such as cold-water fish and fish oil. Digestion and Assimilation of Dietary Fats In the Western diet fats constitute approximately 40% of energy in the diet. In the human diet, the majority of fa t consumed, whether of animal or plant origin, is in the form of TAG. Triacylglycerols are the majo r biological form of storage lipid, composed of three FA esterified to a glycerol backbone Long-chain FA, such as oleic acid (18:1) and palmitic acid (16:0) are the major FA present in dietary TAG, although FA can vary

PAGE 36

18 in chain length from C2 to C24 and from sa turated FA to unsaturated FA with six or more double bonds. In addition to TAG, smaller amounts of phospholipids, cholesterol, and other sterols are consumed in the diet. An average adult on a Western diet consumes approximately 150 grams of TAG and 4-8 gr ams of phospholipids daily. Cholesterol intake can vary depending on diet, but average daily intake of total cholesterol is 400-500 mg (Rizek et al., 1974). Although the majority of TAG digestion occu rs in the small intestine, digestion begins in the stomach. Partial hydrolysis of TAG begins with the actions of lingual or gastric lipase, depending on the species studi ed (Mu and Hoy, 2004). Lingual lipase is secreted by the von Ebner’s glands of the tongue and is transported wi th the food bolus to the stomach (Hamosh and Scow, 1973). Gastri c lipase is secreted from the gastric mucosa. Secretion of either of these lipases can be stimulated mechanically (suckling and swallowing), neurally (sympathetic agonis ts), and by diet (high fat) (Hamosh, 1978). The relative contribution of these lipases to fat hydrolysis is species dependent. For example, rodents have a relatively high activ ity of lingual lipase and low activity of gastric lipase, whereas, in pr imates, gastric lipase has hi gh activity (Mu and Hoy, 2004). Both lingual and gastric lipases show a stereo -specific preference for cleaving TAG at the sn-3 position, regardless of the FA present, although shor tand medium-chain FA are hydrolyzed at a faster rate th an long-chain FA (Jensen et al ., 1983). This preferential cleavage gives rise to diglycerides and nonesterified fatty acids (NEFA) as major digestion products. Approximately 10-30% of dietary fat is partia lly hydrolyzed in the stomach which facilitates further digestion in the small intestine (Hamosh and Scow, 1973). In addition, the churning action of the stomach creates a coarse emulsion

PAGE 37

19 stabilized by phospholipids, and proteolytic digestion in the st omach serves to release fats from food particles where they are generally associated with proteins (Gurr et al., 2002). The major digestion of dietary TAG results from the actions of pancreatic lipase. Entry of TAG, TAG degradation products, and acidic stomach contents into the duodenum causes gall bladder emptying a nd secretion of pancreatic lipase and cholecystokinin (Meyer and Jones, 1974). B ile acids serve to emulsify the fats and increase the available surface area for enzy matic action, where pancreatic lipase and colipase act to hydrolyze TAG. Colipase att aches to the ester bond of the TAG, which in turn strongly binds the lipase (Patton, 1981). Pancreatic lipas e cleaves the sn-1 and sn-3 bonds specifically, leading to the formation of 2-monoacylglycerols and free FA, with small amounts of 1,2and 2,3-diacylglycerols as intermediate products (Mattson and Volpenhein, 1964). Although pancreatic lipas e attacks primarily at stereospecific locations, the relative rate of hydrolysis depends on the FA pres ent. The lipase has much slower activity when long-cha in FA, particularly the n-3 polyunsaturated FA (20:5 and 22:6), are located in the sn-3 posi tion (Ikeda et al., 1995). Additionally, 2-monoacylglycerols can isomerize to 1-monogl ycerides to a small extent in aqueous conditions, allowing for the formation of a sm all percentage of glycerol and free FA (Mattson and Volpenhein, 1962). Phospholipids undergo a similar hydrolysis as TAG, however the specific enzyme, phospholipase A2, cleaves FA from the sn-2 position of the phosphoglyceride (van Deenen and deHass, 1963). Dietary cholesterol enters the duodenum as both free and esterified cholestero l. Prior to absorption, the esterified cholesterol is hydrolyzed to free cholestero l and NEFA by choleste rol esterase (Hyun et

PAGE 38

20 al., 1969). Cholesterol esterase may also ai d in the hydrolysis of TAGs that contain long-chain PUFA (Carlier et al., 1991). Lipid absorption in humans begins in the distal duodenum and is completed in the jejunum. Non-esterified fatty acids and 2-monoacylglycerols, along with phospholipids, enter into bile micelles, forming mixed micelle s. This solubilization allows the non-polar lipids to travel through the unstirred water laye r and reach the brush-border membrane of the enterocyte (Dietschy et al., 1971; Wilson et al., 1971). The pH of the unstirred water layer promotes protonation of NEFA, allowi ng them to more easily leave the micelles and move to the epithelial cell membrane. Once in close contact with the brush-bor der, the 2-monoacylglycerols, NEFA, and free cholesterol cross the microvi llus membrane. In the past, it had been thought that FA pass into the enterocyte via passive diffusion due to high intraluminal and low cytosolic concentrations of lipids (Keelan et al., 1992). More recently, however, it has been proposed that a specific transp ort protein facilitate s the movement of FA into the cell. Two such proteins that may be involved in intestinal lipid transport are plasma membrane fatty acid binding protein and fatty acid tran slocase (Frohnert and Bernlohr, 2000). Bile salts and some cholesterol are not absorbed and pass to the ileum, where they are recycled via the porta l blood to the liver. Once within the entero cyte, FAs are re-esterified into TAG and phospholipids in a multi-step process. First, FAs bind to a cytosolic fatty acid binding protein (FABPc), allowing for targeting to the ER (Cartwright et al., 2000). There, acyl-CoA synthetase, a membrane-associated enzyme, activates FAs to their acyl-CoA thioesters via an ATP-dependent mechanism. This activati on effectively traps FA within the cell,

PAGE 39

21 maintaining the concentration gradient a nd increasing the rate of TAG synthesis (DiRusso and Black, 1999). Si nce the major forms of abso rbed lipids in humans and other non-ruminants are 2-monoacylglycerols an d NEFAs, resynthesis of approximately 80% of the TAG occurs via the monoacylglycerol pathway (Lehner et al., 1993). In this pathway, the first step is the acylation of 2-monoglygerides with fatty-acyl-CoA to diacylglycerols by monoacylglycerol acylt ransferase (MGAT). Monoacylglycerol acyltransferase has a preference for medium-c hain saturated and long-chain unsaturated 2-monoacylglycerols (Coleman and Haynes, 1984), but all acyl -CoA studied are incorporated with similar efficiency (B ugaut et al., 1984). The reaction produces predominantly 1,2-diacylglycerols, with onl y about 10% 2,3-diacylglycerols formed (Lehner and Kuksis, 1996). This stereospecifi city allows for the final and rate limiting step in TAG synthesis. Di acylglycerol acyltrans ferase (DGAT), which will not act on the 2,3-isomer, acetylates diacylglycerol in an acyl-CoA dependent manner (Coleman, 1988). Similarly to MGAT, DGAT shows substrat e specificity for di-unsaturated or mixed-diacylglycerols over disaturates. During fat absorption, the resynthesized TAG are packaged in the enterocyte into lipoproteins, making the lipids st able for transport in the aqueous environment of the blood. The human intestine secr etes mainly chylomicrons (CM) and very low-density lipoproteins (VLDL). During fa sting, VLDLs are the main lipoproteins secreted by the small intestine, whereas CMs are secreted during fat feeding (Ockner et al., 1969). Chylomicrons are the main route of transpor t for long-chain dietary FAs. Medium-chain FA (C<12) are absorbed in the non-esterifie d form, passing directly into the portal blood system. This occurs because shortand medium-chain FA are more likely to occupy

PAGE 40

22 position three of the TAG and are therefore hydrolyzed in the small intestine and not retained as 2-monoacylglycerols (Sethi et al ., 1993). Soon after di etary lipids enter the enterocyte, fat droplets can be seen in the ER from the formation of TAG. The rough ER is the site of synthesis of phospholipids a nd apolipoproteins, whic h provide a coat to stabilize the lipid droplet. Specifically, apoli poprotein B48 (apo B48) associates with the TAG during its synthesis, forming the immatu re CM (Cartwright et al., 2000). In the smooth ER, the immature CM accumulates furt her TAG via the actions of microsomal transfer protein (MTP). The CMs then migrate through the Golgi apparatus, where glycosylation takes place (Leblond and Benne tt, 1977) before the fully-formed CM are exported in secretory vesicles The CM-containing vesicle travels to the basolateral surface of the enterocyte, fuses with the pl asma membrane, and is secreted into the extracellular space by exocytosis (Sabesin and Frase, 1977). Very low-density lipoproteins, as mentioned above, are formed in the small intestine when the levels of lipids are too low to form CMs. Very lowdensity lipoproteins differ from CMs in their density, size, lipid content, and composition, and although both are formed in the same organelles, the two particles are not mixed in individual Go lgi vesicles (Mahley et al., 1971). Lipoproteins secreted from the intestine do not enter the blood stream directly. Instead, they are secreted in to minute lymph vessels, known as lacteals, due to their milky appearance when filled with lipid. From there, the CM and VLDL enter the circulation in the subclavian vein via the thoracic duct (Mu and Hoy, 2004). Once in the blood stream, intestinal lipoproteins come into contact with other plasma lipoproteins, where transfer of protein and TAG occurs (R edgrave and Small, 1979). In particular,

PAGE 41

23 CM and VLDL acquire apolipoprotein CII (apo C-II), which is essential for further metabolism. As CM and VLDL pass through cap illaries, they come into contact with and bind to lipoprotein lipase (LPL), which is expr essed in extrahepatic tissues that use FA, such as adipose tissue, skeletal and cardiac muscle, and the mammary gland (Ginsberg, 1998). Lipoprotein lipase, with apo C-II as a cofactor, hydrolyzes the TAG in the particle, generating NEFA that can diffuse into the tissue for further metabolism or storage (Frayn, 1998). The TAG depleted CM remnant is rapidly removed from plasma and is metabolized by the liver. Once VLDL ha ve interacted with LPL, they also lose surface apolipoproteins C and E, and become low-density lipoprotein (LDL) particles once only apo B remains. The apo B of LD L is recognized by the LDL receptor on the surface of most cells, allowing for LDL uptak e and metabolism within peripheral cells. The LDL particles are the majo r carriers of blood c holesterol in humans, pigs, and guinea pigs; however, in most mammalian species, high -density lipoprotein (HDL) serves this function. The reverse transport from peripheral cells to the liver is an important physiological process necessary to counteract the deposition of cholestero l in tissues from VLDL and LDL cholesterol. In reverse transport, HDL, primarily synthesized by the liver (Wang and Briggs, 2004), takes choleste rol from peripheral tissues a nd transports it to the liver for metabolism. In 1968, it was first rec ognized that reverse cholesterol transport involved the active transport of cholesterol, as cellular free cholesterol was converted to the insoluble ester outside of the cell. The enzyme involved in this process is lecithin:cholesterol acy l-transferase (LCAT), and is a co mponent of HDL that increases the cholesterol esters within this lipoprotein fraction (Glomset, 1968). The rate of LCAT

PAGE 42

24 is affected primarily by the surface properties of individual lecithin molecules (Pownell et al., 1985). Two additional proteins contri bute to HDL remodeling, both by working down concentration gradients in an energy-i ndependent manner. P hospholipid transfer protein supplies lecithin to HDL (Tollefson et al., 1988), and a choles terol ester transfer protein can move cholesterol esters made by LCAT to othe r lipoproteins, particularly LDL (Tall, 1993). The TAG portion of HDL can be catabolized by the extrcellular hepatic triacylglycerol lipase, and the chol esterol is removed by the liver via several different mechanisms (Nagata et al., 1988; Wa ng and Briggs, 2004). It is only tissues that actively uptake or synthesize cholesterol that contribute to the reverse cholesterol transport pathway. Dietary Fats in Relation to Health Dietary Fats in Relation to Weight Control According to the World Health Organizati on (World Health Re port, 2002), obesity rates have risen over three-fold since 1980 in most developed and developing countries worldwide. Current estimates count more than one billion adults as overweight and at least 300 million as clinically obese. In the US, approximately 30% of adults are categorized as obese, which is defined as at least 20% heavie r than their ideal weight. Obesity is associated with increased early mort ality and an increased risk for a variety of diseases, including metabolic, cardiovascular an d gastrointestinal di seases. Because of this, the World Health Organiza tion has listed obesity as one of the top ten global health problems in Western cultures (World Health Report, 2002). Analysis of epidemiological data suggests that dietary fat plays a role in obesity, though the mode of action is not clear (Bray and Popkin, 1998; Astrup et al., 2000). It is evident however, that it is not a simple re lationship. Longititudinal measurements of

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25 food intake in both the US and the UK show that fat intake has not increased as a proportion of dietary energy over the last 30 years, unlike th e rising trends in obesity (Heini and Weinsier, 1997; Niel sen et al., 2002). Obesity may be due to the types of fat consumed or the interaction of dietary fats or FAs with other dietary compounds. If high intakes of dietary fat are a fact or in the development of obesity, then reducing the fat in the diet w ould be expected to produce wei ght loss. Studies examining the relationship between fat in the diet and changes in body weight reported several conclusions. When animals were fed a high fa t diet, almost all sp ecies develop obesity, as demonstrated in primates, rodents, pigs, dogs, and cats (West and York, 1998). Exceptions include animals with a strong genetic component to obesity, such as C57/BL mice and Osborne-Mendel rats (Bray et al., 2004 ). Reductions in body weight of subjects consuming a low-fat diet are modest and te nd to extend over only a short period of time (Jeffery et al., 1995), and the higher the body mass index (BMI) of the subject, the greater the weight loss (Astrup et al ., 2000). There is also a pos itive relationship between the percent reduction in dietary fat and the decr ease in energy intake, suggesting that the major mechanisms for weight loss associated with reduced-fat diets may be primarily through a lower energy intake (Bra y et al., 2002). Dietary fat, therefore, may not be an independent cause for obesity. Excessive energy intake, whatever the source, and decreased energy expenditure generally are cons idered the main causes of obesity (Foreyt and Poston, 2002). Current recommendations sti ll call for a low-fat intake, due to fat’s higher caloric density than other nutrients, coupled with less energy expenditure of an increasingly sedentary populati on (Astrup et al., 2002). When this theory is considered, it is evident that the type or composition of dietary fat may have li ttle effect on obesity.

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26 However, the type of fat consumed can play significant, differential, and more direct roles in other health and disease states. Dietary Fats and Blood Cholesterol Dyslipidemia is a condition in which plas ma concentrations of LDL cholesterol and TAG are elevated and HDL cholesterol is lower than found in normal, healthy individuals. According to the most recent guidelines set by the US National Cholesterol Education Program (NCEP), total choleste rol should be <200 mg/dL, with LDL cholesterol <100 mg/dL and HDL choles terol >40 mg/dL (Grundy et al., 2004). Abnormally high LDL cholesterol and low HDL cholesterol outside the recommended values are considered significant risk fact ors for cardiovascular disease, therefore maintaining optimum blood concentrations is be neficial. Dietary fat has the ability to modify blood cholesterol components in both a positive and negative manner, depending on the types of fat consumed. As a group, consumption of saturated FAs raises total and LDL cholesterol in blood, but individual saturated fats can have differing effects (Reddy and Katan, 2004). Several feeding studies have demonstrated that individuals consuming diets high in saturated fat had increased concentrations of both HDL and LDL cholesterol (Kromhout et al., 1995). Myristic (14:0) and lauric (12:0) acids have a greater effect on elevating LDL cholesterol than palmitic acid (16:0), but, among these, palmitic acid is greatest in the food supply. Stearic acid, in contrast, decreases plasma and liver cholesterol concentrations, primarily by reducing intestin al cholesterol absorption. The mechanism by which stearic acid reduces cholesterol is thought to be by reduc ing solubility of cholesterol and alteri ng the population of microflora that can synthesize secondary bile acids (Cowles et al., 2002).

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27 In the human diet, the predominant monouns aturated fatty acid (MUFA) is oleic acid (18:1, n-9). It is found at high levels in olive oil, canola oil, and nuts. The Mediterranean diet, which is not low in fat but is associated with a healthy blood lipid profile, contains a high percentage of fat as ol eic acid. When saturated fats in the diet are replaced with oleic acid, total and LDL chol esterol concentrations are lowered (Gardner and Kramer, 1995). This seems to be caused by a passive mechanism; when saturated fats are decreased and MUFA are increase d, the fat induced-suppression of LDL receptor activity is less and LDL uptake into cells is in creased (Dietschy et al ., 1993). Effects of MUFA on HDL cholesterol are less clear. Some studies have indicated that MUFA have no effects on blood HDL concentrations (Delap lanque et al., 1991; Ma ta et al., 1992). This combined with the LDL-lowering e ffects suggest that MUFA can shift the LDL:HDL ratio towards a healthier profile. However, upon extensive meta-analysis, the effects of MUFA on HDL cholesterol in blood could not be confirmed (Gardner and Kramer, 1995). Polyunsaturated fatty acids of the n-6 fam ily, particularly LA, lower total and LDL cholesterol when they are supplied in the diet in place of saturated fats (Kris-Etherton and Yu, 1997). In addition to the passive mech anism described for MUFA, PUFA actively increase receptor-dependent LDL uptake, altho ugh this is a small effect (Dietschy et al., 1993. Dietschy, 1998). In some studies in wh ich n-6 PUFA replaced saturated fats, a significant decrease in HDL cholesterol was re ported (Shepherd et al., 1978; Jackson et al., 1984), although this is not a consistent effect (Iacono and Dougherty, 1991). When n-3 PUFA were supplemented w ith the regular diet, LDL chol esterol was raised in some studies (Harris et al., 1988; Fumeron et al., 1991) and HDL cholesterol was either

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28 unchanged or slightly increased (Harris, 1989). These effects tend to be more pronounced in hyperlipidemic subjects. For example, an increase in LDL cholesterol occured in isolated hypertrig lyceridemiac subjects when more than 10 g of n-3 FA were supplemented per day (Schmidt and Dyerber g, 1994). When very long-chain n-3 FA, such as EPA and DHA were specifically suppl emented, they not only have the ability to significantly lower serum TAG, but also to increase LDL cholesterol more so than supplementing with LNA or a mixt ure of n-3 FA (Harris, 1997). Trans -FA are geometrical isomers of unsaturated FA that assume a saturated fatlike configuration. The predominant source in the human diet is from hydrogenated oils, such as margarine and partially hydrogena ted soybean oil, commonly found in baked goods and deep fat-fried fast foods (Hu et al ., 2001). Metabolic studi es have shown that consumption of trans -FA has the ability to negatively alter the lipid profile to a greater extent than saturated fats, because they not only increase small, dense LDL cholesterol (Mauger et al., 2003), but also decrease HDL c holesterol (Judd et al., 1994; de Roos et al., 2003). This leads to an in crease in the ratio of total to HDL cholesterol that is approximately double that observed with sa turated fats (Willett and Ascherio, 1994). Additionally, diets high in trans-FA are asso ciated with raised TAG concentrations (Katan and Zock, 1995), an independent risk marker of cardiovascular disease. Dietary Fats and Cardiovascular Disease Most of the FAs in the Western diet ar e derived from meats, oils, and dairy products, leading to a large intake of satu rated and MUFAs, with a relatively small proportion of PUFA consumed. Saturated fat and cholesterol repres ent two of the most established dietary risk fact ors for cardiovascular dis ease (CVD), whereas MUFA and PUFA are likely to provide beneficial effect s with increased amounts in the diet. These

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29 effects are partially due to the effects on the blood lipid profile, but the risks associated with intake of certain fats are greater than would be exp ected from cholesterol effects alone. As stated above, intake of certain sa turated fats can increase LDL and decrease HDL cholesterol, creating an atherogenic lipid profile. A recent analysis of the Nurses’ Health Study revealed that intake of short to medium chain FA was not associated with increased coronary heart disease (CHD) risk. In the same analysis, however, intakes of longer chain saturates, particularly stearic acid, were associated with an increased risk of CHD (Hu et al., 1999). Additionally, stearic acid may negatively impact other markers of atherogenesis. Stearic acid can increase lipoprotein(a) con centration (Aro et al., 1997), and may activate Factor VII (Mitropoulos et al., 1994) and impair fi brinolysis (Ferguson et al. 1970). On the positive side however, when compared with consumption of palmitic acid, stearic acid decreases plat elet volume, platelet aggreg ation, and coagulation factor VII activity (Kelly et al., 2001). Due to the many negative health imp lications of a diet high in saturated fats, there is a consensus to reduce the intake of saturated fats to less than 10% of the total daily energy suppl y (American Heart Association, 2000). The Seven Countries Study gave the first epidemiological evid ence for a negative correlation between dietary intake of MUFA and mortality from CHD. Mortality was noticeably low in Mediterranean countries, wher e olive oil is the main source of fat (Keys et al., 1986). In addition to the positive effects on plasma LDL cholesterol levels, diets rich in olive oil can improve endothelial function, as compared to a high saturated fat diet (Fuentes et al., 2001), and attenuate postpran dial endothelial dysf unction that follows a fatty meal (Vogel et al., 2000). Intake of MUFA is also protective against LDL

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30 oxidation. Due to their structure, MUFA are more stable and less susceptible to lipid peroxidation. A high intake of MUFA resu lts in a greater incorporation into LDL cholesterol (Mata et al., 1996). Oxidation of LDL cholesterol prevents its recognition by the LDL receptor and subsequent uptake in to cells. It is instead taken up by the scavenger receptors of macropha ges, leading to the accumula tion of cholesterol and the formation of fatty streaks. These processe s promote the development of atherosclerosis (Westhuyzen, 1997). Several studies have s hown that dietary sour ces of MUFA other than olive oil are associated w ith an increased CHD risk (Pos ner et al., 1991; Esrey et al., 1996). However, these studies did not correct for potential confounding effects, such as the intake of other FAs and antioxidants. Epidemiological evidence supports a role fo r dietary LA in reducing the risk of CHD. High adipose LA in healthy men is associated with lower CHD mortality (Riemersma et al., 1986), while low dietary intake of LA predisposes to myocardial infarction (Simpson et al., 1982). A more recent study of Japanese subjects found reduced serum LA in patients with ischemic st roke as compared to healthy controls (Iso et al., 2002). Similarly to LA, LNA intake was inversely associated with mortality from CHD in the Multiple Factor Intervention Trial (Dolecek, 1992). Large prospective studies in both men and women have found that LNA protected against both cardiac deaths and nonfatal myocardial infarction (Ascherio et al., 1996; Hu et al., 1999). The effects of LNA on plasma lipids are not larg e; therefore the reduction in CHD risk may have more to do with cardiac functio n, such as arrhythmia, inflammation, and thrombosis.

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31 The low rate of CVD seen in several co mmunities consuming a diet rich in fish (Bang et al., 1980; Kromhout et al., 1985; Hirai et al., 1989; Oomen et al., 2000) has prompted investigations into how fish and its nutritional component s may lower the risk of CVD. Fish, particularly fatty fish such as tuna, mackerel, and salmon, are rich in the n-3 FA EPA and DHA (Parkinson et al., 1994). High serum and adipose tissue long-chain n-3 PUFA have been associated with reduced risk of fatal myocardial infarction (Simon et al., 1995; Pedersen et al., 2000; Lemaitre et al., 2003), primary cardiac arrest (Siscovick et al., 1995), and sudden cardiac death (Albert et al., 2002). Upon meta-analysis of 11 randomized controlle d trials comparing long-chain n-3 PUFA intake to placebo or control diets, intake of long-chain n-3 PUFA was associated with lower cardiac fatalities in patients with C HD (Bucher et al., 2002). However, these FA did not protect against nonfatal cardiac even ts or total morbidity (Erkkila et al., 2003; Lemaitre et al., 2003), suggesting that th e hypolipidemic effects of DHA and EPA on atherosclerosis are distinct fr om those effects associated with arrhythmic myocardial dysfunction. In a canine m odel with dogs made susceptible to fatal ventricular fibrillation and sudden cardiac death, infusi on of EPA and DHA reduced cardiac deaths by preventing ventricular fibrillation (Billman et al., 1997; Billman et al., 1999). As the fat infusion was given only one hour prior to inducing ischemia, the effects are not likely by membrane incorporation of n-3 PUFA, but rather by direct acti on of nonesterified PUFA on the myocytes. In support of this, i nduction of arrhythmias in cultured neonatal rat myocytes was abolished by the addition of EPA or DHA to the culture medium (Kang and Leaf, 1996), and supplementation of four g/d of EPA and DHA increased heart rate variability in survivors of myocardial infarction, reduci ng the risk of subsequent

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32 arrhythmic events (Christensen et al., 1996) The anti-arrhythmic actions of EPA and DHA seem to be associated with the ability of these FA to prevent calcium overload in cardiac myocytes during periods of stress (Leaf and Kang, 1997). As stated previously, trans -FA negatively impact the bl ood lipid profile in several ways. However, the relations hip between consumption of trans -FA and cardiovascular risk is greater than is predicted based on these lipid changes (Ascherio et al., 1999), suggesting effects on other distinct risk mark ers for CVD. In humans, it has been shown that trans -FA increase lipoprotein (a) le vels (Nestel, et al., 19 92; Sundram et al., 1997), which are positively associated with in creased risk of CHD (Utermann, 1989). Additional studies have examined the effects of trans -FA on markers of low-grade chronic inflammation. The Nurses’ Health Study showed that a high intake of trans -FA was positively associated with concentrations of tumor necrosis factor receptors 1 and 2 (Mozaffarian et al., 2004). A dietary interv ention study in which 8% of dietary energy from carbohydrates, oleic acid, or stearic plus trans -FA was replaced with trans -FA supported this epidemiological data, and additionally found increases in plasma C-reactive protein, IL-6 a nd E-selectin with the trans -FA diet (Baer et al., 2004). Conjugated Linoleic Acid Conjugated linoleic acid (CLA ) is the collective term for a group of positional and geometric conjugated dienoic isomers of LA These FA are considered conjugated because, unlike other FA, the double bonds occur on adjacent carbons, and are not separated by a methylene group. To date, 16 CLA isomers have been identified (Rickert et al 1999), with double bonds ranging in positio n from carbons 6 and 8 to carbons 12 and 14. The double bonds can occur in pairs of geometric isomers as cis cis cis trans trans cis and trans trans However, only two isomers ( cis -9, trans -11 CLA and

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33 trans -10, cis -12 CLA) are known to possess biologi cal activity (Pariza et al., 2000). Sources of CLA in the human diet are ru minant products and synthetic supplements, though the specific makeup of CLA differs among sources. In milk, cheese, and ruminant meat, which can contain 2-8 mg of CLA/g lipid depending on the source (Lin et al 1995; Chin et al., 1992), approximately 80% of the CLA is cis -9, trans -11 CLA and 10% is trans -10, cis -12 CLA (Fogerty et al., 1988). Base d on this, recent studies suggest average intakes of 150-200 mg of CLA per day (Jiang et al., 1999; Ritzenthaler et al., 2001), with intakes as high as 650 mg/day on a diet rich in animal fats (Park et al., 1999a). Conjugated linoleic acid dietary supplements, produced by the chemical isomerization of LA, contain predominantly cis -9, trans -11 CLA and trans -10, cis -12 CLA in equal amounts (Gaullier et al., 2002). The cis -9, trans -11 CLA isomer is produced as an intermediate in the rumen during the biohydrogenation of dietary LA. A key anaerobic bacterium in this process is Butyrivibrio fibrisolvens (Kepler et al., 1966). The cis -12 bond of LA is acted upon by the microbial isomerase, forming cis -9, trans -11 CLA. In some of the literature, this isomer is also referred to as rumenic acid. This CLA product can leave the rumen and be directly absorbed, or it ca n be further metabolized by ru minal microbial hydrogenases, forming trans -vaccenic acid ( trans -11, 18:1) before being co mpletely hydrogenated to stearic acid (Kepler et al., 1966). This pr oduct may also exit the rumen and be absorbed and transported to periph eral tissues. In the mammary tissue and muscle, a 9-desaturase is present that can act on trans -vaccenic acid to produce cis -9, trans -11 CLA (Holman and Mahfouz, 1980; Pollard et al., 1980). Th is has been shown to occur in several mammalian species, including ruminants (Griin ari et al., 2000), mice (Santora et al.,

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34 2000), and humans (Turpeinen et al., 2002). Certain ruminal bacteria also have the capability to convert LA to trans -10, cis -12 CLA by isomerizing the cis -9 bond (Griinari and Bauman, 1999). This can be hydrogenated to form trans -10 octadecenoic acid, which may be absorbed and transported to peripheral tissues, but since mammals do not possess a 12-desaturase, it would not be converted back to trans -10, cis -12 CLA. Numerous beneficial physio logical effects have been attributed to CLA. The seminal observation came when CLA isolated from grilled beef inhibited chemically-induced skin neoplasia in mice (Ha et al., 1987). This discovery led to research examining the effects of CLA on cancer (Ha et al., 1990), immune function (Miller et al., 1994), atherosclerosis (Lee et al., 1994), weight gain and food intake (Chin et al., 1994), and body composition (Park et al ., 1997). As previously stated, the two biologically active isomers of CLA are cis -9, trans -11 and trans -10, cis -12 CLA. Though derived from the same parent molecu le, the two isomers are structurally and functionally distinct. Both isomers contain a trans double bond, creating a straighter carbon chain, as opposed to the “kink” created by the cis configuration. Many enzymes recognize specific configurations in FA; therefore it is not su rprising that differences in bond position and orientation of CLA isomers give them differing biological activities. Numerous studies now indicate that the various physiological and bi ological effects of CLA may be due to the se parate actions of the cis -9, trans -11 and trans -10, cis -12 isomers (Pariza et al., 2000). Dietary CLA modulates body composition through decreases in adiposity and increases in lean mass in various animal models. The effects in mice are the most dramatic, with a 50-60% reduction in total ad ipose mass in animals fed mixed isomers of

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35 CLA over a 4-5 week period, as compared to mi ce fed the control diet (Park et al., 1997). The effect in mice can be sustained, even afte r removal of CLA from the diet (Park et al. 2001). Additionally, when the trans -10, cis -12 isomer was fed to mice, it was more effective in lowering adipose tissue mass than cis -9, trans -11 CLA (Park et al., 1999b). Similar reductions in adipose mass have been noted in Sprague-Dawley and Zucker lean rats fed CLA, although the effects are not as large as in mice (25-30% reduction) (Sisk et al., 2001). In contrast to lean rats, obese Zuck er rats exhibit an ad ipose-enhancing effect of dietary CLA (Szymczyk et al., 2000). In pigs, CLA-feeding decreased fat deposition and increased lean tissue (Dugan et al., 1997; Thiel-Cooper et al., 2001). In humans, however, the results are not as clear. Several studies have shown no effects of mixed CLA or individual isomers in the diet on ch anges in body composition in human subjects (Terpstra, 2004), and to date, no studies ha ve shown changes in body weight (Larsen et al., 2003). Conversely, studies feeding mixed CLA and the trans -10, cis -12 isomer have reported reductions in body fat mass but no changes in body mass index (Blankson et al., 2000; Smedman and Vessby, 2001; Riserus et al., 2004). These changes are much less than those observed in pigs and mice; howev er, pigs and mice are generally fed at least five times more CLA per kilogam of body weight than humans (House et al., 2005). Comparable doses as used in animal studies would correspond to a daily intake of 130 g in humans (Larsen et al., 2003). Long term supplementation with mixed CLA isomers by healthy overweight individuals also seems to be well to lerated, although reductions of body fat mass may or may not be maintained (G aullier et al., 2005; Larsen et al., 2006). Mechanisms by which CLA reduces adiposit y may involve pathways that involve energy expenditure. This is shown by incr eased metabolic rates and reduced nighttime

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36 respiratory quotients in mice fed CLA for si x weeks (West and York, 1998). Effects of CLA also have been linked with the induction of adipocyt e apoptosis, both in vivo (Tsuboyama-Kasaoka et al., 2000) and in vitro (Evans et al., 2000), and with decreased uptake of TAG into adipocytes, particularly due to the suppressi on of lipoprotein lipase activity by trans -10, cis -12 CLA (Park et al., 2001). Obesity puts an individual at a greater risk for other diseases, including type II diabetes. It would be expect ed then, that reduction in fa t mass due to CLA intake would help decrease this risk. Evidence, however s upports an additional di rect effect of CLA on diabetes, which can vary, depending on the sp ecies studied. In animal models of diabetes, such as the Zucker diabetic fatty ra t, CLA-enriched diets reduce fasting glucose, insulinemia (Houseknecht et al., 1998), tryg lyceridemia and blood NEFA concentrations (Belury and Vanden Huvel, 1999) as compared with controls. These beneficial effects may be due, in part, to enhanced muscle upt ake of glucose (Ryder et al., 2001). It is important to note that these effects are seen when a mixture of CLA isomers are fed. When fed butter enriched with cis -9, trans -11 CLA, little or no effect was seen, indicating that the effects on glucos e tolerance are likely due to the trans -10, cis -12 isomer (Ryder et al., 2001). In contrast with diabetic animals, CLA modestly increases fasting serum insulin in nondiabetic pigs (S tangl et al., 1999 ), mice (Tsuboyama-Kasaoka et al., 2000), and humans (Medina et al., 2000). These negative effects on insulin resistance may result from decreased plasma leptin concentrations (Wang and Jones, 2004) or an increase in TAG concen tration in muscle due to feeding trans -10, cis -12 CLA (Terpstra, 2004).

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37 Similar to other classes of FA, CLA can affect the blood lipid profile and cardiovascular risk factors. In rabbits fed an atherogenic diet, supplementation with CLA lowered serum TAG and LDL cholesterol concentr ations, as compared to controls (Lee et al., 1994). These animals fed CLA also showed a decrease in atherosclerotic plaque formation. Another study with rabbits showed a regression of establis hed atherosclerosis, despite an increase in total cholesterol and de crease in HDL cholesterol (Kritchevsky et al., 2000). In a similar model in hamsters fed the cis -9, trans -11 isomer, there was no effect on plasma lipids (Gavino et al., 2000). Culturing of human platelets with either CLA isomer inhibited induced platelet aggr egation (Truitt et al., 1999), but in human subjects supplemented with a mixture of CLA isomers, no difference was observed in platelet aggregation or prothr ombin time (Benito et al., 2001 ). Together, these findings potentially implicate trans -10, cis -12 CLA to have positive effects on the blood lipid profile and coronary risk factors. In addition to the effects on disease states CLA can alter lipid metabolism. When consumed, CLA is incorporated into membra ne phospholipids and alters FA homeostasis, particularly in the liver (Bel ury, 2002). Conjugated linoleic acid that is not broken down via -oxidation is desaturated and elongated to other conjugated meta bolites (Belury and Kempa-Steczko, 1997). The competition of CLA with LA for 6-desaturase may result in decreased AA, and can explain the redu ced eicosanoid production in several systems (Belury, 2002; Brown and McIntosh, 2003). It has also been found that mice (Degrace et al., 2003) and hamsters (de Deckere et al., 1999) supplemented with CLA, particularly the trans -10, cis -12 isomer, develop enlarge d, fatty livers. This eff ect has been attributed to an increase in liver TAG, cholesterol, ch olesterol esters, and NE FA (Kelley et al.,

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38 2004), though the mechanism is unclear. Supplementation with trans -10, cis -12 CLA in vivo and in vitro in various animal and human models leads to an increase in the ratio of saturated to monounsaturated fats (House et al ., 2005). This is likely due to a reduction in stearoyl-CoA desaturase, which catalyzes the biosynthesis of MUFA from stearic and palmitic acids (Lee et al., 1998). Trans -10, cis -12 CLA also inhibits transcription of other genes involved in de novo FA synthesi s, desaturation, and TAG synthesis, which may partially explain its effects on changes in lipid metabolism in the liver (Baumgard et al., 2002). Roles of the Peroxisome Proliferator-Activated Receptors in Lipid Metabolism Peroxisome proliferator-activated recept ors (PPAR) belong to the steroid hormone receptor superfamily that are ligand-activated transcription factors (Wahli and Martinez, 1991), and act by modulating a network of respon sive genes. They have been identified in many species, including Xenopus (Dreyer et al., 1992), mouse (Issemann and Green, 1990), rat (Gottlicher et al., 1992), and huma n (Sher et al., 1993). The name PPAR derives from the ability of the first-iden tified member to induce hepatic peroxisome proliferation in mice, but this phenomenon se ems to be rodent-specific, and does not occur in other mammals. The PPARs consist of a family of three isoforms: PPAR , and – / (Issemann and Green, 1990; Dreyer et al., 1992; Kliewer et al., 1994). Though encoded by separate genes, and different in their tissue dist ribution and metabolic actions, all three isoforms are structurally similar a nd can be activated by FA and their metabolic derivatives, making them the first recognized lipid sensors in the body (Schoonjans et al., 1996). Genes whose expression is modified by PPARs are nu merous and control glucose homeostasis, cell cycle, inflammation, immune response, and lipid metabolism (Desvergne and Wahli, 1999).

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39 Similar to other nuclear receptors, the PPA Rs possess structural features composed of functional domains. The DNA-binding domai n consists of two zinc fingers that specifically bind peroxisome pro liferator response elements ( PPRE) in enhancer sites of regulated genes (Wahli and Martinez, 1991) The PPRE are specific DNA sequences formed by the direct repeat of a hexanuc leotide sequence (AGGTCA), separated by one or two nucleotides (Torra et al., 2001). Un like other steroid receptors which function as homodimers, to bind to the PPRE, PPAR must form a heterodimer with the retinoid X receptor (RXR) in the cytoplasm, allowing for transport to the nucleus (Miyata et al., 1994). The ligand binding domain appears to be quite large in comparison with other nuclear receptors (Nolte et al., 1998; Xu et al., 1999), potentially allowing PPARs to interact with a broad range of structural ly distinct natural and synthetic ligands. As PPARs play a critical role in lipid metabolism, the search for natural ligands began with the FAs and eicosanoids. Cell-base d transactivation assays and direct binding studies have identified and characterized the endogenous receptor effectors. In general, all isoforms of PPAR are more responsive to n-6 and n-3 PUFA than to saturated or monounsaturated FAs (Krey et al., 1997). Howeve r, the affinities for the receptor vary, suggesting a role for site-speci fic availability and metabolism of particular FA, as well as different affinities for the specific PPAR is oforms (Sampath and Ntambi, 2005). It has been shown that FA such as LA, LNA, and AA can activate PPAR at a concentration of 100 M (Lehmann et al., 1997). Additionally, EPA is a much more potent activator of PPAR than arachidonic acid in primary hepa tocytes (Ren et al., 1997). Since the concentration of NEFA in human blood can be greater than 1 mM, these FA can be considered potent endogenous ligands for PPAR It is important to note, however, that

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40 the intracellular concentrations of PUFA are not known. Like PPAR PPAR has affinity for the PUFAs, as well as metabolic derivatives of PUFAs, such as 9-hydroxyoctadecadienoic acid (HODE) and 13-HODE (Nagy et al., 1998), and CLA (Hontecillas et al., 2002). Peroxisome proliferator-activated receptor also interacts with saturated and unsaturated FA, but with a liga nd specificity that is intermediate between that of PPAR and PPAR (Berger and Moller, 2002). Even with the abundance of natural ligands for PPARs, the emphasis in r ecent years has been on the development of synthetic ligands, due to their greater therapeutic and commerci al value. Fibrates, which are ligands for PPAR and thiazolidinediones (TZD), which are ligands for PPAR are two classes of drugs used to treat hy pocholesterolemia and type II diabetes. PPAR The first PPAR discovered (Issemann and Green, 1990), PPAR is expressed predominantly in the liver, kidney, heart, brown fat, and skeletal musc le (Braissant et al., 1996; Auboeuf et al., 1997), as well as in m onocytic (Chinetti et al., 1998), vascular endothelial (Inuoe et al ., 1998), and vascular smooth muscle cells (Staels et al., 1998). It plays an important role in lipid metabolis m via regulation of the expression of genes involved in cellular free FA uptake, -oxidation, and cellular chol esterol trafficking (Li et al., 2002). It has been reported that PPAR is greatly induced duri ng fasting or starvation in which a switch from carbohydrates and fats to mostly fats as an energy source is required. During fasting, FA released from the adipose tissue are taken up by the liver, where they are re-esterified to TAG or broken down via -oxidation to ketones. Peroxisome proliferator-activated receptor induces expression of fatty acid translocase (Motojima et al., 1998) and fatty acid transp ort protein (Martin et al., 1997), genes involved in transport of FA into the cell, as well as CPT-I (Brady et al., 1999), which

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41 catalyzes the rate limiting step for transport of FA into the mitochondria for oxidation. Activation of PPAR also directly upregulates genes involved in peroxisomal -oxidation, including acet yl-CoA synthase (Schoonj ans et al., 1995) and ACO (Tugwood et al., 1992). The importance of PPAR in this response has been demonstrated by studies involving PPAR -null mice, which are unable to induce the change in energy source, resulting in hypoglycemia, hyperlipidemia, hypoketonemia and fatty liver (Kersten et al., 1999). In rodents, activation of PPAR induces peroxisome proliferation, hepatomegaly, and hepato carcinogenesis (Issemann and Green, 1990). Fortunately, these effects are not present in humans, possibly due to the 10-fold greater concentrations of PPAR in rodent as compared to human liver (Palmer et al., 1998) or to differences in the PPREs of responsive genes, such as ACO (Lambe et al., 1999). Fibrates have been a commonly prescribed drug to treat dyslipidemia in humans for over 30 years, but the direct role of PPAR in the lipid-lowering ac tions of fibrates has only recently been established. In humans fibrate administrati on lowers plasma TAG and increases plasma concentrations of HD L and its major constituents, apolipoproteins A-I (apo A-I) and A-II (apo A -II) (Malmendier and Delcroix, 1985; Mellies et al., 1987). Peroxisome proliferator-activated receptor activation affects seve ral key genes in HDL metabolism, including apo A-I, apo A-II, ABCA1, LPL, and scavenger receptor class B type I (Fruchart, 2001). Peroxisome proliferator-act ivated receptor also has been shown to down-regulate apo C-III (Hertz et al., 1995; Staels et al., 1995), a protein that inhibits TAG hydrolysis by LPL, further contributing to the lipid-lowering effects of fibrates.

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42 Surprisingly, the role of PPAR in cardiovascular disease appears to be negative. In a mouse model, over-expression of PPAR in the heart increases FA oxidation and decreases glucose use, similar to that s een in the diabetic heart. Upon fibrate administration, these mice develop greater ca rdiomyopathy than the wild-type controls (Finck et al., 2002). Peroxisome proliferator-activated receptor null mice do not show this effect (Finck et al., 2003). This know ledge, when combined with research indicating that PPAR and apolipoprotein E double knockout mi ce are resistant to insulin-resistance and atherosclerotic lesions induced by a hi gh-fat diet (Tordjman et al., 2001), suggests that PPAR senses FAs and induces their use, ther eby playing a potential causative role in cardiovascular disease. Unlike humans, in rodent models, fibrate administration decreases apo A-I and apo A-II expression, suggesting differential regulation in the different species (Berthou et al., 1995). Ov erall reduction in TAG and increase in HDL cholesterol in humans, even with potential fo r negative cardiovascular events, would still result in less fat accumulation in the vessel wall s, and would be beneficial to heart health. PPAR / Peroxisome proliferator-activated receptor / (hereafter referred to as PPAR ) has been slighted in its importance in the body because of its ubiquitous expression and unavailability of selective ligan ds, despite the fact that it is the predominant isoform in skeletal muscle – one of the most insulin responsive and metabolically demanding tissues of the body. The importance of PPAR in FA metabolism was first realized from studies using knockout animals. Most PPAR null mice die during early embryogenesis, and the numbers that do survive show a marked decrea se in fat mass (Peters et al., 2000). In exercised or fasted PPAR null mice, the liver, but not the muscle glycogen levels

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43 deplete as compared to wild-type litter mate s, indicating that a factor other than PPAR may be in control of energy homeostasis (Muoio et al., 2002). Recently, synthetic, highly selective PPAR -agonists have been developed, and its role in FA catabolism and energy homeostasis has been further elucidated (Peters et al., 2000; Barak et al., 2002). Activation of PPAR increases FA oxidation in human and rodent myocytes, showing the redundancy of PPARs and in FA homeostasis (Muoio et al., 2002). In geneti cally obese ob/ob mice, PPAR activation not only enhances -oxidation in skeletal muscle, but prot ects against diet-induced obesity, improves glucose tolerance, and improves insulin sensit ivity, showing its potenti al as a target in treating and preventing obesity and type II diabetes (Tanaka et al., 2003). Similarly to PPAR PPAR activation up-regulates adenosine triphosphate-binding cassette tr ansporter-A1 (ABCA1) gene expression and increases cholesterol efflux from cells and increases HDL cholesterol in mice (Leibowitz et al., 2000) and non-human primates (O liver et al., 2001). The PPAR selective agonist GW501516, in particular, has shown therap eutic potential for the treatment of dyslipidemia, by dramatically improving the serum lipid profile of insulin-resistant rhesus monkeys. This occurs through d ecreases in concentrations of blood TAG and insulin and increases in HDL to a greater extent than is achieved with fibrates in fasting individuals. Activation of PPAR has the added normal-lipidemic effect of lowering the blood concentrations of small dense LD L cholesterol (Oli ver et al., 2001). Recent research suggests a role for PPAR in the heart as well. In cultured human cardiomyocytes, PPAR is highly expressed, and its activa tion leads to an increase in FA oxidation (Cheng et al., 2004), leading to a potential increase in energy to an energy

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44 demanding organ, and implicating PPAR as a modulator of cardiac energy homeostasis. Regarding foam cell formation, research with different PPAR activators have given different results. In one study, activation in creased cholesterol e fflux through the ABCAI pathway (Oliver et al., 2001), whereas a nother study demonstrated enhanced lipid accumulation (Vosper et al., 2001). Though these discrepancies may be due to different experimental models and stru cturally different agonists, further research can help elucidate the role of PPAR PPAR Because it is primarily f ound in adipose tissue, PPAR is a prime suspect in the regulation of lipid metabolism. In suppor t of this, many studies have shown the importance of PPAR in the formation and functioning of adult fat cells (Rosen et al., 2000). As obesity is a primary risk factor fo r incidence of the meta bolic syndrome, it is highly likely that PPAR plays a role in the associated diseases and their treatment. Thiazolidinediones are pharm acologic activators of PPAR which significantly improve insulin sensitivity in humans with type II diabetes (Sood et al., 2000). The mechanism of action, however, still remains unclear, especially considering the fact that muscle is the major insulin responsive tissue, and PPAR is present at very low levels in muscle and liver and high in adipose. Resolving this apparent paradox had pr oved difficult. Most research has stemmed from clinical trials a nd rodent models of obesity and diabetes. Unlike the readily available PPAR null mice, PPAR knockout mice die early in gestation, preventing valuable loss-of-function st udies (Barak et al., 1999). By the use of microarrays for gene expression profiling, se veral key metabolic genes were identified, all primarily in the adipocyte. Changes induced by TZD administration to Zucker

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45 diabetic fatty rats include modulation of ge nes involved in glucose uptake, lipid uptake and storage, and energy expend iture (Way et al., 2001). Th e small increases in glucose disposal by the adipose, coupled with greater sequestration of fat into adipose, thereby relieving some of the metabol ic burden of muscle and liver and allowing for greater glucose use by these tissues, is a potential explanation for the profound activity of the TZD class of drugs. In addition to the action of PPAR ligands on adipose, there is mounting evidence that these compounds can exert some effect s on other tissues. aP2/DTA mice, whose white and brown adipose tissue has been el iminated by fat-specific expression of diphtheria toxin A chain, de velop hyperglycemia, hyperinsulinemia, and hyperlipidemia indicative of insulin-resistant diabetes (Ross et al., 1993). Thiazolidinedione administration to these animals improves the serum lipid profile, but results are conflicting on the effects on glucose toleran ce, with one study showing decreases in insulin (Burant et al., 1997) and another showing no change (Chao et al., 2000). Using tissue-specific PPAR knock-out mice, the question of whether TZDs directly or indirectly affect insulin resistance has been researched. Targeted deletion of PPAR in adipose results in adipose hypertrophy, elevated plasma NEFA and TAG, increased hepatic gluconeogenesis and insu lin resistance, without change s in insulin sensitivity of muscle (He et al., 2003). These observations indicate that change s in adipose function via PPAR result in changes in he patic function with minima l effects in muscle. In addition to the effects on adipose tissu e and insulin resistance, activation of PPAR seems to play a role in atherosclerosis. Again, the positive effects of TZD treatment on decreased risk of atherosclerosi s may be secondary to the improvement in

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46 lipid profile, but PPAR activation may also have a dire ct effect on the formation and progression of atherosclerotic lesions. Pe roxisome proliferator-activated receptor activation inhibits leukocyte-e ndothelial cell interaction, a crit ical inflammatory response in the formation of atherosclerotic plaque s (Jackson et al., 1999). Activation by TZDs also inhibits the expression of vascular ce ll adhesion molecule (Pasceri et al., 2000) and E-selectin (Nawa et al., 2000), which would reduce the “homing” of monocyte and macrophage cells to atherosclerotic plaques.

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47 CHAPTER 3 EFFECTS OF N-3 AND N-6 FATTY AC IDS ON LIPID META BOLIZING GENES AND HIGH-DENSITY LIPOPROTEIN CHOLESTEROL PRODUCTION IN CULTURED HUMAN AND RAT HEPATOCYTES Introduction Dietary polyunsaturated fatty acids (PUF A) have been shown to lower blood triglycerides, alter the blood lipid profile, decrease intram uscular lipid droplet size, improve insulin sensitivity, and enhance gluc ose utilization (Jump and Clarke, 1999). Since the observation that PUFAs could inhi bit hepatic lipogenesis in mice (Allmann and Gibson, 1965), numerous studies have demonstrat ed that diets rich in PUFAs influence metabolic changes by coordinately suppressing lipid synthesis in the liver and enhancing fatty acid oxidation in both liver and skeletal muscle (J ump and Clarke, 1999). The PUFA induction of genes encoding proteins involved in lipid oxidation include 3-hydroxy, 3-methylglutaryl CoA synthase (Rodriguez et al., 1994), carnitine palmitoyltransferase, fatty acid binding pr oteins and peroxisoma l acyl-CoA oxidase (ACO; Reddy and Hashimoto, 2001). With the di scovery of a new member of the steroid hormone receptor superfamily, the peroxisome proliferator-activated receptor (PPAR; Issemann and Green, 1990) and the discovery th at certain fatty acids (FA) and their derivatives can specifically bind PPARs (Gottlich er et al., 1992), the possibility arose that PUFAs mediate metabolic effects via altera tion of PPAR activity. In the liver, the predominant isoform is PPAR ; therefore this isoform has become the primary focus of studies involving the liver.

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48 The objective of this study was to examine the short term effects of FAs of differing levels of saturation and bond position on lipid metabolizing gene expression and high-density lipoprotein (HDL) cholesterol production in He pG2 and H-4-II-E cells. Based on both dietary and in vitro studies of lipid metabolism, we hypothesized that FAs of differing saturation and double bond position may have differing effects on ACO, 3-hydroxy, 3-methylglutaryl CoA reductase (HMG -R), and apolipoprotein A-I (Apo A-I) gene expression. Also, because several fa tty acids and their derivatives are known ligands for PPARs, we hypothesized that fa tty acids may act on lipid metabolizing genes through activation of PPAR in the liver. Materials and Methods Materials Polystyrene tissue culture dishes (100 x 20 mm) were purchased from Corning (Corning Glass Works, Corning, NY). The antibiotic/antimycotic (ABAM), sodium pyruvate, fatty acid-free bovine serum albumin (BSA), stear ic acid (ST), WY 14,643, and MK886 were from Sigma Chemical Co. (St. Louis, MO). Minimum Essential Medium (MEM), phenol red-free MEM, Hanks Balanced Salt Solution (HBSS) and TriZol reagent were from GIBCO BRL (Carlsbad, CA). The fetal bovine serum (FBS) was from Atlanta Biologicals (Norcross, GA). Linoleic, linolenic, and eicosa pentaenoic acids were from Cayman Chemicals (Ann Arbor, MI). BioTrans nylon membrane and [ -32P] deoxycytidine triphosphate (SA 3000 Ci/nmol) were from MP Biolomedicals (Atlanta, GA). The Enzyme Color Solution, Reacti ng Solution, and HDL Calibrator were from Wako Diagnostics (Richmond, VA).

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49 Cell Culture and Treatment HepG2 (ATCC # HB-8065; Manassas, VA) and H-4-II-E (ATCC # CRL-1548; Manassas, VA) cells were suspended in 10 mL of growth medium (MEM), containing 2.2 g/L sodium bicarbonate, 1.0 mM sodium pyruva te, 1% (v/v) ABAM and 10% FBS. Cells were cultured at 37C in a humid ified atmosphere containing 95% O2 and 5% CO2. Cultures were replenished with fresh medium every 2 d until cells were approximately 90% confluent. Cells were washed twice with HBSS, and cultured in fresh serum-free medium containing appropriate tr eatments for an additional 24 h. Stock solutions of fatty acids were stored at -20C. At preparation of treatments, fatty acids were mixed with serum-free cu lture medium containing 33 mg/mL of fatty acid-free BSA to a concentration of 1 mM. Th is mixture was incubated for 2 h at 37C to allow complexation of the fatty acids with BSA and then further diluted in culture medium to a final treatment concen tration of 100 M of fatty acids. To investigate the effects of supplemen tal PUFAs on hepatic gene expression and cholesterol synthesis, HepG2 a nd H-4-II-E cells were treated with stearic (ST), linoleic (LA), linolenic (LNA) or eicosapentoenoic (E PA) acid (100 M). Sub-confluent cells were incubated with serum-free medium alone (Control) or with appropriate treatments (listed above) complexed with BSA, for a peri od of 24 h. Cells were rinsed twice with 10 mL of HBSS. The remaining cell monolayer wa s then lysed in 3 mL of TriZol reagent, and stored at -80C for subsequent mRNA analys is. The same fatty acid treatments were repeated, using phenol red-free MEM. After incubation, conditioned media were collected and stored at -20oC until lipid extraction and HDL cholesterol analysis. To determine whether fatty acid e ffects on gene expression involves PPAR activation, confluent HepG2 a nd H-4-II-E cells were treat ed with ST (100 M), the

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50 PPAR agonist WY 14,643 (10 M), or a comb ination of fatty acid and WY 14,643. Additional sets of culture dishes were incubated with ST alone, the PPAR inhibitor MK886 (10 M; Kehrer et al., 2001), or a co mbination of ST and MK886. After a 24 h incubation, cells were washed twice with 10 mL of HBSS, lysed with TriZol, and stored at -80oC until mRNA analysis. RNA Isolation and Analysis Total cellular RNA was isolated from cells using TriZol reagent according to the manufacturer’s instru ctions. Ten micrograms of tota l RNA was fractioned in a 1.0% agarose formaldehyde gel following previous ly described protocols (Ing et al., 1996) using the MOPS buffer (Fisher Scientific, Pitts burgh, PA) and transferred to a Biotrans nylon membrane by downward capillary transfer in 20X SSC (3 M NaCl, 0.3 M sodium citrate, pH 7.0) using the TurboBlotting sy stem (Schleicher and Schuel, Keene, NH). Nylon membranes were cross-linked by expos ure to a UV light source for 90 sec and baked at 80C for 1 h. Membranes were in cubated for 2 h at 50C in ultrasensitive hybridization buffer (ULTRAhyb; Ambion, Au stin, TX) followed by an overnight incubation at 50C in the same ULTRAhyb solution containing the 32P-labeled acyl-CoA oxidase (ACO), 3-hydroxy, 3-methylgl utaryl CoA reductase (HMG-R) and apolipoprotein A-I (Apo A -I) cDNA probes. Probes were generated by RT-PCR for ACO (forward 5’-CCGGAGCTGCTTACACACA T-3’; reverse 5’-GGTCATACGTGGC TGTGGTT-3’), HMG-R (forward 5’-TCCTTGGTGATGGGAGCTTGTTGTG-3’; reverse 5’-TGCGAACCCTTCAGATGTTTCGAGC -3’), human Apo A-I (forward 5’-AAGACAGCGGCAGAGACTAT-3’; revers e 5’-ATCTCCTCCTGCCACTTCTT-3’), and rat Apo A-I (forward 5’-AAGG ACAGCGGCAGAGACTA-3’; reverse 5’-CCACAACCTTTAGATGCCTT-3’). The sizes and sequences of these cDNA probes

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51 were verified by DNA sequencing prior to their use in Northern blot analysis. Filters were sequentially washed in 2X SSC (1X= 0.15 M sodium chloride, 0.015 M sodium citrate)-0.1% SDS and in 0.1x SSC-0.1% SDS twice each at 50C and then exposed to X-ray film to detect radiolabeled bands. Equal loading of total RNA for each experimental sample was verified by comparison to 18S rRNA ethidium bromide staining. Lipid Extraction Total lipids were extracted from conditioned media as described by Bligh and Dyer (1959), with modifications. For each sample, 2 mL of conditioned media was aliquotted into a 20 mL glass screw-top vial. Fourteen mL of chloroform:methanol (2:1, v/v) was then added and the vials were vortexed for 5 minutes. The vials were then centrifuged at 1700 rpm for 5 minutes. The bottom lipid-contai ning chloroform layer was transferred to a clean, dry, pre-weighed vial, placed in a 37oC water bath, and dried under nitrogen gas. Dry samples were placed in a 50oC oven for 10 minutes and placed in a desiccator to cool to room temperature. Samples were we ighed, and lipid weight was determined by difference. The sample was resuspended in chloroform and stored at -20oC until HDL cholesterol analysis. HDL Cholesterol Assay Lipid extracts from conditioned media were analyzed using a commercially available L-Type HDL-C kit, following the ma nufacturer’s directions. Briefly, using a 96-well plate, 3 L of sample was pipetted in to each well. Two hundred seventy L of Enzyme Color Solution (R1) was added, and the plate was incubated for 5 minutes at 37oC. Ninety L of Reacting Solution (R2) was then added, and the plate was incubated another 5 minutes at 37oC. The absorbance at 600 nm was measured using the

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52 SpectraMax 340 PC microplate reader (Mol ecular Devices, Sunnyvale, CA), and the concentration of the samples was calculated by plotting against a standard curve. Statistical Analysis All hybridization signals as measured by densitometry were evaluated by least squares analysis of variance (ANOVA) using the General Linear Model (GLM) procedure of the SAS software package (SAS Institute Inc, Cary, NC). In each experiment, treatments were run in duplic ate, and the whole experiment was also duplicated, giving n=4 plates per treatment The general model for mRNA analysis included experiment, treatment, and experi ment x treatment interaction. In mRNA analyses, densitometric values for target gene s were expressed as ratios of target gene densitometric values over the corresponding 18S rRNA densito metric values. For HDL cholesterol concentration, the sources of variation included experiment, treatment, experiment x treatment interac tion, and plate (experiment x treat ment). The plate, nested within experiment and treatment, was consid ered a random variable and therefore the plate variance was used as an error term to te st the effects of experiment, treatment, and experiment x treatment interaction. Treat ment means were further compared using preplanned orthogonal contrasts. These contrasts were contro l vs. fat treatment (ST, LA, LNA, EPA), saturated fat (ST) vs. PUFA (LA, LNA, EPA), n-6 (LA) vs. n-3 (LNA, EPA); and LNA vs. EPA. For all responses, th e two cell lines were analyzed separately. Results Effects of Fatty Acids on HepG2 Cells Steady-state levels of AC O mRNA were not affected by any FA treatment in HepG2 cells (P = 0.3; Figure 3-1). Concentr ations of HMG-R mRNA transcript were greater (+24%, P = 0.006) in HepG2 cells treat ed with ST than in PUFA-treated cells

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53 (Figure 3-2). Concentrations of Apo A-I mRNA transcript were greater (+15%, P = 0.05) in HepG2 cells treated with FA than in control cells (Figure 3-3). There were no differences in HDL cholesterol concentra tion among any of the treatments (P = 0.9; Figure 3-4). Effects of Fatty Acids on H-4-II-E Cells In the H-4-II-E cells, ACO mRNA expre ssion was greater (+26%, P = 0.004) in ST-treated cells as compared to PUFA-treat ed cells (Figure 3-5). Concentrations of HMG-R mRNA were greater in ST-t reated as compared to PUFA-treated cells (+27%; P = 0.002), in n-3 (EPA and LNA)-treated as co mpared to n-6 (LA)-treated cells (+30%; P = 0.004), and in EPA-treated as compared to LNA-treated cells (+49%; P < 0.001), with the EPA treatment showing the greates t induction of HMG-R mRNA transcript (Figure 3-6). Similarly, st eady-state levels of Apo A -I mRNA were increased in ST-treated cells as compared to PUFA -treated cells (+39%; P < 0.001) and in EPA-treated cells as compared to LNA-treat ed cells (+31%; P = 0.008; Figure 3-7). As compared to n-6 FA, n-3 FA increased (+79%; P = 0.0002) HDL cholesterol concentration by H-4-II-E cells, with the e ffect predominantly deriving from the large increase (+84%; P < 0.0001) in production wi th LNA as compared to EPA (Figure 3-8). Role of PPAR in Stearic Acid-Induced Effects on Gene Expression Co-incubation of HepG2 cells with ST and 10 M WY 14,643, a specific PPAR agonist, decreased (-9%; P = 0.04) ACO mRNA expression as compared to ST alone. There was no detectible effect on ACO mRNA with the use of the a gonist alone (P = 0.8; Figure 3-9). WY 14,643 decreased both basal (-32%; P = 0.0002) and ST-induced (-10%; P = 0.02) expression of HMG-R mR NA (Figure 3-10). Use of the PPAR agonist

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54 alone (P = 0.5) or in combination with ST (P = 0.4) had no e ffects on Apo A-I mRNA (Figure 3-11). In HepG2 cells, incubation with 10 M MK886, a specific PPAR inhibitor increased (P < 0.05) basal production of all three gene transcripts (Figures 3-12, 3-13, and 3-14). Co-incubation with MK886 had no effects on ST-induced expression of any of the genes. Co-incubation of H-4-II-E cells with WY 14,643 increa sed (+22%; P = 0.04) basal levels and enhanced (+38%; P = 0.0003) th e effect of ST on ACO gene expression (Figure 3-15). Both basal (-45%; P = 0.01) and ST-induced (-32%; P = 0.03) HMG-R mRNA expression were decrease d with the use of the PPAR agonist (Figure 3-16). The abundance of ST-induced Apo A-I mRNA transc ript was enhanced (+29%; P = 0.001) by the use of WY 14,643 (Figure 3-17). Basal le vels of Apo A-I mRNA were unaffected (P = 0.2). In H-4-II-E cells, incubation with MK886 attenuated (-28%; P = 0.001) the effects of ST on ACO mRNA expressi on (Figure 3-18). The PPAR inhibitor increased (+29%; P = 0.003) the basal concentration of HMG-R mRNA transcript, but had no effects (P = 0.8) on ST-induced gene expression (Figur e 3-19). The concentration of both basal (-96%; P = 0.01) and ST-induced (-39%; P = 0.03) Apo A-I mRNA transcript was reduced by the use of MK886 (Figure 3-20). Discussion Dietary fat has been implicated as a ma jor factor in many areas of health and disease. However, it has been suggested by numerous studies that all fats may not have the same effects. In this study, both human and rat hepatoma cells were used as models, as it also has been suggested that species di fferences exist in fat metabolism (Bergen and

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55 Mersmann, 2005). In HepG2 (human) liver cells, ACO mRNA expression was unaffected by any FA treatment. In contra st, in the H-4-II-E (rat) liver cells, ACO mRNA expression was induced by ST only. Other studies, however, have shown up-regulation of ACO mRNA in rat liver by di etary PUFAs as well as by saturated fats (Berthou et al., 1995). In HepG2 cells, it has been shown that PUFAs of differing saturation and length can regulate ACO mRNA in a dose-dependent and differential manner (Rise and Galli, 1999). In a human re tinoblastoma cell line, low concentrations of supplemental n-3 PUFA increased ACO mR NA, whereas high concentrations of the FA decreased it (Langelier et al., 2003). Consis tent with our findings in rat cells, pigs fed a tallow-based diet high in saturated fat had an increased concentration of ACO mRNA as compared to fish-oil fed animals (Ding et al., 2003). 3-hydroxy, 3-methylglutaryl CoA reducta se is the rate limiting enzyme in cholesterol synthesis, an d its inhibition is the target of th e statin class of drugs, used in the treatment of hyperlipidemias. In this study, we showed that in HepG2 cells, HMG-R mRNA was up-regulated by ST as compared to the PUFAs, whereas in the H-4-II-E cells, it was up-regulated by both ST and EPA. Consis tent with our findings in rodent cells, in C3H mice fed diets differing in fat com position, HMG-R mRNA wa s increased to a greater extent in mice fed the PUFA diet than in those fed the saturated fat diet (Cheema and Agellon, 1999). In Reuber H35 rat hepato ma cells, incubation wi th either saturated fats or PUFAs increased HMG-R enzyme activ ity (Garcia-Pelayo et al., 2003). Enzyme activity of HMG-R also has been shown to be increased in mice fed a diet high in PUFAs (Kuan and Dupont, 1989).

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56 Apolipoprotein A-I is the predominant lipop rotein associated with HDL cholesterol and is essential for its normal metabolism. Deletion of the Apo A-I gene in humans results in very low plasma concentrations of HDL cholesterol a nd premature coronary artery disease (Schaefer et al., 1982). Dietary fat has the ability to modulate plasma lipids, and may act, in part, by effects on apo lipoproteins. In this study, we showed that, in HepG2 cells, Apo A-I mRNA was up-regulated by all FA. However, no effects were seen in HDL cholesterol concen tration in the culture media. This is supported in a study by Dashti and coworkers (2002) in which HDL concentration was not different between LAand saturated fat-treated HepG2 cells. In Golden-Syrian hamsters, an effective model for human diet and blood lipid interact ions, canola and soybean oils increased Apo A-I mRNA as compared to a butter diet, though HDL concentrations were lowered in the diets containing unsaturat ed as compared to saturated fats (Dorfman et al., 2005). In the H-4-II-E cells, ST increase d Apo A-I mRNA concentra tion as compared to the PUFA-treated cells. In contrast to current fi ndings, Sprague-Dawley rats fed diets high in saturated fat or PUFAs showed no differences in Apo A-I amounts (Hat ahet et al., 2003). However, the saturated fat diet contained primarily palmitic acid, not stearic acid, as in this study. As fatty acids and their derivatives have been identified as potential ligands for peroxisome proliferator-activated receptors ( PPAR), we investigated the possibility that fatty acid effects in the two ce ll lines may be mediated by PPAR Incubation of HepG2 cells with WY 14,643, a PPAR agonist, had no effects on basal expression of ACO or Apo A-I mRNA. In the H-4-II-E cells, howev er, incubation with the agonist not only enhanced ST-induced ACO and Apo A-I mR NA expression but also increased basal

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57 expression of ACO mRNA. Not unexpectedly, use of the PPAR inhibitor, MK886, was able to cause the opposite effect, blocking the effects of ST on ACO and Apo A-I mRNA expression in H-4-II-E cells. Al though ACO is an established PPAR responsive gene (Tugwood, et al., 1992), species differences do exist. It is questionable whether the PPAR response element of human ACO is activ e (Woodyatt et al., 1999). Dietary studies have shown that rodents are re sponsive to the effects of PPAR activation, but non-rodent species, such as pr imates and guinea pigs, are resistant or unresponsive to some of the negative effect s (Bentley et al., 1993; Ca ttley et al., 1998). In a comprehensive analysis of gene expre ssion in human and rat hepatoma cells by microarray analysis, only rat ACO mR NA was responsive to WY 14,643 (Vanden Heuvel et al., 2003). Other genes that may be differentially regulat ed in human and rat liver include cytosolic aspartate aminotransfe rase (Tomkiewicz et al., 2004), peroxisomal 3-oxoacyl-CoA thiolase (Lawrence et al., 2001 ), and catalase (Amm erschlaeger et al., 2004). Additionally, different PPAR agonists may regulate lipid metabolism in a compound-dependent manner. A recent study by Duez and coworkers (2005) showed that, in mice, fenofibrate and gemfibrozil, both stimulate ACO mRNA expression, but only fenofibrate greatly induces Apo A-I gene expression. Interes tingly, although effects of PPAR activation or inhibition on ACO and A po A-I mRNA were different between the human and rat cell lines, effects on HMG-R mRNA were similar. In both cell lines, activation of PPAR by WY 14,643 caused a decrease in basal and ST-induced HMG-R mRNA expression. Inhibition of PPAR by MK886 increased HMG-R mRNA expression to a level similar to that induced by ST treatment alone, suggesting that ST

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58 effects are mediated by PPAR The findings of this study, in combination with other reports, strongly suggest a sp ecies-specific role for PPAR in gene regulation. Summary In HepG2 cells, ST up-regulated HMG-R gene expression as compared to PUFAs. As compared to control, in this cell line, a ll FA in this experime nt up-regulated Apo A-I gene expression. When PPAR was selectively activated, th e effect of ST on ACO gene expression was decreased, whereas both basa l and ST-induced HMG-R gene expression were decreased. Inc ubation with the PPAR inhibitor was able to decrease the basal production of all three genes, but had no effects on ST-induced gene expression. In H-4-II-E cells, ST up-regulated ACO, HMG-R, and Apo A-I gene expression as compared to the PUFAs. Selective activation of PPAR increased basal levels of ACO and further enhanced the effect of ST on ACO and Apo A-I mRNA. Conversely, selective activation of PPAR decreased basal levels of HMG-R and blocked the effect of ST on HMG-R mRNA. Incubation with the PPAR inhibitor was able to decrease the effects of ST-induced ACO and Apo A-I mRNA, as well as decrease the basal concentration of Apo A-I mRNA and increa se the basal concen tration of HMG-R mRNA. Together, these results indicate that FAs likely regulate li pid metabolizing genes in the liver through a PPAR -dependent mechanism. However, due to different responses in the human and rat hepatoma ce ll lines, the net effect s are likely species specific.

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59 Figure 3-1. Effect of long-chain FA on ACO mRNA expression in HepG2 cells. Ten micrograms of total cellular RNA isol ated from control and FA-treated HepG2 cells were subjected to Nort hern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal cont rasts. Contrast 1: Control vs. (ST + LA + LNA + EPA), P = 0.9; Contrast 2: ST vs. (LA + LNA + EPA), P = 0.09; Contrast 3: LA vs. (LNA + EPA), P = 0.7; Contrast 4: LNA vs. EPA, P = 0.2. Treatments ControlSTLALNAEPA ACO mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 3.2 kb 18S C LA ST LNA EPA ACO rRNA (A) (B)

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60 Figure 3-2. Effect of long-chain FA on HM G-R mRNA expression in HepG2 cells. Ten micrograms of total cellular RNA isol ated from control and FA-treated HepG2 cells were subjected to Nort hern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal cont rasts. Contrast 1: Control vs. (ST + LA + LNA + EPA), P = 0.8; Contrast 2: ST vs. (LA + LNA + EPA), P = 0.006; Contrast 3: LA vs. (LNA + EPA), P = 0.6; Contrast 4: LNA vs. EPA, P = 0.9. Treatments ControlSTLALNAEPA HMG-R mRNA (Normalized to 18S) 0.0 0.2 0.4 0.6 0.8 1.0 C LA ST LNA EPA 4.1-4.7kb HMG-R 18S rRNA (A) (B)

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61 Figure 3-3. Effects of long-chain FA on Apo A-I mRNA expression in HepG2 cells. Ten micrograms of total cellular RNA isol ated from control and FA-treated HepG2 cells were subjected to Nort hern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal cont rasts. Contrast 1: Control vs. (ST + LA + LNA + EPA), P = 0.05; Contrast 2: ST vs. (LA + LNA + EPA), P = 0.7; Contrast 3: LA vs. (LNA + EPA) P = 0.3; Contrast 4: LNA vs. EPA, P = 0.4. Treatments ControlSTLALNAEPA Apo A-I mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 0.5 0.6 C LA ST LNA EPA Apo A-I 18S 0.9 kb rRNA (A) (B)

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62 Figure 3-4. Effects of long-chain FA on HDL cholesterol production in HepG2 cells. Data represents least square means SEM calculated over two experiments. To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: Control vs. (S T + LA + LNA + EPA), P = 0.8; Contrast 2: ST vs. (LA + LNA + EPA), P = 0.4; Cont rast 3: LA vs. (LNA + EPA), P = 0.9; Contrast 4: LNA vs. EPA, P = 0.9. Treatment ControlSTLALNAEPA HDL Cholesterol (mg/dL) 0 1 2 3 4 5 Blah

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63 Figure 3-5. Effects of long-chain FA on ACO mRNA expression in H-4-II-E cells. Ten micrograms of total cellular RNA isol ated from control and FA-treated H-4-II-E cells were subjected to No rthern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal cont rasts. Contrast 1: Control vs. (ST + LA + LNA + EPA), P = 0.2; Contrast 2: ST vs. (LA + LNA + EPA), P = 0.004; Contrast 3: LA vs. (LNA + EPA), P = 0.6; Contrast 4: LNA vs. EPA, P = 0.07. Treatments ControlSTLALNAEPA ACO mRNA (Normalized to 18S) 0.00 0.02 0.04 0.06 0.08 0.10 0.12 0.14 C LA ST LNA EPA ACO 18S 3.2 kb rRNA (A) (B)

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64 Figure 3-6. Effects of long-chain FA on HMGR mRNA expression in H-4-II-E cells. Ten micrograms of total cellular RNA is olated from control and FA-treated H-4-II-E cells were subjected to No rthern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal cont rasts. Contrast 1: Control vs. (ST + LA + LNA + EPA), P = 0.9; Contrast 2: ST vs. (LA + LNA + EPA), P = 0.002; Contrast 3: LA vs. (LNA + EPA), P = 0.004; Contrast 4: LNA vs. EPA, P < 0.001. Treatments ControlSTLALNAEPA HMG-R mRNA (Normalized to 18S) 0.00 0.05 0.10 0.15 0.20 0.25 C LA ST LNA EPA HMG-R 18S 4.5 kb rRNA (A) (B)

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65 Figure 3-7. Effects of long-chain FA on Apo A-I mRNA expression in H-4-II-E cells. Ten micrograms of total cellular RNA is olated from control and FA-treated H-4-II-E cells were subjected to No rthern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal cont rasts. Contrast 1: Control vs. (ST + LA + LNA + EPA), P = 0.6; Contrast 2: ST vs. (LA + LNA + EPA), P < 0.001; Contrast 3: LA vs. (LNA + EPA), P = 0.6; Contrast 4: LNA vs. EPA, P = 0.008. Treatments ControlSTLALNAEPA Apo A-I mRNA (Normalized to 18S) 0.00 0.01 0.02 0.03 0.04 0.05 0.06 0.07 C LA ST LNA EPA 18S Apo A-I 0.9 kb rRNA(A) (B)

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66 Figure 3-8. Effects of long-chain FA on HDL cholesterol production in H-4-II-E cells. Data represents least square means SEM calculated over two experiments. To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: Control vs. (S T + LA + LNA + EPA), P = 0.3; Contrast 2: ST vs. (LA + LNA + EPA), P = 0.06; C ontrast 3: LA vs. (LNA + EPA), P = 0.0002; Contrast 4: LNA vs. EPA, P < 0.0001. Treatment ControlSTLALNAEPA HDL Cholesterol (mg/dL) 0 2 4 6 8 10 12 14 blah

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67 Figure 3-9. Effect of WY 14,643 on ACO mRNA response to ST in HepG2 cells. Ten micrograms of total cellular RNA isolat ed from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM proce dure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine tr eatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (ST + ST+A), P = 0.2; Contrast 2: ST vs. ST+A, P = 0.04; Contrast 3: Control vs. Agonist, P = 0.8. Treatments ControlSTAgonistST + A ACO mRNA (Normalized to 18S) 0.00 0.05 0.10 0.15 0.20 0.25 0.30 C ST PPAR Agon ST + Agon 18S ACO 3.2 kb rRNA (A) (B)

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68 Figure 3-10. Effect of WY 14,643 on HMG-R mRNA response to ST in HepG2 cells. Ten micrograms of total cellular RNA isolated from control and treated HepG2 cells were subjected to Nort hern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (ST + ST+A), P = 0.003; C ontrast 2: ST vs. ST+A, P = 0.02; Contrast 3: Control vs. Agonist, P = 0.002. Treatments ControlSTAgonistST + A HMG-R mRNA (Normalized to 18S) 0.00 0.05 0.10 0.15 0.20 0.25 0.30 C ST PPAR Agon ST + Agon HMG-R 18S 4.1-4.7kb rRNA (A) (B)

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69 Figure 3-11. Effect of WY 14,643 on Apo A-I mRNA response to ST in HepG2 cells. Ten micrograms of total cellular RNA isolated from control and treated HepG2 cells were subjected to Nort hern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (ST + ST+A), P = 0.6; C ontrast 2: ST vs. ST+A, P = 0.4; Contrast 3: Control vs. Agonist, P = 0.5. Treatments ControlSTAgonistST + A Apo A-I mRNA (Normalized to 18S) 0.00 0.05 0.10 0.15 0.20 0.25 C ST PPAR Agon ST + Agon Apo A-I 18S 0.9 kb rRNA (A) (B)

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70 Figure 3-12. Effect of MK886 on ACO mRNA re sponse to ST in HepG2 cells. Ten micrograms of total cellular RNA isolated from control and FA treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM proce dure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (ST + ST+I), P = 0.15; Contrast 2: ST vs. ST+I, P = 0.6; C ontrast 3: Control vs. Inhib, P = 0.03. Treatments ControlSTInhibST + I ACO mRNA (Normalized to 18S) 0.00 0.05 0.10 0.15 0.20 0.25 0.30 0.35 3.2 kb C ST PPAR Inhib ST + Inhib rRNA ACO 18S(A) (B)

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71 Figure 3-13. Effect of MK886 on HMG-R mRNA response to ST in HepG2 cells. Ten micrograms of total cellular RNA isolat ed from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM proce dure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (ST + ST+I), P = 0.5; Contrast 2: ST vs. ST+I, P = 0.4; Cont rast 3: Control vs. Inhib, P = 0.01. Treatments ControlSTInhibST + I HMG-R mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 0.5 C ST PPAR Inhib ST + Inhib HMG-R 18S 4.1-4.7kb rRNA (A) (B)

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72 Figure 3-14. Effect of MK886 on Apo A-I mRNA response to ST in HepG2 cells. Ten micrograms of total cellular RNA isolat ed from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM proce dure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (ST + ST+I), P = 0.01; Contrast 2: ST vs. ST+I, P = 0.6; C ontrast 3: Control vs. Inhib, P = 0.02. Treatments ControlSTInhibST + I Apo A-I mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 0.5 0.6 C ST PPAR Inhib ST + Inhib Apo A-I 18S 0.9 kb rRNA (A) (B)

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73 Figure 3-15. Effect of WY14,643 on ACO mRNA response to ST in H-4-II-E cells. Ten micrograms of total cellular RNA isolat ed from control and treated H-4-II-E cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM proce dure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (ST + ST+A), P = 0.007; Contrast 2: ST vs. ST+A, P = 0003; Contrast 3: Control vs. Agonist, P = 0.04. Treatments ControlSTAgonistST + A ACO mRNA (Normalized to 18S) 0.00 0.02 0.04 0.06 0.08 C ST PPAR Agon ST + Agon ACO 18S 3.2 kb rRNA (A) (B)

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74 Figure 3-16. Effect of WY 14,643 on HMG-R mRNA response to ST in H-4-II-E cells. Ten micrograms of total cellular RNA isolated from control and treated H-4-II-E cells were subjected to No rthern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (ST + ST+A), P = 0.12; Contrast 2: ST vs. ST+A, P = 0.03; Contrast 3: Contro l vs. Agonist, P = 0.01. Treatments ControlSTAgonistST + A HMG-R mRNA (Normalized to 18S) 0.00 0.02 0.04 0.06 0.08 0.10 C ST PPAR Agon ST + Agon HMG-R 18S 4.5 kb rRNA (A) (B)

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75 Figure 3-17. Effect of WY 14,643 on Apo A-I mRNA response to ST in H-4-II-E cells. Ten micrograms of total cellular RNA isolated from control and treated H-4-II-E cells were subjected to No rthern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control Agonist) vs. (ST + ST+A), P < 0.0001; Contrast 2: ST vs. ST+A, P = 0.001; Contrast 3: Contro l vs. Agonist, P = 0.2. Treatments ControlSTAgonistST + A Apo A-I mRNA (Normalized to 18S) 0.00 0.01 0.02 0.03 0.04 0.05 C ST PPAR Agon ST + Agon Apo A-I 18S 0.9 kb rRNA (A) (B)

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76 Figure 3-18. Effect of MK886 on ACO mRNA re sponse to ST in H-4-II-E cells. Ten micrograms of total cellular RNA isolat ed from control and treated H-4-II-E cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM proce dure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine tr eatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (ST + ST+I), P = 0.5; Contrast 2: ST vs. ST+I, P = 0.001; Contrast 3: Control vs. Inhib, P = 0.06. Treatments ControlSTInhibST + I ACO mRNA (Normalized to 18S) 0.00 0.05 0.10 0.15 0.20 0.25 0.30 C ST PPAR Inhib ST + Inhib ACO 18S 3.2 kb rRNA (A) (B)

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77 Figure 3-19. Effect of MK886 on HMG-R mRNA response to ST in H-4-II-E cells. Ten micrograms of total cellular RNA isolat ed from control and treated H-4-II-E cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM proce dure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (ST + ST+I), P = 0.004; Contrast 2: ST vs. ST+I, P = 0.8; Contrast 3: Control vs. Inhib, P = 0.003. Treatments ControlSTInhibST + I HMG-R mRNA (Normalized to 18S) 0.00 0.02 0.04 0.06 0.08 0.10 0.12 C ST PPAR Inhib ST + Inhib HMG-R 18s 4.5 kb rRNA (A) (B)

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78 Figure 3-20. Effect of MK886 on Apo A-I mRNA response to ST in H-4-II-E cells. Ten micrograms of total cellular RNA isolat ed from control and treated H-4-II-E cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM proce dure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (ST + ST+I), P = 0.003; Contrast 2: ST vs. ST+I, P = 0.03; Contrast 3: Control vs. Inhib, P = 0.01. Treatments ControlSTInhibST + I Apo A-I mRNA (Normalized to 18S) 0.00 0.01 0.02 0.03 0.04 0.05 C S PPAR Inhib ST + Inhib 18s 0.9 kb rRNA Apo A-I (A) (B)

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79 Figure 3-21. Regulation of lipid metabolizi ng genes and HDL cholesterol production by long-chain fatty acids. In HepG2 cells, HMG-R mRNA was up-regulated by ST as compared to the PUFAs. All fatty acids up-regulated Apo A-I mRNA as compared to control. Activation of PPAR attenuated the effects of ST on ACO and HMG-R gene expression. In H4-II-E cells, ST up-regulated ACO, HMG-R, and Apo A-I gene expression as compared to the PUFAs. Activation of PPAR increased basal expression of ACO and enhanced ST effects on ACO and Apo A-I mRNA. Both basal and ST-induced HMG-R mRNA levels were decreased by PPAR activation. Inhibition of PPAR decreased basal expression of Apo A-I and attenuated ST-induced expression of ACO and Apo A-I mRNA. Basal concentrations of HMG-R mRNA were increased by PPAR inhibition. As compared to n6 PUFA, n-3 PUFA increased HDL cholesterol production, with the effect predominantly deriving from the increase due to LNA. ACO mRNA HMG-R mRNA Apo A-I mRNA ACO mRNA HMG-R mRNA Apo A-I mRNA HDL-C PPAR ST LA LNA EPA Activation Inhibition HepG2 H-4-II-E ?

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80 CHAPTER 4 EFFECTS OF ISOMERS OF CONJUGAT ED LINOLEIC ACID ON LIPID METABOLIZING GENES AND HIGH-DENS ITY LIPOPROTEIN CHOLESTEROL PRODUCTION IN CULTURED HU MAN AND RAT HEPATOCYTES Introduction Conjugated linoleic acid (C LA) is a collective term for positional and geometric isomers of linoleic acid (LA). Though over 16 individual isomers ha ve been identified (Rickert et al 1999), only cis -9, trans -11 CLA and trans -10, cis -12 CLA are known to possess biological activity (Pariza et al., 2000). Cis -9, trans -11 CLA is the predominant CLA produced as an intermediate in the ru men during the biohydrogenation of dietary LA and is commonly found in dairy products and ruminant meat. Dietary sources of trans -10, cis -12 CLA derive predominantly from synthetic partial biohydrogenation and is found in margarines, shortenings, and s upplements (Gaullier et al., 2002). First identified in grilled beef as a potential anti-carcinogen (Pariza and Hargraves, 1985), numerous health benefits have been attributed to CLA mixt ures, including actions as an antiadipogenic (Park et al, 1997), antidiabe togenic (Houseknecht et al., 1998), and antiatherosclerotic (Kritchevsky et al., 2004) agent. More rece ntly, studies involving individual isomers have shown that the two ma in isoforms can have different effects on metabolism and cell function and may act throu gh different signaling pathways (Wahle et al., 2004). The metabolic responses to cis -9, trans -11 and trans -10, cis -12 CLA may differ, but both isomers have implications fo r human health. Most studies have been performed in animal models, with species diffe rences observed. In particular, only some

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81 of the findings attributed to animal models pertain to human subjects, and even when comparing studies in humans, results are often inconclusive (Terpstra, 2004). The objective of this study was to examine the short term effects of the two biologically active isomers of CLA on lipid metabolizing gene expression and high-density lipoprotein (HDL) cholesterol production in HepG2 (human) and H-4-II-E (rat) hepatoma cell lines. Based on both dietary and in vitro studies of lipid metabolism, we hypothesized that the different isomers of CLA may have diffe ring effects on acylCoA oxidase (ACO), 3-hydroxy, 3-methylgl utaryl CoA reductase (HMG-R), and apolipoprotein A-I (Apo A-I) gene expression. Also, because several fatty acids and their derivatives are known ligands for peroxisome proliferator-activated receptors (PPAR), we hypothesized that CLA isomers may act on lipid-metabo lizing genes through activation of PPAR in the liver. Materials and Methods Materials Polystyrene tissue culture dishes (100 x 20 mm) were purchased from Corning (Corning Glass Works, Corning, NY). The antibiotic/antimycotic (ABAM), sodium pyruvate, fatty acid-free bovi ne serum albumin (BSA), WY 14,643, and MK886 were from Sigma Chemical Co. (St. Louis, MO). Minimum Essential Medium (MEM), phenol red-free MEM, Hanks Balanced Salt Solution (HBSS) and TriZol reagent were from GIBCO BRL (Carlsbad, CA). The feta l bovine serum (FBS) was from Atlanta Biologicals (Norcross, GA). Linoleic acid, cis -9, trans -11 CLA, and trans -10, cis -12 CLA were from Cayman Chemicals (Ann Ar bor, MI). BioTrans nylon membrane and [ -32P]deoxycytidine triphosphate (SA 3000 Ci/n mol) were from MP Biolomedicals

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82 (Atlanta, GA). The Enzyme Color Solution, Reacting Solution, and HDL Calibrator were from Wako Diagnostics (Richmond, VA). Cell Culture and Treatment HepG2 (ATCC # HB-8065; Manassas, VA) and H-4-II-E (ATCC # CRL-1548; Manassas, VA) cells were cultured and fatty acids were complexed as described in chapter 3. To investigate the effects of supplemental CLA on hepatic gene expression and cholesterol synthesis, HepG2 and H4-II-E cells were treated with LA, cis -9, trans -11 CLA, or trans -10, cis -12 CLA (100 M). Sub-confluent cells were incubated with serum-free medium alone (Control) or with appropriate treat ments (listed above) complexed with BSA, for a period of 24 h. Cells were then rinsed twice with 10 mL HBSS. The remaining cell monolayer was then lysed in 3 mL TriZol reagent, and stored at -80C for subsequent mRNA analysis. The same fatty acid (FA) treatments were repeated, using phenol red-free MEM. After incubation, conditioned media were collected and stored at -20oC until lipid extraction and HDL cholesterol analysis. To investigate whether CLA effects on gene expression involves PPAR activation, confluent HepG2 and H-4 -II-E cells were treated with trans -10, cis -12 CLA isomer (100 M), the PPAR agonist WY 14,643 (10 M), or a combination of trans -10, cis -12 CLA and WY 14,643. Additional sets of culture dishes we re incubated with trans -10, cis -12 CLA alone, the PPAR antagonist MK886 (10 M; Kehrer et al., 2001), or a combination of trans -10, cis -12 CLA and MK886. After 24 h of incubation, cells were washed twice with 10 mL HBSS, ly sed with TriZol, and stored at -80oC until mRNA analysis.

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83 RNA Isolation and Analysis Total cellular RNA was isolated from cells using TriZol reagent according to the manufacturer’s instru ctions. Ten micrograms of tota l RNA was fractioned in a 1.0% agarose formaldehyde gel following previous ly described protocols (Ing et al., 1996) using the MOPS buffer (Fisher Scientific, Pitts burgh, PA) and transferred to a Biotrans nylon membrane by downward capillary transfer in 20X SSC (3 M NaCl, 0.3 M sodium citrate, pH 7.0) using the TurboBlotting sy stem (Schleicher and Schuel, Keene, NH). Nylon membranes were cross-linked by expos ure to a UV light source for 90 sec and baked at 80C for 1 h. Membranes were in cubated for 2 h at 50C in ultrasensitive hybridization buffer (ULTRAhyb; Ambion, Au stin, TX) followed by an overnight incubation at 50C in the same ULTRAhyb solution containing the 32P-labeled ACO, HMG-R, and Apo A-I cDNA probes. Probe s were generated by RT-PCR for ACO (forward 5’-CCGGAGCTGCTTACACACAT-3 ’; reverse 5’-GGTCATACGTGGCTGT GGTT-3’), HMG-R (forward 5’-TCCTTG GTGATGGGAGCTTGTTGTG-3’; reverse 5’-TGCGAACCCTTCAGATGTTTCGAGC-3’), human Apo A-I (forward 5’-AAGACA GCGGCAGAGACTAT-3’; reverse 5’-ATC TCCTCCTGCCACTTCTT-3’), and rat Apo A-I (forward 5’-AAGGACAGCGGCAGAGAC TA-3’; reverse 5’-CCACAACCTTTAG ATGCCTT-3’). The sizes and sequences of these cDNA probes were verified by DNA sequencing prior to their use in Northern blot analysis. Filters were sequentially washed in 2X SSC (1X= 0.15 M sodium chloride, 0.015 M sodium citrate)-0.1% SDS and in 0.1x SSC-0.1% SDS two times each at 50C and then exposed to X-ray film to detect radiolabeled bands. Equal loading of to tal RNA for each experimental sample was verified by comparison to 18S rR NA ethidium bromide staining.

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84 Lipid Extraction Total lipids were extracted from conditioned media as described by Bligh and Dyer (1959), with modifications. For each sample, 2 mL of conditioned media was aliquotted into a 20 mL glass screw-top vial. Fourteen mL of chloroform:methanol (2:1, v/v) was then added and the vials were vortexed for 5 minutes. The vials were then centrifuged at 1700 rpm for 5 min. The bottom lipid-containing chloroform layer was transferred to a clean, dry, pre-weighed vial, placed in a 37oC water bath, and dried under nitrogen gas. Dry samples were placed in a 50oC oven for 10 minutes and placed in a desiccator to cool to room temperature. Samples were we ighed, and lipid weight was determined by difference. The sample was resuspended in chloroform and stored at -20oC until HDL cholesterol analysis. HDL Cholesterol Assay Lipid extracts from conditioned media were analyzed using a commercially available L-Type HDL-C kit, following the ma nufacturer’s directions. Briefly, using a 96-well plate, 3 L of sample was pipetted in to each well. Two hundred seventy L of Enzyme Color Solution (R1) was added, and the plate was incubated for 5 minutes at 37oC. Ninety L of Reacting Solution (R2) was then added, and the plate was incubated another 5 minutes at 37oC. The absorbance at 600 nm was measured using the SpectraMax 340 PC microplate reader (Mol ecular Devices, Sunnyvale, CA), and the concentration of the samples was calculated by plotting against a standard curve. Statistical Analysis All hybridization signals as measured by densitometry were evaluated by least squares analysis of variance (ANOVA) using the General Linear Model (GLM) procedure of the SAS software package (SAS Institute Inc, Cary, NC). In each

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85 experiment, treatments were run in duplic ate, and the whole experiment was also duplicated, giving n=4 plates per treatment The general model for mRNA analysis included experiment, treatment, and experi ment x treatment interaction. In mRNA analyses, densitometric values for target gene s were expressed as ratios of target gene densitometric values over the corresponding 18S rRNA densito metric values. For HDL cholesterol concentration, the sources of variation included experiment, treatment, experiment x treatment interac tion, and plate (experiment x treat ment). The plate, nested within experiment and treatment, was consid ered a random variable and therefore the plate variance was used as an error term to te st the effects of experiment, treatment, and experiment x treatment interaction. Treat ment means were further compared using preplanned orthogonal contrasts. These contrasts were control vs. fat treatment (LA, cis 9, trans -11 CLA, trans -10, cis -12 CLA), LA vs. CLA ( cis -9, trans -11 CLA, trans -10, cis -12 CLA), and cis -9, trans -11 CLA vs. trans -10, cis -12 CLA. For all responses, the two cell lines were analyzed separately. Results Effects of Conjugated Linoleic Acid on HepG2 Cells Concentrations of ACO mRNA transcript were greater (+17%; P = 0.03) in HepG2 cells treated with CLA as compared to LA In addition, ACO mRNA concentration was increased (+22%; P = 0.009) in HepG2 cells treated with trans -10, cis -12 CLA as compared to the cis -9, trans -11 isomer (Figure 4-1). St eady-state levels of HMG-R mRNA were increased (+38%; P = 0.0003) in cells treated with CLA as compared to those treated with LA. Concentration of HMGR mRNA was also increa sed (+22%; P = 0.009) in cells treated with trans -10, cis -12 as compared to cis -9, trans -11 CLA (Figure 4-2). On average, the CLA isomer s increased (+21%; P = 0.03) Apo A-I mRNA

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86 transcript as compared to LA. Similar to the other genes studied, trans -10, cis -12 CLA increased (+22%; P = 0.03) Apo A-I ge ne transcript as compared to cis -9, trans -11 CLA (Figure 4-3). As compared to LA, incuba tion with CLA decreased (-18%; P = 0.05) HDL cholesterol production in HepG2 cells. This effect was predominantly derived from cis -9, trans -11 CLA, which decreased (-29%; P = 0.02) HDL cholesterol concentration in the media as compared to trans -10, cis -12 CLA (Figure 4-4). Effects of Conjugated Linole ic Acid on H-4-II-E Cells In H-4-II-E cells, ACO mRNA transcript was increased (+23%; P = 0.0005) by all FA as compared to control, though among th e FA studied, there were no differences (Figure 4-5). As compared to LA, CL A increased (+38%; P = 0.001) HMG-R gene expression; however, there were no differences between the two isom ers of CLA (P = 0.2; Figure 4-6). On aver age, treating H-4-II-E cells with FA decreased (-30%; P = 0.01) Apo A-I mRNA concentration, but there were no differences among the FA (Figure 4-7). There was no effect (P = 0.6) of any of the FA on HDL cholesterol production (Figure 4-8). Role of PPAR in trans -10, cis -12 CLA-Induced Effects on Gene Expression Incubation of HepG2 cells with 10 m WY 14,643, a specific PPAR agonist, had no effect on either basal (P = 0.99) or trans -10, cis -12 CLA-induced (P = 0.3) ACO mRNA expression (Figure 4-9). Basal leve ls of HMG-R mRNA were increased (+23%; P = 0.02) by use of the PPAR agonist, but there was no effect (P = 0.4) on trans -10, cis -12 CLA-induced gene expression (Figure 4-10). WY 14,643 incr eased (+38%; P < 0.0001) Apo A-I mRNA basal concentrations to levels similar to that induced by trans -10, cis -12 CLA, but had no additive effect (P = 0.9; Figure 4-11).

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87 In HepG2 cells, incubation with 10 m MK886, a specific PPAR inhibitor, had no effect on either basal or trans -10, cis -12 CLA-induced expression of any of the genes studied (ACO, Figure 4-12; HMG-R, Fi gure 4-13, Apo A-I, Figure 4-14). Incubation of H-4-II-E cells with WY 14,643 increased (+24%; P = 0.02) basal expression and attenuated (27%; P = 0.02) the effects of trans -10, cis -12 CLA on ACO gene expression (Figure 4-15). There were no effects of the agonist on basal (P = 0.9) or trans -10, cis -12 CLA-induced (P = 0.2) expression of HMG-R or Apo A-I mRNA concentrations (HMG-R, Figure 4-16; Apo A-I, Figure 4-17). In H-4-II-E cells, incubation with MK 886 had no effects (P = 0.3) on ACO gene expression (Figure 4-18). The PPAR inhibitor decreased (-25%; P = 0.02) basal expression of HMG-R mRNA, but had no effects (P = 0.1) on trans -10, cis -12 CLA-induced gene expression (Figure 4-19). Basal levels were unaffected (P = 0.9), but co-incubation with MK886 a ttenuated (-35%; P = 0.002) trans -10, cis -12 CLA-induced Apo A-I mRNA concentration (Figure 4-20). Discussion Numerous beneficial physio logical effects have been attributed to CLA, though these effects may be both isomer and species specific. One of the potential mechanisms by which CLA modulates health and diseas e states is through changes in lipid metabolism. To address these facts, in this study, the two biologically active isomers of CLA were studied in both human and rat he patoma cell lines. In HepG2 (human) cells, ACO mRNA expression was up-regulated by CLA as compared to LA treated cells, with an increase additionally s een with incubation of trans -10, cis -12 CLA as compared with cis -9, trans -11 CLA. In contrast, all FA increas ed ACO mRNA as compared to control in H-4-II-E cells, but there were no differences among the FA studied. Several animal

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88 and cell models have also show n similar effects. Feeding mi ce a mix of (Peters et al., 2001) or individual CLA isomers (Warren et al., 2003; Degrace et al., 2004) increases expression of ACO mRNA as compared to c ontrol mice. In two studies, the increases seen in ACO mRNA levels in CLA-fed mice al so coincided with increases in enzyme activity (Takahashi et al., 2003; Ide, 2005). In FaO cells, a rat hepatoma cell line derived from H4IIEC3 cells (Bayly et al., 1993), AC O gene expression was increased with 200 M cis -9, trans -11 CLA. This effect was not see n, however, with lower concentrations of CLA (Moya-Camarena et al., 1999). In a recent study using a hamster model, ACO activity was increased by trans -10, cis -12 CLA as compared to control or cis -9, trans -11 CLA (Macarulla et al., 2005). Together with our findings, these studies suggest a potential role for CLA isomers in increasing liver peroxisomal -oxidation. Fatty acids have the ability to modulate serum cholesterol levels, though the exact site and mode of regulation may vary from one model to another. One potential gene involved is HMG-R, the rate limiting enzyme in cholesterol synthesis. In HepG2 cells, CLA isomers increased HMG-R mRNA transcript as compared to the parent molecule, LA. Between the CLA isomers, trans -10, cis -12 CLA up-regulated HMG-R mRNA concentrations as compared with the cis -9, trans -11 isomer. This differed in the H-4-II-E cells, where, on average, CLA increased ge ne expression as compared to LA, but no differences were seen between the two CLA isomers. Although the effects of saturated and polyunsaturated fats on HM G-R gene expression and en zyme activity have been examined, few studies have explored the role of CLA. In a recent study, HMG-R activity was decreased in rats fed diacylglycerol-en riched structured lip ids containing CLA as compared to those fed lipids without CLA or corn oil (Kim et al., 2006). Though this

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89 differs from our findings, gene expression was not measured, and as with other target genes, it would not be surprising if species and model-specific diffe rences exist relative to transcriptional and/or posttranscripti onal regulation of the HMG-R enzyme. Another factor involved in normal lipoprot ein profiles and metabolism is Apo A-I, the predominant apolipoprotein associated w ith HDL cholesterol. Dietary fat has the ability to modulate plasma lipids, and may act in part, by effects on apolipoproteins. In general, CLA-induced changes in the blood li pid profile observed in various models are conflicting. In our study, Apo A-I gene tran script was increased by the CLA isomers as compared to LA, and by trans -10, cis -12 CLA as compared to cis -9, trans -11 CLA in HepG2 cells. High-density lipoprotein (HDL) cholesterol production in HepG2 cells was decreased by the CLA isomers as compared to LA, with the effect predominantly derived from the decrease due to cis -9, trans -11 CLA. In H-4-II-E cel ls, steady-state levels of Apo A-I mRNA was decreased by all FA treatme nts; however, there were no differences among the FA studied. These decreases in ge ne expression did not result in increased HDL cholesterol production, as levels were not different among any treatments. The different responses seen in the two cell lines is reflected by conflicting responses in other species studied. Mice supplemented with cis -9, trans -11 or trans -10, cis -12 CLA showed no differences in Apo A-I mRNA concentrations in control or CLA-fed animals (Warren et al., 2003). In contrast, in apo-E deficient mice, dietary trans -10, cis -12 CLA decreased plasma Apo A-I levels as compared to cis -9, trans -11 CLA (Arbones-Mainar et al., 2006). Similar to the decrease in HDL choles terol production in HepG2 cells, rabbits fed a mixture of CLA isomers showed an increase in total serum cholesterol and a decrease in HDL cholesterol (Kritchevsky et al., 2000) Additionally, in the Syrian Golden

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90 hamster, diets containing trans -10, cis -12 CLA increased HDL cholesterol as compared with LA or cis -9, trans -11 CLA diets (Mitchell et al., 2005 ). However, some studies in rat models have shown no effect of dietary CLA on serum HDL cholesterol (Kloss et al., 2005). In a human dietary study, both Apo A-I ge ne transcript and HDL cholesterol were decreased by CLA-enriched butter as compared with pre-supplement levels (Desroches et al., 2005). Fatty acids and their derivatives have been identified as potential ligands for PPAR. As several CLA isomers have been identified as high-affinity liga nds and activators of PPAR (Moya-Camarena et al., 1999), we investigat ed the possibility that CLA effects in the two cell lines may be mediated by PPAR Incubation of HepG2 cells with WY 14,643, a specific PPAR agonist, showed no effects on trans -10, cis -12 CLA-induced gene expression, although it di d increase basal expression of HMG-R and Apo A-I gene transcript s. Inhibition of PPAR had no effects on any of the genes in HepG2 cells. In H-4-II-E cells, however, the PPAR response differed. Activation of PPAR had no effects on HMG-R or Apo A -I mRNA concentrations, but basal concentrations of ACO were increased. In contrast, MK886 decreased basal levels of HMG-R mRNA expression and at tenuated the effect of trans -10, cis -12 CLA-induced Apo A-I concentration. Although ACO is an established PPAR responsive gene (Tugwood, et al., 1992), species differences do exist. It is questionable whether the PPAR response element of human ACO is activ e (Woodyatt et al., 1999). Dietary studies have shown that rodents are re sponsive to the effects of PPAR activation, but non-rodent species, such as pr imates and guinea pigs, are resistant or unresponsive to some of the negative effect s (Bentley et al., 1993; Ca ttley et al., 1998). In a

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91 comprehensive analysis of gene expre ssion in human and rat hepatoma cells by microarray analysis, only rat ACO mR NA was responsive to WY 14,643 (Vanden Heuvel et al., 2003). Other genes that may be differentially regulat ed in human and rat liver include cytosolic aspartate aminotransfe rase (Tomkiewicz et al., 2004), peroxisomal 3-oxoacyl-CoA thiolase (Lawrence et al., 2001 ), and catalase (Amm erschlaeger et al., 2004). Consistent with our findi ngs in the rat cell line, A po A-I gene expression was not different from controls in mi ce fed fenofibrate, a potent PPAR activator (Warren et al., 2003). However, different PPAR agonists may regulate lipid metabolism in a compound-dependent manner. A recent study by Duez and coworkers (2005) showed that, in mice, fenofibrate and gemfibroz il, both stimulated ACO mRNA expression, but only fenofibrate greatly induced Apo A-I gene expression. The lack of effect of PPAR activation or inhibition on trans -10, cis -12 CLA-induced gene expression in our study may be due to possible interactions between CLA and PPAR which may serve as a PPAR -independent mediator in response to CLA supplementation, as shown in PPAR -null mice (Peters et al., 2001). Warren a nd coworkers also supported this idea, with reports that PPAR expression in mice decreased with trans -10, cis -12 CLA, while ACO mRNA expression increased (Warren et al., 2003). Findings from our studies and others suggest that it is pr obable that the effects of CLA are not solely dependent upon PPAR Summary In HepG2 cells, ACO, HMG-R, and Apo A-I steady-state mRNA levels were up-regulated by both CLA isomers as compared to LA, and the greatest gene induction was seen with the trans -10, cis -12 CLA isomer. Selective activation or inhibition of PPAR had no effect on trans -10, cis -12 CLA-induced gene expression. However,

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92 incubation of HepG2 cells with the PPAR agonist increased basal production of HMG-R and Apo A-I mRNA concentration. Cons istent with the low level of endogenous expression of PPARs in HepG2 cells (Hsu et al., 2001), both the PPAR activator and PPAR inhibitor had marginal effects on basal and trans -10, cis -12 CLA-stimulated lipid metabolizing gene expression in the human hepatoma cell line. In H-4-II-E cells, all of the FA studied increased expressi on of ACO mRNA and decreased expression of Apo A-I mRNA. On average, CLA isomers increased HMG-R gene expression as compared with LA, alt hough there was no difference between the two isomers. Selective activation of PPAR increased basal expression and attenuated trans -10, cis -12 CLA-induced expression of ACO mRNA concentration. Activation of PPAR had no effect on HMG-R or Apo A-I ge ne transcripts. Inhibition of PPAR decreased basal expression of HMGR gene transcript and attenuated trans -10, cis -12 CLA effects on Apo A-I mRNA. Taken t ogether, these results indicate that trans -10, cis -12 CLA likely regulates lipid metabo lizing genes in th e liver through a PPAR -dependent mechanism. However, due to different responses in the human and rat hepatoma cell lines, the net effects are likely species specific.

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93 Figure 4-1. Effect of CLA on ACO mRNA expression in He pG2 cells. Ten micrograms of total cellular RNA isolated from co ntrol and FA-treated HepG2 cells were subjected to Northern blot analysis, a nd resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two expe riments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: Control vs. (LA + cis -9, trans -11 CLA + trans -10, cis -12 CLA), P = 0.09; Contrast 2: LA vs. ( cis -9, trans -11 CLA + trans -10, cis -12 CLA), P = 0.03; Contrast 3: cis -9, trans -11 CLA vs. trans -10, cis -12 CLA, P = 0.009. Treatment ControlLACLA-c9,t11CLA-t10,c12 ACO mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 0.5 C LA CLA c 9, t 11 CLA t 10, c 12 18S 3.2kb (A) (B) ACO rRNA

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94 Figure 4-2. Effect of CLA on HMG-R mR NA expression in HepG2 cells. Ten micrograms of total cellular RNA isol ated from control and FA-treated HepG2 cells were subjected to Nort hern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal cont rasts. Contrast 1: Control vs. (LA + cis -9, trans -11 CLA + trans -10, cis -12 CLA), P = 0.5; Contrast 2: LA vs. cis -9, trans -11 CLA + trans -10, cis -12 CLA), P = 0.0003; Contrast 3: cis -9, trans -11 CLA vs. trans -10, cis -12 CLA, P = 0.009. Treatment ControlLACLA-c9,t11CLA-t10,c12 HMG-R mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 0.5 C LA CLA c 9, t 11 CLA t 10, c 12 4.1-4.7kb 18S (A) (B) HMG-R rRNA

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95 Figure 4-3. Effect of CLA on Apo A-I mRNA expression in HepG2 cells. Ten micrograms of total cellular RNA isol ated from control and FA-treated HepG2 cells were subjected to Nort hern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal cont rasts. Contrast 1: Control vs. (LA + cis -9, trans -11 CLA + trans -10, cis -12 CLA), P = 0.9; Contrast 2: LA vs. cis -9, trans -11 CLA + trans -10, cis -12 CLA), P = 0.03; Contrast 3: cis -9, trans -11 CLA vs. trans -10, cis -12 CLA, P = 0.03. Treatment ControlLACLA-c9,t11CLA-t10,c12 Apo A-I mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 0.5 C LA CLA c 9, t 11 CLA t 10, c 12 18S 0.9kb (A) (B) Apo A-I rRNA

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96 Figure 4-4. Effect of CLA on HDL chol esterol production by HepG2 cells. Data represents least square means SEM cal culated over two experiments. To further examine treatment effects, m eans were separated using orthogonal contrasts. Contrast 1: Control vs. (LA + cis -9, trans -10 CLA + trans -10, cis -12 CLA), P = 0.5; Contrast 2: LA vs. ( cis -9, trans -10 CLA + trans -10, cis -12 CLA), P = 0.05; Contrast 3: cis -9, trans -10 CLA vs. trans -10, cis -12 CLA, P = 0.02. Treatment ControlLACLA c9,t11CLA t10,c12 HDL Cholesterol (mg/dL) 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 Blah

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97 Figure 4-5. Effect of CLA acid on ACO mRNA expression in H-4-II-E cells. Ten micrograms of total cellular RNA isol ated from control and FA-treated H-4-II-E cells were subjected to No rthern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal cont rasts. Contrast 1: Control vs. (LA + cis -9, trans -11 CLA + trans -10, cis -12 CLA), P = 0.0005; Contrast 2: LA vs. ( cis -9, trans -11 CLA + trans -10, cis -12 CLA), P = 0.3; Contrast 3: cis -9, trans -11 CLA vs. trans -10, cis -12 CLA, P = 0.07. Treatment ControlLACLA 9,11CLA 10,12 ACO mRNA (Normalized to 18S) 0.00 0.05 0.10 0.15 0.20 0.25 3.2kb 18S C LA CLA c 9, t 11 CLA t 10, c 12 ACO rRNA (A) (B)

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98 Figure 4-6. Effect of CLA on HMG-R mRNA expression in H-4-II-E cells. Ten micrograms of total cellular RNA isol ated from control and FA-treated H-4-II-E cells were subjected to No rthern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal cont rasts. Contrast 1: Control vs. (LA + cis -9, trans -11 CLA + trans -10, cis -12 CLA), P = 0.2; Contrast 2: LA vs. ( cis -9, trans -11 CLA + trans -10, cis -12 CLA), P = 0.001; Contrast 3: cis -9, trans -11 CLA vs. trans -10, cis -12 CLA, P = 0.2. Treatment ControlLACLA 9,11CLA 10,12 HMG-R mRNA (Normalized to 18S) 0.00 0.02 0.04 0.06 0.08 0.10 0.12 4.5kb 18S C LA CLA c 9, t 11 CLA t 10, c 12 HMG-R rRNA (A) (B)

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99 Figure 4-7. Effect of CLA on Apo A-I mRNA expression in H-4-II-E cells. Ten micrograms of total cellular RNA isol ated from control and FA-treated H-4-II-E cells were subjected to No rthern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal cont rasts. Contrast 1: Control vs. (LA + cis -9, trans -11 CLA + trans -10, cis -12 CLA), P = 0.01; Contrast 2: LA vs. ( cis -9, trans -11 CLA + trans -10, cis -12 CLA), P = 0.3; Contrast 3: cis -9, trans -11 CLA vs. trans -10, cis -12 CLA, P = 0.3. Treatment ControlLACLA-c9,t11CLA-t10,c12 Apo A-I mRNA (Normalized to 18S) 0.00 0.01 0.02 0.03 0.04 0.05 0.06 0.9kb 18S C LA CLA c 9, t 11 CLA t 10, c 12 Apo A-I rRNA (A) (B)

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100 Figure 4-8. Effect of CLA on HDL choles terol production by H-4-II-E cells. Data represents least square means SEM cal culated over two experiments. To further examine treatment effects, m eans were separated using orthogonal contrasts. Contrast 1: Control vs. (LA + cis -9, trans -10 CLA + trans -10, cis -12 CLA), P = 0.2; Contrast 2: LA vs. ( cis -9, trans -10 CLA + trans -10, cis -12 CLA), P = 0.1; Contrast 3: cis -9, trans -10 CLA vs. trans -10, cis -12 CLA, P = 0.1. Treatment ControlLACLA c9,t11CLA t10,c12 HDL Cholesterol (mg/dL) 0 2 4 6 8 10 12 14 16 18 c

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101 Figure 4-9. Effect of WY 14,643 on ACO mRNA response to trans -10, cis -12 CLA in HepG2 cells. Ten micrograms of total ce llular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (CLA + CLA+A), P = 0.01; Contrast 2: CLA vs. CLA+A, P = 0.99; Contrast 3: Control vs. Agonist, P = 0.3. Treatment ControlCLA 10,12AgonistCLA + Agonist ACO mRNA (Normalized to 18S) 0.00 0.05 0.10 0.15 0.20 0.25 0.30 18S 3.2kb C CLA t 10, c 12 PPAR Agonist CLA + Agonist ACO rRNA (A) (B)

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102 Figure 4-10. Effect of WY 14,643 on HMG-R mRNA response to trans -10, cis -12 CLA in HepG2 cells. Ten micrograms of tota l cellular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (CLA + CLA+A), P = 0.01; Contrast 2: CLA vs. CLA+A, P = 0.4; Contrast 3: Control vs. Agonist, P = 0.02. Treatment ControlCLA 10,12AgonistCLA + Agonist HMG-R mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 18S 4.1-4.7kb C CLA t 10, c 12 PPAR Agonist CLA + Agonist HMG-R rRNA (A) (B)

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103 Figure 4-11. Effect of WY 14,643 on Apo A-I mRNA response to trans -10, cis -12 CLA in HepG2 cells. Ten micrograms of tota l cellular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (CLA + CLA+A), P = 0.0006; Cont rast 2: CLA vs. CLA+A, P = 0.9; Contrast 3: Control vs. Agonist, P < 0.0001. Treatment ControlCLA 10,12AgonistCLA + Agonist Apo A-I mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 18S 0.9kb C CLA t 10, c 12 PPAR Agonist CLA + Agonist Apo A-I rRNA (A) (B)

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104 Figure 4-12. Effect of MK886 on ACO mRNA response to trans -10, cis -12 CLA in HepG2 cells. Ten micrograms of total ce llular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (CLA + CLA+I), P = 0.5; C ontrast 2: CLA vs CLA+I, P = 0.8; Contrast 3: Control vs. Agonist, P = 0.8. Treatment ControlCLA 10,12InhibCLA + Inhib ACO mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 18S 3.2kb C CLA t 10, c 12 PPAR Inhib CLA + Inhib (A) (B) ACO rRNA

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105 Figure 4-13. Effect of MK886 on HMG-R mRNA response to trans -10, cis -12 CLA in HepG2 cells. Ten micrograms of total ce llular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (CLA + CLA+I), P = 0.007; C ontrast 2: CLA vs. CLA+I, P = 0.8; Contrast 3: Contro l vs. Agonist, P = 0.8. Treatment ControlCLA 10,12InhibCLA + Inhib HMG-R mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 0.5 0.6 4.1-4.7kb 18S C CLA t 10, c 12 PPAR Inhib CLA + Inhib (A) (B) HMG-R rRNA

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106 Figure 4-14. Effect of MK886 on Apo A-I mRNA response to trans -10, cis -12 CLA in HepG2 cells. Ten micrograms of total ce llular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (CLA + CLA+I), P = 0.02; C ontrast 2: CLA vs CLA+I, P = 0.5; Contrast 3: Control vs. Agonist, P = 0.5. Treatment ControlCLA 10,12InhibCLA + Inhib Apo A-I mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 0.5 0.6 C CLA t 10, c 12 PPAR Inhib CLA + Inhib 0.9kb 18S Apo A-I rRNA (A) (B)

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107 Figure 4-15 Effect of WY 14,643 on ACO mRNA response to trans -10, cis -12 CLA in H-4-II-E cells. Ten micrograms of tota l cellular RNA isolated from control and treated H-4-II-E cells were subject ed to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (CLA + CLA+A), P = 0.07; Contrast 2: CLA vs. CLA+A, P = 0.02; Contrast 3: Control vs. Agonist, P = 0.02. Treatment ControlCLA 10,12AgonistCLA + Agonist ACO mRNA (Normalized to 18S) 0.00 0.02 0.04 0.06 0.08 0.10 0.12 0.14 18S 3.2kb C CLA t 10, c 12 PPAR Agonist CLA + Agonist ACO rRNA (A) (B)

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108 Figure 4-16. Effect of WY 14,643 on HMG-R mRNA response to trans -10, cis -12 CLA in H-4-II-E cells. Ten micrograms of to tal cellular RNA isolated from control and treated H-4-II-E cells were subject ed to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (CLA + CLA+A), P = 0.4; C ontrast 2: CLA vs. CLA+A, P = 0.2; Contrast 3: Contro l vs. Agonist, P = 0.9. Treatment ControlCLA 10,12AgonistCLA + Agonist HMG-R mRNA (Normalized to 18S) 0.00 0.02 0.04 0.06 18S 4.5kb C CLA t 10, c 12 PPAR Agonist CLA + Agonist HMG-R rRNA (A) (B)

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109 Figure 4-17. Effect of WY 14,643 on Apo A-I mRNA response to trans -10, cis -12 CLA in H-4-II-E cells. Ten micrograms of to tal cellular RNA isolated from control and treated H-4-II-E cells were subject ed to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (CLA + CLA+A), P = 0.1; C ontrast 2: CLA vs. CLA+A, P = 0.1; Contrast 3: Contro l vs. Agonist, P = 0.8. Treatment ControlCLA 10,12AgonistCLA + Agonist Apo A-I mRNA (Normalized to 18S) 0.00 0.02 0.04 0.06 0.08 18S 0.9kb C CLA t 10, c 12 PPAR Agonist CLA + Agonist Apo A-I rRNA (A) (B)

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110 Figure 4-18. Effect of MK886 on ACO mRNA response to trans -10, cis -12 CLA in H-4-II-E cells. Ten micrograms of tota l cellular RNA isolated from control and treated H-4-II-E cells were subject ed to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (CLA + CLA+I), P = 0.003; C ontrast 2: CLA vs. CLA+I, P = 0.3; Contrast 3: Control vs. Inhib, P = 0.3. Treatment ControlCLA 10,12InhibitorCLA + Inhib ACO mRNA (Normalized to 18S) 0.00 0.05 0.10 0.15 0.20 18S 3.2kb C CLA t 10, c 12 PPAR Inhib CLA + Inhib ACO rRNA (A) (B)

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111 Figure 4-19. Effect of MK886 on HMG-R mRNA response to trans -10, cis -12 CLA in H-4-II-E cells. Ten micrograms of tota l cellular RNA isolated from control and treated H-4-II-E cells were subject ed to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (CLA + CLA+I), P = 0.9; C ontrast 2: CLA vs CLA+I, P = 0.1; Contrast 3: Control vs. Inhib, P = 0.02. Treatment ControlCLA 10,12InhibitorCLA + Inhib HMG-R mRNA (Normalized to 18S) 0.00 0.02 0.04 0.06 0.08 0.10 0.12 0.14 18S 4.5kb C CLA t 10, c 12 PPAR Inhib CLA + Inhib HMG-R rRNA (A) (B)

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112 Figure 4-20. Effect of MK886 on Apo A-I mRNA response to trans -10, cis -12 CLA in H-4-II-E cells. Ten micrograms of tota l cellular RNA isolated from control and treated H-4-II-E cells were subject ed to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (CLA + CLA+I), P = 0.4; C ontrast 2: CLA vs CLA+I, P = 0.002; Contrast 3: Control vs. Inhib, P = 0.9. Treatment ControlCLA 10,12InhibitorCLA + Inhib Apo A-I mRNA (Normalized to 18S) 0.00 0.02 0.04 0.06 0.08 18S 0.9kb C CLA t 10, c 12 PPAR Inhib CLA + Inhib Apo A-I rRNA (A) (B)

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113 Figure 4-21. Regulation of lipid metabolizi ng genes and HDL cholesterol production by CLA. In HepG2 cells, ACO, HMG-R, and Apo A-I gene expression were up-regulated by CLA isomers as compared to LA, with the greatest induction seen with t 10, c 12 CLA. Activation of PPAR increased basal expression of HMG-R and Apo A-I mRNA. HDL chol esterol production was decreased by c 9, t 11 CLA. In H-4-II-E cells, all of th e fatty acids increased expression of ACO mRNA and decreased expression of Apo A-I mRNA. On average, CLA isomers increased HMG-R gene expression. Activation of PPAR increased basal expression of ACO mR NA. Inhibition of PPAR decreased basal expression of HMG-R mRNA and attenuated t 10, c 12 CLA-effects on Apo A-I gene expression. ACO mRNA HMG-R mRNA Apo A-I mRNA HDL-C ACO mRNA HMG-R mRNA Apo A-I mRNA HDL-C PPAR Activation Inhibition HepG2 H-4-II-E LA CLA CLA c 9, t 11 t 10, c 12

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114 CHAPTER 5 EFFECTS OF CIS AND TRANS ISOMERS OF OCTADECENOIC ACID ON LIPID METABOLIZING GENES AND HIGH-DENS ITY LIPOPROTEIN CHOLESTEROL PRODUCTION IN CULTURED HU MAN AND RAT HEPATOCYTES Introduction Trans -fatty acids are geometrical isomers of unsaturated fatty acids (FA) that assume a saturated fat-like configurati on that differs from that of their cis counterparts. The predominant source of trans fats in the human diet is hydrogenated oils, such as margarine and partially hydrogenated soybean oil, commonly found in baked goods and deep fat-fried fast foods (Hu et al., 2001). Metabolic studies in several species have shown that trans -FA can negatively alter the lipid pr ofile to a greater extent than saturated fats, because they not only incr ease the concentration of small, dense low-density lipoprotein (LDL) cholesterol (M auger et al., 2003), but also decrease high-density lipoprotein (HDL) c holesterol concentration in some studies (Judd et al., 1994; de Roos et al., 2003). Additionall y, epidemiological evidence has reported trans -FA intake to be associated with increase d risk for cardiovascular disease (Ascherio et al., 1999). Few studies, however, have examined the role that individual trans -FA may have in modulating lipid metabolism. As ha s been reported with other fatty acids, it is possible that cis and trans isomers of octadecenoic acid may also have differential effects on lipid metabolism. The objective of this study was to examine the short term effects of cis and trans isomers of octadecenoic acid on lipid metabo lizing gene expression and HDL cholesterol production in HepG2 (human) and H-4-II-E (rat) hepatoma cell lines. Based on both

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115 dietary and in vitro studies of lipid metabolism, we hypothesized that the different cis and trans isomers may have differing effects on acyl-CoA oxidase (ACO), 3-hydroxy, 3-methylglutaryl CoA reductase (HMG-R), and apolipoprotein A-I (Apo A-I) gene expression. Also, because several fatty acids and their derivatives are known ligands for peroxisome proliferator-activated receptor s (PPAR), we hypothesized that these fatty acids may act on lipid metabolizing genes through activation of PPAR in the liver. Materials and Methods Materials Polystyrene tissue culture dishes (100 x 20 mm) were purchased from Corning (Corning Glass Works, Corning, NY). The antibiotic/antimycotic (ABAM), sodium pyruvate, fatty acid-free bovine serum albumin (BSA), cis -vaccenic acid (c11), trans -vaccenic acid (t11), WY 14,643, and MK886 were from Sigma Chemical Co. (St. Louis, MO). Minimum Essential Medi um (MEM), phenol red-free MEM, Hanks Balanced Salt Solution (HBSS) and TriZol reagent were from GIBCO BRL (Carlsbad, CA). The fetal bovine serum (FBS) was from Atlanta Biologicals (Norcross, GA). Oleic (c9) and elaidic (t9) acids were from Cayman Chemicals (Ann Arbor, MI). BioTrans nylon membrane and [ -32P]deoxycytidine triphosphate (SA 3000 Ci/nmol) were from MP Biolomedicals (Atlanta, GA). The En zyme Color Solution, Reacting Solution, and HDL Calibrator were from Wa ko Diagnostics (Richmond, VA). Cell Culture and Treatment HepG2 (ATCC # HB-8065; Manassas, VA) and H-4-II-E (ATCC # CRL-1548; Manassas, VA) cells were cultured and fatty acids were complexed as described in chapter 3. To investigate the effects of supplemental octade cenoic fatty acids of differing bond position and orientation on hepatic gene expression and chol esterol synthesis,

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116 HepG2 and H-4-II-E cells were treated with oleic (c9; cis -9,18:1), elaidic (t9; trans -9, 18:1), cis -vaccenic (c11; cis -11, 18:1) or trans -vaccenic ( trans -11; 18:1) acids (100 M). Sub-confluent cells were incubated with se rum-free medium alone (Control) or with appropriate treatments (listed above) complexed with BSA, for a period of 24 h. Cells were then rinsed twice with 10 mL HBSS. The remaining cell monolayer was then lysed in 3 mL TriZol reagent, and stored at 80C for subsequent mRNA analysis. The same FA treatments were repeated, using phenol red-free MEM. After incubation, conditioned media were collected and stored at -20oC until lipid extraction and HDL cholesterol analysis. To investigate whether FA effect s on gene expression involves PPAR activation, confluent HepG2 and H-4-II-E cells were treated with the appropriate FA (100 M), the PPAR agonist WY 14,643 (10 M), or a combination of FA and WY 14,643. In the HepG2 cells, c11 was used, as this FA lead to the greatest responses in gene expression, whereas in the H-4-II-E cells, t11 was used for the same reason. Additional sets of culture dishes were incuba ted with FA alone, the PPAR inhibitor MK886 (10 M; Kehrer et al., 2001), or a combination of FA and MK886. After 24 h of incubation, cells were washed twice with 10 mL HBSS, ly sed with TriZol, and stored at -80oC until mRNA analysis. RNA Isolation and Analysis Total cellular RNA was isolated from cells using TriZol reagent according to the manufacturer’s instru ctions. Ten micrograms of tota l RNA was fractioned in a 1.0% agarose formaldehyde gel following previous ly described protocols (Ing et al., 1996) using the MOPS buffer (Fisher Scientific, Pitts burgh, PA) and transferred to a Biotrans

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117 nylon membrane by downward capillary transfer in 20X SSC (3 M NaCl, 0.3 M sodium citrate, pH 7.0) using the TurboBlotting sy stem (Schleicher and Schuel, Keene, NH). Nylon membranes were cross-linked by expos ure to a UV light source for 90 sec and baked at 80C for 1 h. Membranes were in cubated for 2 h at 50C in ultrasensitive hybridization buffer (ULTRAhyb; Ambion, Au stin, TX) followed by an overnight incubation at 50C in the same ULTRAhyb solution containing the 32P-labeled ACO, HMG-R, and Apo A-I cDNA probes. Probe s were generated by RT-PCR for ACO (forward 5’-CCGGAGCTGCTTACACACAT-3 ’; reverse 5’-GGTCATACGTGGCTGT GGTT-3’), HMG-R (forward 5’-TCCTTG GTGATGGGAGCTTGTTGTG-3’; reverse 5’-TGCGAACCCTTCAGATGTTTCGAGC-3’), human Apo A-I (forward 5’-AAGACA GCGGCAGAGACTAT-3’; reverse 5’-ATC TCCTCCTGCCACTTCTT-3’), and rat Apo A-I (forward 5’-AAGGACAGCGGCAGAGAC TA-3’; reverse 5’-CCACAACCTTTAG ATGCCTT-3’). The sizes and sequences of these cDNA probes were verified by DNA sequencing prior to their use in Northern blot analysis. Filters were sequentially washed in 2X SSC (1X= 0.15 M sodium chloride, 0.015 M sodium citrate)-0.1% SDS and in 0.1x SSC-0.1% SDS two times each at 50C and then exposed to X-ray film to detect radiolabeled bands. Equal loading of to tal RNA for each experimental sample was verified by comparison to 18S rR NA ethidium bromide staining. Lipid Extraction Total lipids were extracted from conditioned media as described by Bligh and Dyer (1959), with modifications. For each sample, 2 mL of conditioned media was aliquotted into a 20 mL glass screw-top vial. Fourteen mL of chloroform:methanol (2:1, v/v) was then added and the vials were vortexed for 5 minutes. The vials were then centrifuged at 1700 rpm for 5 min. The bottom lipid-containing chloroform layer was transferred to a

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118 clean, dry, pre-weighed vial, placed in a 37oC water bath, and dried under nitrogen gas. Dry samples were placed in a 50oC oven for 10 minutes and placed in a desiccator to cool to room temperature. Samples were we ighed, and lipid weight was determined by difference. The sample was resuspended in chloroform and stored at -20oC until HDL cholesterol analysis. HDL Cholesterol Assay Lipid extracts from conditioned media were analyzed using a commercially available L-Type HDL-C kit, following the ma nufacturer’s directions. Briefly, using a 96-well plate, 3 L of sample was pipetted in to each well. Two hundred seventy L of Enzyme Color Solution (R1) was added, and the plate was incubated for 5 minutes at 37oC. Ninety L of Reacting Solution (R2) was then added, and the plate was incubated another 5 minutes at 37oC. The absorbance at 600 nm was measured using the SpectraMax 340 PC microplate reader (Mol ecular Devices, Sunnyvale, CA), and the concentration of HDL cholesterol in each samples was calculated by plotting against a standard curve. Statistical Analysis All hybridization signals as measured by densitometry were evaluated by least squares analysis of variance (ANOVA) using the General Linear Model (GLM) procedure of the SAS software package (SAS Institute Inc, Cary, NC). In each experiment, treatments were run in duplic ate, and the whole experiment was also duplicated, giving n=4 plates per treatment The general model for mRNA analysis included experiment, treatment, and experi ment x treatment interaction. In mRNA analyses, densitometric values for target gene s were expressed as ratios of target gene densitometric values over the corresponding 18S rRNA densito metric values. For HDL

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119 cholesterol concentration, the sources of variation included experiment, treatment, experiment x treatment interac tion, and plate (experiment x treat ment). The plate, nested within experiment and treatment, was consid ered a random variable and therefore the plate variance was used as an error term to te st the effects of experiment, treatment, and experiment x treatment interaction. Treat ment means were further compared using preplanned orthogonal contrasts. These contrasts were control vs. fat treatment (c9, t9, c11, t11), double bonds in position 9 (c9, t9) vs. 11(c11, t11), c9 vs. t9, and c11 vs. t11. For all responses, the two cell lin es were analyzed separately. Results Effects of Octadecenoic Acids on HepG2 Cells In HepG2 cells, incubation with monounsaturated FA with bonds in the 11 position increased (+14%; P = 0.005) ACO mRNA expr ession as compared to MUFA with bonds in the 9 position. This effect was primarily due to the increase (+16%; P = 0.01) in ACO gene expression from c11 as compared to t11 (Figure 5-1). Steady-state levels of HMG-R mRNA were unaffected (P = 0.6) by any FA treatment (Figure 5-2). No differences were detected in Apo A-I gene ex pression due to double bond position (P = 0.2) or c9 as compared to t9 (P = 0.3). However, the c11 isomer up-regulated (+19%; P = 0.04) Apo A-I gene expression as co mpared to the t11 isomer (Figure 5-3). None of the FA studied had effects (P = 0.4) on HDL cholesterol production by HepG2 cells (Figure 5-4). Effects of Octadecenoic Acids on H-4-II-E Cells On average, incubation of H-4-II-E cells with FA up-regulated (+10%; P = 0.01) ACO mRNA expression as compared to contro l. Additionally, the c9 isomer increased (+11%; P = 0.02) ACO mRNA levels as compar ed to t9, and the t11 isomer increased

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120 (+10%; P = 0.03) gene expression as compared to c11 (Figure 5-5). The t11 FA isomer increased (+21%; P = 0.005) HMG-R gene expr ession as compared to the c11 isomer (Figure 5-6). As compared to control, in cubation with FA decreased (-10%; P = 0.002) Apo A-I mRNA concentrations, although ther e were no differences among the FA studied (Figure 5-7). High-de nsity lipoprotein cholesterol production by H-4-II-E cells was unaffected (P = 0.3) by a ny FA studied (Figure 5-8). Role of PPAR in Vaccenic Acid-Induced Effects on Gene Expression Incubation of HepG2 cells with 10 m WY 14,643, a specific PPAR agonist, had no effect on either basal (P = 0.06) or c 11-induced (P = 0.3) AC O mRNA expression (Figure 5-9). Basal levels of HMG-R mRNA were increased (+16%; P = 0.05) by use of the PPAR agonist, but there was no effect (P = 0.4) on c11-induced gene expression (Figure 5-10). Co-incubation with WY 14,643 enhanced (+33%; P = 0.007) the effect of c11 on Apo A-I mRNA concentration (Figure 5-11). In HepG2 cells, incubation with MK886, a specific PPAR inhibitor, had no effects (P > 0.2) on basal or c11-induced expression of any of the genes studied (A CO, Figure 5-12; HMG-R, Figure 5-13; Apo A-I, Figure 5-14). In H-4-II-E cells, incubation with WY 14,643 enhanced (+52%; P = 0.001) the effect of t11 on ACO gene expression (Fi gure 5-15). A similar effect was seen in HMG-R mRNA levels, with activation of PPAR increasing (+44%; P = 0.05) t11-induced gene expression (Fig ure 5-16). Activation of PPAR enhanced (+55%; P = 0.002) the effect of t11 on Apo A-I gene expression, as compared to t11 alone (Figure 5-17). Incubation with MK886 decreased (-55%; P = 0.0007) the basal level and attenuated (-169%; P < 0.0001) t11-induced ACO mRNA levels in H-4-II-E cells (Figure

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121 5-18). Inhibition of PPAR had no effects (P > 0.1) on ba sal or t11-induced HMG-R or Apo A-I gene expression (HMG-R, Figur e 5-19; Apo A-I, Figure 5-20). Discussion Dietary trans fatty acids have been implicated as potent negative factors in the development of numerous disease states, including dyslipidemia and cardiovascular disease. These changes may be mediated, in part, by the effects of fats on lipid metabolism; however, these effects may be differ ent for different isomers. In this study, we investigated the effects of cis and trans isomers of octadecenoic acid in human and rat hepatoma cell lines. In HepG2 (human ) cells, MUFA with the double bond in the 11 position increased ACO mRNA expression as co mpared to those FA with the bond in the 9 position. In particular, cis -vaccenic acid (c11) increased ACO gene expression. In contrast, in the H-4-II-E (rat) cells, all FA up-regulated ACO mRNA concentrations, with oleic acid (c9) and trans -vaccenic acid (t11) increasing gene expression as compared to elaidic acid (t9) and cis -vaccenic acid (c11), respectively. In contrast with our findings in gene expression, elaidic acid was shown to be a better substrate than oleic acid for fat oxidation, particularly peroxisomal oxidation, in rat hepatocytes (Guzman et al., 1999). However, in a human dietary study, supplementation with trans -FA increased and cis -FA decreased fat oxidation (as measured by indirect calorimetry) as compared to a saturated fat control diet (Lovejoy et al., 2002). Add itionally, in INS-1 cells, similar effects were seen as in H-4-II-E cells, with palmitate oxidation increasing when cells were incubated with oleic, elaidic, cis -vaccenic or trans -vaccenic acids. Though gene expression was not measured, trans -vaccenic acid increased oxidation to a greater extent than its cis counterpart (Alstr up et al., 2004).

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122 As numerous studies have s hown effects of trans-FA on cholesterol, we examined the mechanisms by which these fats may modul ate cholesterol producti on. One potential gene involved is HMG-R, the rate limiting enzyme in cholesterol synthesis. In HepG2 cells, HMG-R mRNA expression was unaffected by any FA st udied. In the rat cells, however, trans -vaccenic (t11) acid increased gene expression as compared to cis -vaccenic (c11) acid. Though evidence sugge sts that FA can affect cholesterol production, few studies have examined the role of HMG-R in this response. In support of our findings, dietary oleic aci d had no effect on HMG-R activity in Golden Syrian hamsters (Kurushima et al., 1995a; Kurushima et al., 1995b). These studies, however did not examine the effects of trans -FA. A recent study in mice showed no effects of dietary trans 18:1 fatty acids on HMG-R gene expression (Cassagno et al., 2005). Another factor involved in normal lipoprot ein profiles and metabolism is Apo A-I, the predominant apolipoprotein associat ed with HDL cholesterol. Dietary trans -FA have the ability to modulate plasma lipids, and may act, in part, by their effects on apolipoproteins. The effects of trans -FA on cholesterol production have been examined extensively, but results seem to depe nd on the model used. In HepG2 cells, cis -vaccenic acid (c11) increased Apo A-I mRNA levels as compared to trans -vaccenic acid (t11). These changes in gene expression did not co rrelate with HDL chol esterol production, as none of the FA treatments had any effects. In H-4-II-E cells, treatmen t with all of the FA decreased Apo A-I mRNA concentration, a lthough there were no differences among the FA studied. As with the human cells, these changes in gene expr ession did not affect HDL cholesterol production, as c oncentrations were not different among the treatments. In contrast with our findings, HDL cholesterol was decreased by trans -FA in two human

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123 dietary studies (Mensink and Katan, 1990; Tholstrup et al., 2006), although in the second study, the saturated to monounsaturated fat ratio of the diets may have played a significant role. In monkeys, dietary el aidic acid decreased Apo A-I and HDL cholesterol as compared to a saturated fat diet (Khosla et al., 1997). In HepG2 cells, oleic acid had no effects on Apo A-I or HDL chol esterol production (Das hti and Wolfbower, 1987), whereas elaidic acid may increase HDL cholesterol (Dashti et al., 2000). Numerous studies, however, suppor t our findings. In hamsters dietary oleic acid had no effects on HDL cholesterol production as compared to saturated fats (Kurushima et al., 1995b), LA (Kurushima et al., 1995 a; Nicolosi et al., 2004), or trans -FA (Nicolosi et al., 1998). When comparing vaccenic and elaidic acids, no effects on HDL cholesterol production were seen (Meijer et al., 2001). Serum and liver cholesterol concentrations were not different in Wist ar rats fed diets high in cis -FA, trans -FA or saturated FA (Colandre et al, 2003). A recent study in mice fed 3% of dietary energy as trans 18:1 FA showed no changes in total or HDL choles terol (Cassagno et al., 2005). In two human studies, HDL cholesterol was unaffected by diets rich in trans -FA as compared with those high in cis -FA (Judd et al., 1994; Lovejoy et al., 2002). As FA and their derivatives have been identified as potential ligands for PPARs, we investigated the possibility that FA eff ects in the two cell lines may be mediated by PPAR In HepG2 cells, incubation with WY 14,643, a selective PPAR agonist, increased basal expression of HMG-R mR NA and enhanced c11-induced Apo A-I mRNA expression. Activation of PPAR had no effects on basal or c11-induced ACO gene expression. Inhibition of PPAR by MK886 had no effects on the three genes studied. In contrast with the effects seen in HepG2 cells, activation of PPAR enhanced

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124 the effects of t11 on ACO, HMG-R, and Apo A -I gene expression in the H-4-II-E cells. Inhibition of PPAR decreased the basal levels a nd attenuated the effects of trans vaccenic acid on ACO gene expression. 3-hydr oxy, 3-methylglutaryl CoA reductase and Apo A-I mRNA concentrations were unaffected by inhibition of PPAR Although ACO is an established PPAR responsive gene (Tugwood, et al ., 1992), species differences do exist. It is questionable whether the PPA R response element of human ACO is active (Woodyatt et al., 1999). Dietary studies have shown that rodents are responsive to the effects of PPAR activation, but non-rodent species, such as primates and guinea pigs, are resistant or unresponsive to some of the negative effects (Bentley et al., 1993; Cattley et al., 1998). In a comprehens ive analysis of gene expressi on in human and rat hepatoma cells by microarray analysis, only ra t ACO mRNA was responsive to WY 14,643 (Vanden Heuvel et al., 2003). Other genes that may be differentially regulated in human and rat liver include cytosolic aspartate aminotransferase (Tom kiewicz et al., 2004), peroxisomal 3-oxoacyl-CoA thiolase (L awrence et al., 2001), and catalase (Ammerschlaeger et al., 2004). Additionally, different PPAR agonists may regulate lipid metabolism in a compound-dependent manner. A recent study by Duez and coworkers (2005) showed that, in mice, fenofib rate and gemfibrozil, both stimulate ACO mRNA expression, but only fenofibrate greatly induces Apo A-I gene expression. Summary In HepG2 cells, only treatment with the cis -11 fatty acid up-regulated ACO and Apo A-I mRNA expression. 3-hydroxy, 3-met hylglutaryl CoA reduc tase steady-state mRNA concentrations were una ffected by treatment with any cis or trans MUFAs. High-density lipoprotein cholesterol produc tion was unchanged by any FA studied. Activation of PPAR increased basal concentrations of HMG-R mRNA and enhanced

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125 c11-induced Apo A-I gene expr ession, but no effects of PPAR inhibition were seen on any gene studied in HepG2 cells. These resu lts are consistent with the low levels of endogenous PPAR expression in this cell line (Hsu et al., 2001). In H-4-II-E cells, incubation with cis and trans FA increased ACO mRNA and decreased Apo A-I mRNA levels, but ther e were no differences among FA effects on either gene. The trans -11 isomer increased HMG-R mRNA expression as compared to the cis -11 isomer. None of the FA studied effected HDL cholesterol production by H-4-II-E cells. Selective activation of PPAR in H-4-II-E cells enhanced t11-induced expression of ACO, HMG-R and Apo A-I ge ne transcripts. Inhibition of PPAR decreased basal expression and attenuated t11-induced ACO mRNA concentrations. However, no effects were seen on HMG-R or Apo A-I mRNA. These results indicate that t11-induced ACO gene expr ession may be mediated by PPAR in the H-4-II-E cells, whereas effects on HMG-R and Apo A-I genes may be independent of PPAR As responses to FA and PPAR activation and inhibition were different in the human and rat cells lines, net effects are likely species specific.

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126 Figure 5-1. Effect of cis and trans isomers of octadece noic acid on ACO mRNA expression in HepG2 cells. Ten microgr ams of total cellular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representati ve Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: Control vs. (c9 + t9 + c11 + t11), P = 0.06; Cont rast 2: (c9 + t9) vs. (c11 + t11), P = 0.005; Contrast 3: c9 vs. t9, P = 0.98; Contrast 4: c11 vs. t11, P = 0.01. Treatments Controlc9t9c11t11 ACO mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 C c9t9c11t11 3.2kb 18S ACO rRNA (A) (B)

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127 Figure 5-2. Effect of cis and trans isomers of octadecenoic acid on HMG-R mRNA expression in HepG2 cells. Ten microgr ams of total cellular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representati ve Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: Control vs. (c9 + t9 + c11 + t11), P = 0.99; Cont rast 2: (c9 + t9) vs. (c11 + t11), P = 0.2; Contrast 3: c9 vs. t9, P = 0.7; Contra st 4: c11 vs. t11, P = 0.7. Treatments Controlc9t9c11t11 HMG-R mRNA (Normalized to 18S) 0.00 0.05 0.10 0.15 0.20 0.25 C c9t9c11t11 18S 4.1-4.7kb HMG-R rRNA (A) (B)

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128 Figure 5-3. Effect of cis and trans isomers of octadecenoic acid on Apo A-I mRNA expression in HepG2 cells. Ten microgr ams of total cellular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representati ve Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: Control vs. (c9 + t9 + c11 + t11), P = 0.9; Contrast 2: (c9 + t9) vs. (c11 + t11), P = 0.2; Contrast 3: c9 vs. t9, P = 0.3; Contrast 4: c11 vs. t11, P = 0.04. Treatments Controlc9t9c11t11 Apo A-I mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 0.5 C c9t9c11t11 18S 0.9 kb Apo A-I rRNA (A) (B)

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129 Figure 5-4. Effects of cis and trans isomers of octadecenoic acid on HDL cholesterol production by HepG2 cells. Data repr esents least square means SEM calculated over two experiments. To fu rther examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: Control vs. (c9 + t9 + c11 + t11), P = 0.7; Contrast 2: (c9 + t9) vs. (c11 + t 11), P = 0.8; Contrast 3: c9 vs. t9, P = 0.6; Contrast 4: c11 vs. t11, P = 0.09. Treatment Controlc9t9c11t11 HDL Cholesterol (mg/dL) 0.0 0.5 1.0 1.5 2.0 2.5 3.0 b

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130 Figure 5-5. Effect of cis and trans isomers of octadece noic acid on ACO mRNA expression in H-4-II-E cells. Ten micr ograms of total cell ular RNA isolated from control and treated H-4-II-E cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representati ve Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: Control vs. (c9 + t9 + c11 + t11), P = 0.01; Cont rast 2: (c9 + t9) vs. (c11 + t11), P = 0.3; Contrast 3: c9 vs. t9, P = 0.02; Contra st 4: c11 vs. t11, P = 0.03. Treatments Controlc9t9c11t11 ACO mRNA (normalized to 18S) 0.0 0.1 0.2 0.3 0.4 C c9t9c11t11 3.2kb 18S ACO rRNA (A) (B)

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131 Figure 5-6. Effect of cis and trans isomers of octadecenoic acid on HMG-R mRNA expression in H-4-II-E cells. Ten micr ograms of total cell ular RNA isolated from control and treated H-4-II-E cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representati ve Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: Control vs. (c9 + t9 + c11 + t11), P = 0.99; Cont rast 2: (c9 + t9) vs. (c11 + t11), P = 0.1; Contrast 3: c9 vs. t9, P = 0.4; Contra st 4: c11 vs. t11, P = 0.005. Treatments Controlc9t9c11t11 HMG-R mRNA (normalized to 18S) 0.00 0.02 0.04 0.06 0.08 0.10 0.12 C c9 t9 c11 t11 18S 4.5kb HMG-R rRNA (A) (B)

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132 Figure 5-7. Effect of cis and trans isomers of octadecenoic acid on Apo A-I mRNA expression in H-4-II-E cells. Ten micr ograms of total cell ular RNA isolated from control and treated H-4-II-E cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representati ve Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: Control vs. (c9 + t9 + c11 + t11), P = 0.002; Contrast 2: (c9 + t9) vs. (c11 + t11), P = 0.1; Contrast 3: c9 vs. t9, P = 0.6; Contra st 4: c11 vs. t11, P = 0.3. Treatments Controlc9t9c11t11 Apo A-I mRNA (normalized to 18S) 0.00 0.02 0.04 0.06 0.08 0.10 0.12 0.14 C c9t9 c11 t11 18S 0.9 kb Apo A-I rRNA (A) (B)

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133 Figure 5-8. Effects of cis and trans isomers of octadecenoic acid on HDL cholesterol production in H-4-II-E cells. Data re presents least square means SEM calculated over two experiments. To fu rther examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: Control vs. (c9 + t9 + c11 + t11), P = 0.7; Contrast 2: (c9 + t9) vs. (c11 + t 11), P = 0.5; Contrast 3: c9 vs. t9, P = 0.09; Contrast 4: c11 vs. t11, P = 0.2. Treatment Controlc9t9c11t11 HDL Cholesterol (mg/dL) 0.0 0.5 1.0 1.5 2.0 2.5 f

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134 Figure 5-9. Effect of WY 14,643 on ACO mRNA response to cis -vaccenic acid in HepG2 cells. Ten micrograms of total cellular RNA isolated from control and treated HepG2 cells were subjected to Nort hern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (c11 + c11+A), P = 0.4; C ontrast 2: c11 vs. c11+A, P = 0.3; Contrast 3: Contro l vs. Agonist, P = 0.06. Treatments Controlc11Agonistc11 + Agonist ACO mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 18S 3.2kb C c11 PPAR Agonist c11 + Agonist ACO rRNA (A) (B)

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135 Figure 5-10. Effect of WY 14,643 on HMG-R mRNA response to cis -vaccenic acid in HepG2 cells. Ten micrograms of total ce llular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (c11 + c11+A), P = 0.2; C ontrast 2: c11 vs. c11+A, P = 0.4; Contrast 3: Contro l vs. Agonist, P = 0.05. Treatments Controlc11Agonistc11 + Agonist HMG-R mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 18S 4.1-4.7kb C c1 PPAR Agonist c11 + Agonist (A) (B) HMG-R rRNA

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136 Figure 5-11. Effect of WY 14,643 on Apo A-I mRNA response to cis -vaccenic acid in HepG2 cells. Ten micrograms of total ce llular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (c11 + c11+A), P = 0.5; C ontrast 2: c11 vs. c11+A, P = 0.007; Contrast 3: Contro l vs. Agonist, P = 0.4. Treatments Controlc11Agonistc11 + Agonist Apo A-I mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 18S 0.9 kb C c11 PPAR Agonist c11 + Agonist Apo A-I rRNA (A) (B)

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137 Figure 5-12. Effect of MK886 on ACO mRNA response to cis -vaccenic acid in HepG2 cells. Ten micrograms of total cellular RNA isolated from control and treated HepG2 cells were subjected to Nort hern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). Contrast 1: (Control + Inhib) vs. (c11 + c11+I), P = 0.2; Contrast 2: c11 vs. c 11+I, P = 0.7; Contrast 3: Control vs. Inhib, P = 0.4. Treatments Controlc11Inhibc11 + Inhib ACO mRNA (Normalized to 18S) 0.00 0.05 0.10 0.15 0.20 0.25 0.30 18S 3.2kb C c11 PPAR Inhib c11 + Inhib ACO rRNA (A) (B)

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138 Figure 5-13. Effect of MK886 on HMG-R mRNA response to cis -vaccenic acid in HepG2 cells. Ten micrograms of total ce llular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). Contrast 1: (Control + Inhib) vs. (c11 + c11+I), P = 0.6; Contrast 2: c11 vs. c 11+I, P = 0.4; Contrast 3: Control vs. Inhib, P = 0.2. Treatments Controlc11Inhibc11 + Inhib HMG-R mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 18S 4.1-4.7kb C c11 PPAR Inhib c11 + Inhib HMG-R rRNA (A) (B)

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139 Figure 5-14. Effect of MK886 on Apo A-I mRNA response to cis -vaccenic acid in HepG2 cells. Ten micrograms of total ce llular RNA isolated from control and treated HepG2 cells were subjected to Northern blot analysis, and resulting densitometric values were analyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). Contrast 1: (Control + Inhib) vs. (c11 + c11+I), P = 0.5; Contrast 2: c11 vs. c 11+I, P = 0.6; Contrast 3: Control vs. Inhib, P = 0.9. Treatments Controlc11Inhibc11 + Inhib Apo A-I mRNA (Normalized to 18S) 0.0 0.1 0.2 0.3 0.4 0.5 18S 0.9 kb C c11 PPAR Inhib c11 + Inhib Apo A-I rRNA (A) (B)

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140 Figure 5-15. Effect of WY 14,643 on ACO mRNA response to transvaccenic acid in H-4-II-E cells. Ten micrograms of tota l cellular RNA isolated from control and treated H-4-II-E cells were subject ed to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (t11 + t11+A), P = 0.03; C ontrast 2: t11 vs. t11+A, P = 0.001; Contrast 3: Contro l vs. Agonist, P = 0.4. Treatments Controlt11Agonistt11 + Agonist ACO mRNA (normalized to 18S) 0.0 0.1 0.2 0.3 0.4 0.5 18S 3.2kb C t11 PPAR Agonist t11 + Agonist ACO rRNA (A) (B)

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141 Figure 5-16. Effect of WY 14,643 on HMG-R mRNA response to trans -vaccenic acid in H-4-II-E cells. Ten micrograms of tota l cellular RNA isolated from control and treated H-4-II-E cells were subject ed to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (t11 + t11+A), P = 0.0005; C ontrast 2: t11 vs. t11+A, P = 0.0007; Contrast 3: Contro l vs. Agonist, P = 0.06. Treatments Controlt11Agonistt11 + Agonist HMG-R mRNA (normalized to 18S) 0.00 0.05 0.10 0.15 0.20 18S 4.5kb C t11 PPAR Agonist t11 + Agonist (A) (B) HMG-R rRNA

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142 Figure 5-17. Effect of WY 14,643 on Apo A-I mRNA response to trans -vaccenic acid in H-4-II-E cells. Ten micrograms of tota l cellular RNA isolated from control and treated H-4-II-E cells were subject ed to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (t11 + t11+A), P = 0.3; C ontrast 2: t11 vs. t11+A, P = 0.002; Contrast 3: Contro l vs. Agonist, P = 0.01. Treatments Controlt11Agonistt11 + Agonist Apo A-I mRNA (normalized to 18S) 0.00 0.02 0.04 0.06 18S 0.9 kb C t11 PPAR A g onist t11 + Agonist Apo A-I rRNA (A) (B)

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143 Figure 5-18. Effect of MK886 on ACO mRNA response to transvaccenic acid in H-4-II-E cells. Ten micrograms of tota l cellular RNA isolated from control and treated H-4-II-E cells were subject ed to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (t11 + t11+I), P = 0.004; C ontrast 2: t11 vs t11+I, P < 0.0001; Contrast 3: Control vs. Inhib, P = 0.0007. Treatments Controlt11Inhibt11 + Inhib ACO mRNA (normalized to 18S) 0.00 0.02 0.04 0.06 0.08 18S 3.2kb C t11 PPAR Inhib t11 + Inhib ACO rRNA (A) (B)

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144 Figure 5-19. Effect of MK886 on HMG-R mRNA response to trans -vaccenic acid in H-4-II-E cells. Ten micrograms of tota l cellular RNA isolated from control and treated H-4-II-E cells were subject ed to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (t11 + t11+I), P = 0.08; Contrast 2: t11 vs. t11+I, P = 0.5; Contrast 3: Control vs. Inhib, P = 0.5. Treatments Controlt11Inhibt11 + Inhib HMG-R mRNA (normalized to 18S) 0.00 0.02 0.04 0.06 0.08 0.10 0.12 18S 4.5kb C t11 PPAR Inhib t11 + Inhib HMG-R rRNA (A) (B)

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145 Figure 5-20. Effect of MK886 on Apo A-I mRNA response to transvaccenic acid in H-4-II-E cells. Ten micrograms of tota l cellular RNA isolated from control and treated H-4-II-E cells were subject ed to Northern blot analysis, and resulting densitometric values were an alyzed by the GLM procedure of SAS. A) A representative Northern blot. B) Means SEM calculated over two experiments (n = 4 for each treatment). To further examine treatment effects, means were separated using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (t11 + t11+I), P < 0.0001; C ontrast 2: t11 vs. t11+I, P = 0.2; Contrast 3: Control vs. Inhib, P = 0.1. Treatments Controlt11Inhibt11 + Inhib Apo A-I mRNA (normalized to 18S) 0.00 0.02 0.04 0.06 0.08 0.10 0.12 18S 0.9kb C t11 PPAR Inhib t11 + Inhib Apo A-I rRNA (A) (B)

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146 Figure 5-21. Regulation of lipid metabolizi ng genes and HDL cholesterol production by cis and trans octadecenoic fatty acids. In HepG2 cells, cis -vaccenic acid (c11) up-regulated ACO and Apo A-I gene expression. Ac tivation of PPAR increased basal expression of HMG-R mRNA and enhanced c11-induced Apo A-I gene expression. In H4-II-E cells, all fatty aci ds studied increased ACO and Apo A-I gene expression to the same extent. Trans -vaccenic acid (t11) increased HMG-R mRNA concentr ations. Activation of PPAR enhanced the effects of t11 on ACO, HMG-R and Apo A-I gene expression. Inhibition of PPAR decreased basal expression and attenuated t11induced ACO mRNA levels. ACO mRNA HMG-R mRNA Apo A-I mRNA ACO mRNA HMG-R mRNA Apo A-I mRNA PPAR c 9 t 9 c 11 t 11 Activation Inhibition HepG2 H-4-II-E ?

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147 CHAPTER 6 GENERAL DISCUSSION Dietary fat has been implicated as a ma jor factor in many areas of health and disease. Though historically considered primarily as an energy source and cell membrane constituent, health effects are more likely due to fatty acid effects on gene expression and the subsequent effects on metabo lism. However, it has been suggested by numerous studies that all fatty acids (FA) may not have the same effects. In these studies, both human and rat hepatoma cells we re used as models, as it also has been suggested that species differe nces exist in fatty acid metabolism (Bergen and Mersmann, 2005). In the first experiment, we examined the ro le of fatty acids with differing saturation and bond position on genes involved in -oxidation, HDL choleste rol synthesis, and high-density lipoprotein (HDL) c holesterol secretion into the culture media. In HepG2 (human) cells, acyl CoA oxidase (ACO) mR NA expression was unaffected by stearic (ST), linoleic (LA), linolenic (LNA), or eicosa pentaenoic (EPA) acids. In contrast, in the H-4-II-E (rat) cells, ACO mR NA expression was induced by ST. Consistent with our findings in rat cells, pigs fed a tallow-based diet high in saturated fat had an increased concentration of ACO mRNA as compared to fish oil-fed pi gs (Ding et al., 2003). Other studies, however, have reported up-regulati on of ACO mRNA in rat liver by dietary polyunsaturated fatty acids (PUFA) as well as saturated fats (Ber thou et al., 1995). In HepG2 cells, it has been reported that PU FAs of differing saturation and length can regulate ACO mRNA in a dose-de pendent and differential manner (Rise and Galli, 1999).

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148 In a human retinoblastoma cell line, low c oncentrations of supplemental n-3 PUFA increased ACO mRNA, whereas high concentrati ons of the FA decreased it (Langelier et al., 2003). In numerous animal models and human diet ary and epidemiological studies, fatty acids have been demonstrated to have the ability to modulat e serum cholesterol concentrations in both a positive and negativ e manner. 3-hydroxy, 3-methylglutaryl CoA reductase (HMG-R) is the rate limiting enzyme in cholesterol synthesis, and its inhibition is the target of the stat in class of drugs, used in the trea tment of hyperlipidemias. If select dietary FA stimulate HMG-R expression, th en they may be important FA to limit consumption of in order to prevent elevat ion of blood cholesterol concentrations in humans. In this study, we showed that in HepG2 cells, HMG-R mRNA was up-regulated by ST as compared to the PUFAs, whereas in the H-4-II-E cells, it was up-regulated by both ST and EPA. Consistent with our fi ndings in rodent cells, in Reuber H35 rat hepatoma cells, a cell line closely related to H-4-II-E cells, incubation with either saturated fats or PUFAs increased HMG-R en zyme activity (Garcia-Pelayo et al., 2003). Enzyme activity of HMG-R has also been shown to be increased in mice fed a diet high in PUFAs (Kuan and Dupont, 1989). 3-hydroxy 3-methylglutaryl CoA reductase mRNA was increased to a greater extent in C3H mice fed a PUFA diet than in mice fed a saturated fat diet (Che ema and Agellon, 1999). Apolipoprotein A-I (Apo A-I) is the predominant lipoprotein associated with HDL cholesterol and is essential for its normal me tabolism. Deletion of the Apo A-I gene in humans results in very low blood concentr ations of HDL cholesterol and premature coronary artery disease (Sch aefer et al., 1982). Dietary fat has the ability to modulate

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149 plasma lipids, and may act, in part, by e ffects on apolipoproteins. In this study, we determined that, in HepG2 cells, Apo A-I mR NA was up-regulated by all FA. However, no change was detected in HDL cholesterol conc entration in the culture media. This is supported by a study by Dashti and coworkers (2002) in which HDL concentration was not different between linoleic acid (LA) and saturated fa t-treated HepG2 cells. In Golden-Syrian hamsters, an effective model for human diet and blood lipid interactions, feeding canola and soybean oils (unsaturated FA) increased Apo A-I mRNA expression as compared to a butter-containing diet, t hough HDL concentrations were lowered in the hamsters fed diets containing uns aturated as compared to butter fats (Dorfman et al., 2005). In the H-4-II-E cells, ST increased Apo A-I mRNA concentrat ion to the greatest extent. In contrast to current findings, Spra gue-Dawley rats fed diets high in saturated fat or PUFAs showed no differences in Apo A-I mRNA levels (Hatahet et al., 2003). However, the saturated fat diet contained primarily palmitic acid, not stearic acid, as in this study. Numerous beneficial physio logical effects have been attributed to conjugated linoleic acid (CLA), though these effects ma y be both isomer and species specific. Therefore, in the second experiment, we ex amined the effects of the rwo biologically active CLA isomers on key regulatory genes of lipid metabolism. In HepG2 cells, ACO mRNA expression was up-regulated by CLA as compared to LA-treated cells, with an increase additionally seen with incubation of trans -10, cis -12 CLA as compared with cis -9, trans -11 CLA. In contrast, all FA increas ed ACO mRNA as compared to control in H-4-II-E cells, but there were no differences among the FA studied. Several animal and cell models have also reported similar eff ects. Feeding mice a mix of (Peters et al.,

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150 2001) or individual CLA isomers (Warren et al., 2003; Degrace et al ., 2004) resulted in an increased expression of ACO mRNA as compar ed to control animals. In two studies, the increases detected in ACO mRNA levels in CLA-fed mice also coincided with increases in enzyme activity (Takahashi et al., 2003; Ide, 2005). In FaO cells, a rat hepatoma cell line derived from H4IIEC3 cells (Bayly et al., 1993), ACO gene expression was increased with 200 M cis -9, trans -11 CLA. This effect was not seen, however, with lower concentrations of CLA (Moya-Camarena et al., 1999). In a recent study using a hamster model, ACO activity was increased by trans -10, cis -12 CLA as compared to control or cis -9, trans -11 CLA (Macarulla et al., 2005). Together with our findings, these studies suggest a role fo r CLA in increasing liver peroxisomal -oxidation in the liver. When we examined CLA effects on genes involved in cholesterol synthesis in HepG2 cells, CLA isomers increased HMG-R mRNA transcript as compared to the parent molecule, LA. Between the CLA isomers, trans -10, cis -12 CLA up-regulated HMG-R mRNA concentrations as compared with the cis -9, trans -11 isomer. This differed in the H-4-II-E cells, where, on av erage, CLA increased gene expression as compared to LA, but no differences were s een between the two CLA isomers. Although the effects of saturated fats and PUFA on HMG-R gene expression and enzyme activity have been examined, few studies have explor ed the role of CLA. In a recent study, HMG-R activity was decreased in rats fed diacylglycerol-enriched structured lipids containing CLA as compared to those fed lipids without CLA or those fed corn oil (Kim et al., 2006). Though this differs from our findings, gene expression was not measured,

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151 and as with other target gene s, it would not be surprising if species and model-specific differences exist. In general, CLA-induced changes in the blood lipid profile observed in various models are conflicting. In our study, Apo A-I gene transcri pt was increased by the CLA isomers as compared to LA, and by trans -10, cis -12 CLA as compared to cis -9, trans -11 CLA in HepG2 cells. High-density lipoprotei n cholesterol production in HepG2 cells was decreased by the CLA isomers as compared to LA, with the effect predominantly derived from the decrease due to cis -9, trans -11 CLA. In H-4-II-E cells, steady-state levels of Apo A-I mRNA was decreased by LA and the CLA isomers; however, there were no differences among the FA studied. These decreases in gene expression did not result in increased HDL cholesterol producti on, as levels were not different among the treatments. The different responses seen in the two cell lines is reflected by conflicting responses in other species studi ed. Mice supplemented with cis -9, trans -11 or trans -10, cis -12 CLA showed no differences in Apo A -I mRNA concentrations in control or CLA-fed animals (Warren et al., 2003). In contrast, in apo-E deficient mice, dietary trans -10, cis -12 CLA decreased plasma Apo A-I levels as compared to cis -9, trans -11 CLA (Arbones-Mainar et al., 2006). Simila r to the decrease in HDL cholesterol production in HepG2 cells, rabbits fed a mixtur e of CLA isomers showed an increase in total serum cholesterol and a decrease in HDL cholesterol (Kritchevsky et al., 2000). Additionally, in the Syrian Gold en hamster, diets containing trans -10, cis -12 CLA increased HDL cholesterol as compared with LA or cis -9, trans -11 CLA diets (Mitchell et al., 2005). However, some studies in rat models have shown no effect of dietary CLA on serum HDL cholesterol (Kloss et al., 2005) In a human dietary study, both Apo A-I

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152 gene transcript and HDL cholesterol were decreased by consumpti on of CLA-enriched butter as compared to pre-supplement levels (Desroches et al., 2005). In the third experiment, we examined the effects of cis and trans isomers of octadecenoic acid on lipid metabolism. Dietary trans -FA have been implicated as potent negative factors in the development of nume rous disease states, including dyslipidemia and cardiovascular disease. These changes ma y be mediated, in part, by the effects of fats on lipid metabolism; however, these effect s may be different for different isomers. In HepG2 cells, MUFAs with the double bond in the 11 position increased ACO mRNA expression as compared to those FA with the bond in the 9 position. In particular, cis -vaccenic acid (c11) increased ACO gene e xpression. In contrast, in the H-4-II-E cells, all FA up-regulated ACO mRNA concentrations, with oleic acid (c9) and trans -vaccenic acid (t11) increasi ng gene expression as compared to elaidic acid (t9) and cis -vaccenic acid (c11), respectively. In contrast with our findings in gene expression, elaidic acid was shown to be a better substrate than oleic acid for fat oxidation, particularly peroxisomal oxidation, in rat he patocytes (Guzman et al., 1999). However, in a human dietary study, supplementation with trans -FA increased and cis -FA decreased fat oxidation (as measured by indirect calorimet ry) as compared to a saturated fat control diet (Lovejoy et al., 2002). Additionally, in IN S-1 cells, similar effects were seen as in H-4-II-E cells, with palmitate oxidati on increasing with oleic, elaidic, cis -vaccenic and trans -vaccenic acids. Though gene expression was not measured, trans -vaccenic acid increased oxidation to a greater extent than its cis counterpart (Alstr up et al., 2004). In HepG2 cells, HMG-R mRNA expression was unaffected by any of the FA studied. In the rat cells, however, trans -vaccenic (t11) acid increased gene expression as

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153 compared to cis -vaccenic (c11) acid. Though eviden ce suggests that FA can affect cholesterol production, few studies have examined the role of HMG-R in this response. In support of our findings, dietary oleic acid had no effect on HMG-R activity in Golden Syrian hamsters (Kurushima et al., 1995a; Kurushima et al., 1995b). These studies, however did not examine the effects of trans -FA. A recent study in mice showed no effects of dietary trans 18:1 fatty acids on HMG-R gene expression (Cassagno et al., 2005). Dietary trans -FA have the ability to modulate pl asma lipids, and may act, in part, by effects on apolipoprote ins. The effects of trans -FA on cholesterol production have been examined extensively, but results seem to depend on the model used. In HepG2 cells, cis -vaccenic acid (c11) increased Apo A -I mRNA levels as compared to trans -vaccenic acid (t11). These changes in gene expression did not correlate with HDL cholesterol production, as none of the FA treatments had any effects. In H-4-II-E cells, treatment with all of the monoenes decreased Apo A-I mR NA concentration, although there were no differences among the monoenes studied. As with the human cells, these changes in gene expression did not affect HDL cholesterol production, as concentrations were not different among the treatments. In contrast with our findings, HDL cholesterol concentration was decreased by consumption of trans -FA in two human studies (Mensink and Katan, 1990; Tholstrup et al., 2006), al though in the second st udy, the saturated to monounsaturated fat ratio of the diets may have played a significant role. In monkeys, dietary elaidic acid decreased Apo A-I and HDL cholesterol as compared to a saturated fat diet (Khosla et al., 1997). In HepG2 ce lls, oleic acid had no effects on Apo A-I or HDL cholesterol production (D ashti and Wolfbower, 1987), whereas elaidic acid may

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154 increase HDL cholesterol (Dashti et al., 2000). Numerous studies however, support our findings. In hamsters, dietary oleic acid had no effects on HDL c holesterol production as compared to saturated fats (Kurushima et al., 1995b), LA (Kurushima et al., 1995a; Nicolosi et al., 2004), or trans -FA (Nicolosi et al., 1998). When comparing vaccenic and elaidic acids, no effects on HDL cholesterol pr oduction were seen (M eijer et al., 2001). A recent study in mice fed 3% of dietary energy as trans 18:1 fatty acids reported no change in total or HDL cholesterol (Cassa gno et al., 2005). In two human studies, blood HDL cholesterol concentration wa s unaffected by diets rich in trans -FA as compared with those high in cis -FA (Judd et al., 1994; Lovejoy et al., 2002). Fatty acids and their derivatives have b een identified as potential ligands for peroxisome proliferator-activated receptors (PPAR). Therefore, we tested the hypothesis that fatty acid effects on gene expression may be mediated by PPAR In the three experiments, activation of PPAR by WY 14,643 or inhibition by MK886 had marginal effects on basal or fatty acid-induced gene e xpression in HepG2 cells. These results are consistent with the low levels of endogenous PPAR expression in this cell line (Hsu et al., 2001). In contrast, activation of PPAR consistently increased basal expression and enhanced STand trans -vaccenic-induced ACO gene expression in H-4-II-E cells. Although ACO is an established PPAR responsive gene (Tugwood, et al., 1992), species differences do exist. It is questionable whether the PPAR response element of human ACO is active (Woodyatt et al., 1999). Dietar y studies have show n that rodents are responsive to the effects of PPAR activation, but non-rodent species, such as primates and guinea pigs, are resistant or unresponsive to some of the negativ e effects (Bentley et al., 1993; Cattley et al., 1998). In a comprehe nsive analysis of gene expression in human

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155 and rat hepatoma cells by microarray anal ysis, only rat ACO mR NA was responsive to WY 14,643 (Vanden Heuvel et al., 2003). Ot her genes that may be differentially regulated in human and rat liver include cytosolic aspartate aminotransferase (Tomkiewicz et al., 2004), peroxisomal 3-oxo acyl-CoA thiolase (Lawrence et al., 2001), and catalase (Ammerschlaeger et al., 2004). Consistent w ith our findings in each experiment with the rat cell lin e, Apo A-I gene expression was not different from controls in mice fed fenofibrate, a potent PPAR activator (Warren et al., 2003). However, different PPAR agonists may regulate lipid metabolism in a compound-dependent, as well as species-dependent, manner. A recent study by Duez and coworkers (2005) showed that, in mice, fenofibrate a nd gemfibrozil, both stimulate ACO mRNA expression, but only fenofibrate greatly induces Apo A-I gene expression. Together with our findings, these results indi cate that fatty acids may differentially regulate specific lipid metabolizing genes in the liver through a PPAR -dependent mechanism. However, due to different responses in the human and rat hepatoma cell lines, the net effects are likely species specific.

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156 APPENDIX LS MEANS AND P-VALUES FOR ANALYSIS OF FATTY ACID EFFECTS ON LIPID-METABOLIZING GENES AND HDL CHOLESTEROL PRODUCTION IN HEPG2 AND H-4-II-E CELLS Table A-1. Effects of n-3 and n-6 FA on lip id-metabolizing genes and HDL cholesterol production in HepG2 cells. Response LS Means P-value Control ST LA LNA EPA SEM Exp Trt E x T ACO 0.477 0.551 0.432 0.498 0.412 0.048 0.40 0.33 0.32 HMG-R 0.613 0.754 0.600 0.575 0.564 0.043 0.03 0.06 0.27 Apo A-I 0.396 0.458 0.447 0.464 0.500 0.029 0.002 0.23 0.02 HDL, mg/dL 2.85 2.43 3.30 3.15 3.36 0.802 0.27 0.90 0.35 Table A-2. Effects of n-3 and n-6 FA on lip id-metabolizing genes and HDL cholesterol production in H-4-II-E cells. Response LS Means P-value Control ST LA LNA EPA SEM Exp Trt E x T ACO 0.077 0.109 0.084 0.089 0.069 0.007 0.21 0.02 0.90 HMG-R 0.160 0.201 0.115 0.111 0.218 0.011 0.15 0.00010.99 Apo A-I 0.043 0.059 0.036 0.028 0.041 0.003 0.31 0.00020.22 HDL, mg/dL 6.33 3.20 1.61 13.15 2.08 1.01 0.05 0.04 0.03

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157 Table A-3. Effects of WY 14,643 on mRNA responses to ST in HepG2 cells. Response LS Means P-value Control ST AgonistST + ASEM Exp Trt E x T ACO 0.227 0.245 0.229 0.226 0.005 0.0001 0.11 0.56 HMG-R 0.251 0.274 0.190 0.248 0.007 0.002 0.0001 0.90 Apo A-I 0.179 0.192 0.162 0.168 0.018 0.24 0.65 0.76 Table A-4. Effects of MK886 on mRNA responses to ST in HepG2 cells. Response LS Means P-value Control ST InhibitorST + I SEM Exp Trt E x T ACO 0.157 0.260 0.286 0.289 0.033 0.39 0.07 0.03 HMG-R 0.231 0.301 0.365 0.338 0.029 0.01 0.05 0.15 Apo A-I 0.319 0.437 0.417 0.458 0.025 0.003 0.02 0.18

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158 Table A-5. Effects of WY 14,643 on mRNA responses to ST in H-4-II-E cells. Response LS Means P-value Control ST Agonist ST + A SEM Exp Trt E x T ACO 0.039 0.043 0.050 0.069 0.003 0.01 0.0006 0.002 HMG-R 0.074 0.080 0.051 0.062 0.005 0.002 0.01 0.03 Apo A-I 0.019 0.030 0.022 0.042 0.002 0.0008 0.0001 0.0001 Table A-6. Effects of MK886 on mRNA re sponses to ST in H-4-II-E cells. Response LS Means P-value Control ST InhibitorST + I SEM Exp Trt E x T ACO 0.237 0.258 0.211 0.201 0.008 0.0004 0.005 0.01 HMG-R 0.068 0.101 0.096 0.100 0.005 0.0009 0.003 0.09 Apo A-I 0.028 0.040 0.014 0.028 0.003 0.002 0.003 0.16

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159 Table A-7. Effects of CLA on lipid-metabo lizing genes and HDL cholesterol production in HepG2 cells. Response LS Means P-value Control LA c9, t11 CLA t10, c12 CLA SEM Exp Trt E x T ACO 0.316 0.316 0.335 0.430 0.020 0.0001 0.01 0.07 HMG-R 0.295 0.219 0.311 0.396 0.018 0.54 0.0008 0.16 Apo A-I 0.338 0.287 0.317 0.406 0.023 0.006 0.04 0.01 HDL, mg/dL 2.62 2.76 2.05 2.64 0.151 0.002 0.08 0.03 Table A-8. Effects of CLA on lipid-metabo lizing genes and HDL cholesterol production in H-4-II-E cells. Response LS Means P-value Control LA c9, t11 CLA t10, c12 CLA SEM Exp Trt E x T ACO 0.161 0.202 0.201 0.223 0.007 0.0007 0.002 0.28 HMG-R 0.072 0.058 0.101 0.087 0.006 0.003 0.005 0.14 HMG-R 0.056 0.040 0.047 0.041 0.003 0.0001 0.03 0.02 HDL, mg/dL 2.46 11.43 8.87 1.30 3.28 0.22 0.65 0.65

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160 Table A-9. Effects of WY 14,643 on mRNA responses to trans -10, cis -12 CLA in HepG2 cells. Response LS Means P-value Control CLA Agonist CLA + A SEMExp Trt E x T ACO 0.156 0.227 0.181 0.227 0.0180.0001 0.06 0.99 HMG-R 0.273 0.365 0.354 0.391 0.0190.0001 0.01 0.30 Apo A-I 0.249 0.398 0.400 0.395 0.0130.0001 0.0001 0.0001 Table A-10. Effects of MK886 on mRNA responses to trans -10, cis -12 CLA in HepG2 cells. Response LS Means P-value Control CLA Inhibitor CLA + I SEM Exp Trt E x T ACO 0.260 0.266 0.252 0.273 0.017 0.002 0.84 0.26 HMG-R 0.483 0.529 0.487 0.534 0.013 0.0001 0.04 0.26 Apo A-I 0.539 0.449 0.515 0.471 0.022 0.30 0.07 0.43

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161 Table A-11. Effects of WY 14,643 on mRNA responses to trans -10, cis -12 CLA in H-4-II-E cells. Response LS Means P-value Control CLA AgonistCLA + ASEMExp Trt E x T ACO 0.076 0.113 0.101 0.089 0.0060.85 0.01 0.99 HMG-R 0.051 0.058 0.050 0.050 0.0040.007 0.45 0.15 Apo A-I 0.69 0.56 0.67 0.66 0.0040.0006 0.17 0.12 Table A-12. Effects of MK886 on mRNA responses to trans -10, cis -12 CLA in H-4-II-E cells. Response LS Means P-value Control CLA Inhibitor CLA + I SEM Exp Trt E x T ACO 0.119 0.160 0.136 0.175 0.010 0.003 0.02 0.93 HMG-R 0.130 0.124 0.105 0.110 0.006 0.0002 0.06 0.09 Apo A-I 0.061 0.067 0.061 0.050 0.003 0.0001 0.01 0.01

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162 Table A-13. Effects of cis and trans isomers of octadecenoic acid on lipid-metabolizing genes and HDL cholesterol production in HepG2 cells. Response LS Means P-value Control c9 t9 c11 t11 SEM Exp Trt E x T ACO 0.253 0.258 0.259 0.327 0.275 0.012 0.0001 0.007 0.14 HMG-R 0.168 0.159 0.143 0.194 0.179 0.024 0.0004 0.64 0.28 Apo A-I 0.424 0.431 0.382 0.496 0.401 0.029 0.001 0.13 0.40 HDL, mg/dL 2.29 2.45 2.30 2.16 2.67 0.21 0.002 0.37 0.15 Table A-14. Effects of cis and trans isomers of octadecenoic acid on lipid-metabolizing genes and HDL cholesterol production in H-4-II-E cells. Response LS Means P-value Control c9 t9 c11 t11 SEM Exp Trt E x T ACO 0.256 0.298 0.264 0.276 0.307 0.009 0.0003 0.01 0.004 HMG-R 0.097 0.092 0.096 0.088 0.111 0.003 0.74 0.005 0.38 Apo A-I 0.113 0.100 0.102 0.103 0.107 0.002 0.88 0.02 0.09 HDL, mg/dL 1.50 1.04 1.86 0.98 1.50 0.33 0.42 0.26 0.99

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163 Table A-15. Effects of WY 14,643 on mRNA responses to cis -vaccenic acid in HepG2 cells. Response LS Means P-value Control c11 Agonistc11 + A SEMExp Trt E x T ACO 0.337 0.355 0.370 0.371 0.0110.0001 0.16 0.18 HMG-R 0.264 0.303 0.315 0.325 0.0160.0001 0.11 0.08 Apo A-I 0.239 0.214 0.266 0.320 0.0210.002 0.04 0.11 Table A-16. Effects of MK886 on mRNA responses to cis -vaccenic acid in HepG2 cells. Response LS Means P-value Control c11 Inhibitorc11 + I SEM Exp Trt E x T ACO 0.173 0.221 0.204 0.234 0.025 0.07 0.41 0.61 HMG-R 0.344 0.350 0.305 0.324 0.022 0.005 0.50 0.77 Apo A-I 0.377 0.383 0.375 0.401 0.021 0.005 0.83 0.78

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164 Table A-17. Effects of WY 14,643 on mRNA responses to trans -vaccenic acid in H-4-II-E cells. Response LS Means P-value Control t11 Agonistt11 + A SEMExp Trt E x T ACO 0.227 0.185 0.193 0.384 0.0280.82 0.003 0.55 HMG-R 0.053 0.085 0.082 0.154 0.0090.68 0.0003 0.44 Apo A-I 0.047 0.025 0.025 0.057 0.0050.40 0.004 0.66 Table A-18. Effects of MK886 on mRNA responses to trans -vaccenic acid in H-4-II-E cells. Response LS Means P-value Control t11 Inhibitort11 + I SEM Exp Trt E x T ACO 0.065 0.061 0.042 0.023 0.003 0.14 0.0001 0.97 HMG-R 0.067 0.093 0.076 0.085 0.009 0.89 0.25 0.56 Apo A-I 0.040 0.092 0.051 0.082 0.005 0.01 0.0001 0.002

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194 BIOGRAPHICAL SKETCH Elizabeth Sarah Greene was born in Turner sville, New Jersey in 1980. She is the daughter of Dr. David and Hilary Johnson. Sh e graduated with a Bachelor of Science degree in microbiology and cell science from the University of Florida in May 2002. After graduation, she began work on her Do ctor of Philosophy degree under the guidance of Dr. Lokenga Badinga in the Department of Animal Sciences, at the University of Florida. Her research focused on the effect s of fatty acids on lipid metabolism in human and rat liver. After graduating, Elizabeth pl ans to move to Texa s with her husband and pursue certification in Health Education.


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EFFECTS OF LONG-CHAIN FATTY ACIDS ON LIPID METABOLIZING GENES
AND HIGH-DENSITY LIPOPROTEIN CHOLESTEROL PRODUCTION IN
CULTURED HUMAN AND RAT HEPATOCYTES
















By

ELIZABETH SARAH GREENE


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA


2006







































Copyright 2006

by

Elizabeth Sarah Greene
































To my parents, Dave and Hilary Johnson, for instilling in me a love of learning and
supporting me through my seemingly never-ending quest for knowledge.















ACKNOWLEDGMENTS

I would like to thank my supervisory committee chair, Dr. Lokenga Badinga, for

allowing me the opportunity to be a part of his laboratory. I am grateful for his support

and advice in helping me become a better researcher. I would also like to thank my

committee members, Dr. Joel Brendemuhl, Dr. Bobbi Langkamp-Henken and Dr.

Charles Staples for their guidance and dedication to my education.

I would also like to thank the other members of the laboratory, Carlos

Rodriguez-Sallaberry and Cristina Caldari-Torres for many important problem-solving

and stress-relieving conversations during morning coffee breaks. Special thanks go to

Teri Woodham for being one of the best friends anyone could ever ask for. Additionally,

I could not have succeeded without all of the friends I made during the last four years.

Finally, I thank my husband, Nic, for his love, patience, strength, and never-ending faith

in my abilities. His belief in me made everything possible.
















TABLE OF CONTENTS

page

A C K N O W L E D G M E N T S ................................................................................................. iv

LIST OF TABLES ......... .... ................... ........ ............ ......viii

LIST OF FIGURES ................................... ...... ... ................. .x

ABBREVIATIONS KEY ................................................. ........ .. .............. xiv

ABSTRACT ........ .......................... .. ...... .......... .......... xvii

CHAPTER

1 INTRODUCTION ............... ................. ........... ................. ... ..... 1

2 LITER A TU RE REV IEW .................................................. ............................... 5

Structure and M etabolism of Lipids .................................... .......................... ......... 5
Structure and Nomenclature of Lipids ............... ............................................ 5
B iosynthesis of Fatty A cids............................................ ........................... 7
D egradation of Fatty Acids .............................. ............... ..................... 10
Nutritional and Biological Properties of the Polyunsaturated Fatty Acids................. 13
Dietary Requirements of the Essential Fatty Acids...........................................13
Long-Chain Polyunsaturated Fatty Acids of the n-6 Family..............................16
Long-Chain Polyunsaturated Fatty Acids of the n-3 Family..............................17
Digestion and Assimilation of Dietary Fats.................................... .................. 17
D ietary Fats in R elation to H health ................................................... ............... 24
Dietary Fats in Relation to Weight Control..................................24
Dietary Fats and Blood Cholesterol ....................................... ............... 26
Dietary Fats and Cardiovascular Disease........................ ..................28
C onjugated L inoleic A cid ...................... .............................................. 32
Roles of the Peroxisome Proliferator-Activated Receptors in Lipid Metabolism...... 38
P P A R a .............................................................................................. .40
P P A R /6 ...................................................................4 2
PPARy .................................................44









3 EFFECTS OF N-3 AND N-6 FATTY ACIDS ON LIPID METABOLIZING
GENES AND HIGH-DENSITY LIPOPROTEIN CHOLESTEROL
PRODUCTION IN CULTURED HUMAN AND RAT HEPATOCYTES ..............47

Introduction............ ... ...... ............................... ........................... 47
M materials and M methods ....................................................................... ..................4 8
M materials ......... .................................................................. ..... 48
Cell Culture and Treatm ent ........................................ ........................... 49
R N A Isolation and A nalysis........................................... .......... ............... 50
L ipid E xtraction ........... ... ............................................................ .... .... ... ....5 1
H D L C cholesterol A ssay ................................................ ............ ............... 51
Statistical A analysis .......................... ............ ...........................52
R results ................................................................................... ........ 52
Effects of Fatty Acids on HepG2 Cells ............................. ............... 52
Effects of Fatty A cids on H -4-II-E Cells.................................... ............... .... 53
Role of PPARa in Stearic Acid-Induced Effects on Gene Expression .............53
D discussion ......... ....... .......................................54
S u m m ary ............. ............. ................................................................5 8

4 EFFECTS OF ISOMERS OF CONJUGATED LINOLEIC ACID ON LIPID
METABOLIZING GENES AND HIGH-DENSITY LIPOPROTEIN
CHOLESTEROL PRODUCTION IN CULTURED HUMAN AND RAT
H E PA T O C Y TE S ......... .. ...... .... .. ........ .......................... .. ................ .. 80

Introduction ............... .... ............ ............ .. ......................... 80
M materials and M methods ....................................................................... ..................8 1
M materials .............. ......... ......... ........... ..................... .... 81
Cell Culture and Treatm ent ........................................ ........................... 82
RN A Isolation and A nalysis.......................................... .......................... 83
L ip id E x tra ctio n ............................................................................................. 8 4
H D L C cholesterol A ssay ............................................................ .....................84
Statistical A analysis ...................................... .............................84
R results ..................................................................5
Effects of Conjugated Linoleic Acid on HepG2 Cells .......................................85
Effects of Conjugated Linoleic Acid on H-4-II-E Cells...................................86
Role of PPARa in trans-10, cis-12 CLA-Induced Effects on Gene Expression .86
D isc u ssio n ...................... .. ............. .. ....................................................8 7
S u m m a ry ...................... .. ............. .. ..................................................... 9 1

5 EFFECTS OF CIS AND TRANS ISOMERS OF OCTADECENOIC ACID ON
LIPID METABOLIZING GENES AND HIGH-DENSITY LIPOPROTEIN
CHOLESTEROL PRODUCTION IN CULTURED HUMAN AND RAT
H E P A T O C Y T E S ............................................................................ .............. 114

Introdu action ................................................................................................ ..... 114
M materials and M methods ................................................................. .... .................. 115
M materials ..................................... .......................... ....................... 115









Cell Culture and Treatm ent ..................................................... ...... ......... 115
R N A Isolation and A nalysis.................................... .......................... .. ........ 116
L ipid Extraction .......................................... ........... ... .. ........ .. 117
H D L C cholesterol A ssay ................................................................ ............... 118
Statistical A analysis .................. ............................. ...... .. .. ............ 118
R results ................. ..... ................................ .....................119
Effects of Octadecenoic Acids on HepG2 Cells .......... ................................. 119
Effects of Octadecenoic Acids on H-4-II-E Cells ............................................ 119
Role of PPARa in Vaccenic Acid-Induced Effects on Gene Expression..........120
D isc u ssio n ........................................................................................12 1
S u m m a ry ......................................................................................................12 4

6 GENERAL DISCU SSION ......................................................... .............. 147

APPENDIX LS MEANS AND P-VALUES FOR ANALYSIS OF FATTY ACID
EFFECTS ON LIPID-METABOLIZING GENES AND HDL CHOLESTEROL
PRODUCTION IN HEPG2 and H-4-II-E CELLS .................................................156

L IST O F R E FE R E N C E S ......... ..................... ........................................ .........................165

BIOGRAPHICAL SKETCH ...... ........ ................... ............................ 194
















LIST OF TABLES


Table page

1-1 Biological functions of key genes studied................................................................4

A-i Effects of n-3 and n-6 FA on lipid-metabolizing genes and HDL cholesterol
production in HepG2 cells.......................................... ................... 156

A-2 Effects of n-3 and n-6 FA on lipid-metabolizing genes and HDL cholesterol
production in H -4-II-E cells. ...... .....................................................................156

A-3 Effects of WY 14,643 on mRNA responses to ST in HepG2 cells....................... 157

A-4 Effects of MK886 on mRNA responses to ST in HepG2 cells............................. 157

A-5 Effects of WY 14,643 on mRNA responses to ST in H-4-II-E cells. ....................158

A-6 Effects of MK886 on mRNA responses to ST in H-4-II-E cells. ..........................158

A-7 Effects of CLA on lipid-metabolizing genes and HDL cholesterol production in
H epG 2 cells. ........................................................................159

A-8 Effects of CLA on lipid-metabolizing genes and HDL cholesterol production in
H-4-II-E cells............................................. 159

A-9 Effects of WY 14,643 on mRNA responses to trans-10, cis-12 CLA in HepG2
c e lls .............................................................................1 6 0

A-10 Effects of MK886 on mRNA responses to trans-10, cis-12 CLA in HepG2 cells. 160

A-11 Effects of WY 14,643 on mRNA responses to trans-10, cis-12 CLA in H-4-II-E
c e lls .................................................................................. . 1 6 1

A-12 Effects of MK886 on mRNA responses to trans-10, cis-12 CLA in H-4-II-E
c e lls .................................................................................. . 1 6 1

A-13 Effects of cis and trans isomers of octadecenoic acid on lipid-metabolizing
genes and HDL cholesterol production in HepG2 cells............... ... ............... 162

A-14 Effects of cis and trans isomers of octadecenoic acid on lipid-metabolizing
genes and HDL cholesterol production in H-4-II-E cells. .....................................162









A-15 Effects of WY 14,643 on mRNA responses to cis-vaccenic acid in HepG2 cells. 163

A-16 Effects of MK886 on mRNA responses to cis-vaccenic acid in HepG2 cells.......163

A-17 Effects of WY 14,643 on mRNA responses to trans-vaccenic acid in H-4-II-E
cells .................................................... 164

A-18 Effects of MK886 on mRNA responses to trans-vaccenic acid in H-4-II-E cells. 164
















LIST OF FIGURES


Figure pae

3-1 Effect of long-chain FA on ACO mRNA expression in HepG2 cells......................59

3-2 Effect of long-chain FA on HMG-R mRNA expression in HepG2 cells................60

3-3 Effects of long-chain FA on Apo A-I mRNA expression in HepG2 cells...............61

3-4 Effects of long-chain FA on HDL cholesterol production in HepG2 cells..............62

3-5 Effects of long-chain FA on ACO mRNA expression in H-4-II-E cells..................63

3-6 Effects of long-chain FA on HMG-R mRNA expression in H-4-II-E cells.............64

3-7 Effects of long-chain FA on Apo A-I mRNA expression in H-4-II-E cells.............65

3-8 Effects of long-chain FA on HDL cholesterol production in H-4-II-E cells............66

3-9 Effect of WY 14,643 on ACO mRNA response to ST in HepG2 cells .................67

3-10 Effect of WY 14,643 on HMG-R mRNA response to ST in HepG2 cells ..............68

3-11 Effect of WY 14,643 on Apo A-I mRNA response to ST in HepG2 cells ..............69

3-12 Effect of MK886 on ACO mRNA response to ST in HepG2 cells..........................70

3-13 Effect of MK886 on HMG-R mRNA response to ST in HepG2 cells.....................71

3-14 Effect of MK886 on Apo A-I mRNA response to ST in HepG2 cells...................72

3-15 Effect of WY14,643 on ACO mRNA response to ST in H-4-II-E cells ................73

3-16 Effect of WY 14,643 on HMG-R mRNA response to ST in H-4-II-E cells............74

3-17 Effect of WY 14,643 on Apo A-I mRNA response to ST in H-4-II-E cells............75

3-18 Effect of MK886 on ACO mRNA response to ST in H-4-II-E cells .....................76

3-19 Effect of MK886 on HMG-R mRNA response to ST in H-4-II-E cells ..................77

3-20 Effect of MK886 on Apo A-I mRNA response to ST in H-4-II-E cells ..................78









3-21 Regulation of lipid metabolizing genes and HDL cholesterol production by
long-chain fatty acids ............................................... ...............79

4-1 Effect of CLA on ACO mRNA expression in HepG2 cells...............................93

4-2 Effect of CLA on HMG-R mRNA expression in HepG2 cells..............................94

4-3 Effect of CLA on Apo A-I mRNA expression in HepG2 cells..............................95

4-4 Effect of CLA on HDL cholesterol production by HepG2 cells..............................96

4-5 Effect of CLA acid on ACO mRNA expression in H-4-II-E cells...........................97

4-6 Effect of CLA on HMG-R mRNA expression in H-4-II-E cells ...........................98

4-7 Effect of CLA on Apo A-I mRNA expression in H-4-II-E cells ...........................99

4-8 Effect of CLA on HDL cholesterol production by H-4-II-E cells ....................... 100

4-9 Effect of WY 14,643 on ACO mRNA response to trans-10, cis-12 CLA in
HepG2 cells .................. ......... ................... ....................... 101

4-10 Effect of WY 14,643 on HMG-R mRNA response to trans-10, cis-12 CLA in
HepG2 cells ..... ....................... ................... 102

4-11 Effect of WY 14,643 on Apo A-I mRNA response to trans-10, cis-12 CLA in
HepG2 cells ................ ........ .................. 103

4-12 Effect of MK886 on ACO mRNA response to trans-10, cis-12 CLA in HepG2
c e lls .............................................................................1 0 4

4-13 Effect of MK886 on HMG-R mRNA response to trans-10, cis-12 CLA in
HepG2 cells ................ ........ .................. 105

4-14 Effect of MK886 on Apo A-I mRNA response to trans-10, cis-12 CLA in
HepG2 cells ................... .. ......... ................... 106

4-15 Effect of WY 14,643 on ACO mRNA response to trans-10, cis-12 CLA in
H-4-II-E cells............................................. 107

4-16 Effect of WY 14,643 on HMG-R mRNA response to trans-10, cis-12 CLA in
H -4 -II-E c e lls .................................................................................................... 1 0 8

4-17 Effect of WY 14,643 on Apo A-I mRNA response to trans-10, cis-12 CLA in
H -4 -II-E c e lls .................................................................................................... 1 0 9

4-18 Effect of MK886 on ACO mRNA response to trans-10, cis-12 CLA in H-4-II-E
c e lls .................................................................................. . 1 1 0









4-19 Effect of MK886 on HMG-R mRNA response to trans-10, cis-12 CLA in
H -4 -II-E c e lls .................................................................................................... 1 1 1

4-20 Effect of MK886 on Apo A-I mRNA response to trans-10, cis-12 CLA in
H-4-II-E cells ..................................................................... ......... 112

4-21 Regulation of lipid metabolizing genes and HDL cholesterol production by
C L A ............................................................................................... . 1 1 3

5-1 Effect of cis and trans isomers of octadecenoic acid on ACO mRNA expression
in H ep G 2 cells ................................................. ................ 12 6

5-2 Effect of cis and trans isomers of octadecenoic acid on HMG-R mRNA
expression in HepG2 cells.......................................... ................... 127

5-3 Effect of cis and trans isomers of octadecenoic acid on Apo A-I mRNA
expression in HepG2 cells.......................................... ................... 128

5-4 Effects of cis and trans isomers of octadecenoic acid on HDL cholesterol
production by HepG2 cells................................. .......... .................. 129

5-5 Effect of cis and trans isomers of octadecenoic acid on ACO mRNA expression
in H -4-II-E cells .....................................................................130

5-6 Effect of cis and trans isomers of octadecenoic acid on HMG-R mRNA
expression in H -4-II-E cells .............................................................................131

5-7 Effect of cis and trans isomers of octadecenoic acid on Apo A-I mRNA
expression in H -4-II-E cells .............................................................................132

5-8 Effects of cis and trans isomers of octadecenoic acid on HDL cholesterol
production in H -4-II-E cells ............................................................................133

5-9 Effect of WY 14,643 on ACO mRNA response to cis-vaccenic acid in HepG2
c e lls .................................................................................. . 1 3 4

5-10 Effect of WY 14,643 on HMG-R mRNA response to cis-vaccenic acid in
H ep G 2 cells ................................................ .............. . ............... 13 5

5-11 Effect of WY 14,643 on Apo A-I mRNA response to cis-vaccenic acid in
H ep G 2 cells ............................................................. .............. .... 13 6

5-12 Effect of MK886 on ACO mRNA response to cis-vaccenic acid in HepG2
c e lls .................................................................................. . 1 3 7

5-13 Effect of MK886 on HMG-R mRNA response to cis-vaccenic acid in HepG2
c e lls .......................................................................... 1 3 8









5-14 Effect of MK886 on Apo A-I mRNA response to cis-vaccenic acid in HepG2
c e lls .............................................................................1 3 9

5-15 Effect of WY 14,643 on ACO mRNA response to trans-vaccenic acid in
H -4 -II-E c e lls .................................................................................................... 14 0

5-16 Effect of WY 14,643 on HMG-R mRNA response to trans-vaccenic acid in
H -4 -II-E c e lls .................................................................................................... 14 1

5-17 Effect of WY 14,643 on Apo A-I mRNA response to trans-vaccenic acid in
H -4 -II-E c e lls .................................................................................................... 14 2

5-18 Effect of MK886 on ACO mRNA response to trans-vaccenic acid in H-4-II-E
c e lls .............................................................................1 4 3

5-19 Effect of MK886 on HMG-R mRNA response to trans-vaccenic acid in
H -4 -II-E c e lls .................................................................................................... 14 4

5-20 Effect of MK886 on Apo A-I mRNA response to trans-vaccenic acid in
H -4 -II-E c e lls .................................................................................................... 14 5

5-21 Regulation of lipid metabolizing genes and HDL cholesterol production by cis
and trans octadecenoic fatty acids ........................................ ...... ............... 146


















AA

ABCA1


ACC

ACO

ACP

AI

Apo

BMI

CoA

CHD

CLA

CM

CPT-I or -II

CVD

DGAT

DHA

EPA

ER

ETF


ABBREVIATIONS KEY

arachidonic acid

adenosine triphosphate-binding cassette
transporter-Al

acetyl-CoA carboxylase

acyl-CoA oxidase

acyl carrier protein

adequate intake

apolipoprotein

body mass index

coenzyme A

coronary heart disease

conjugated linoleic acid

chylomicron

carnitine-palmitoyl transferase I or -II

cardiovascular disease

diacylglygerol acyltransferase

docosahexaenoic acid

eicosapentaenoic acid

endoplasmic reticulum

electron transfer flavoprotein









FA fatty acid

FABPc cytosolic fatty acid binding protein

FAS fatty acid synthase

GLA gamma-linolenic acid

HDL high-density lipoprotein

HMG-R 3-hydroxy, 3-methylglutaryl CoA reductase

HODE hydroxyoctadecadienoic acid

IUPAC International Union of Pure and Applied
Chemistry

LA linoleic acid

LCAT lecithin:cholesterol acyltransferase

LNA linolenic acid

LPL lipoprotein lipase

MGAT monoacylglycerol acyltransferase

MTP microsomal transfer protein

MUFA monounsaturated fatty acid

NCEP National Cholesterol Education Program

NEFA non-esterified fatty acid

PPAR peroxisome proliferator-activated receptor

PPRE peroxisome proliferator response element

PUFA polyunsaturated fatty acid

RXR retinoid X receptor

SCD stearyl-CoA desaturase

ST stearic acid









TAG triacylglycerol

TPN total parenteral nutrition

TZD thiazolidinediones

VLDL very low-density lipoprotein















Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy

EFFECTS OF LONG-CHAIN FATTY ACIDS ON LIPID METABOLIZING GENES
AND HIGH-DENSITY LIPOPROTEIN CHOLESTEROL PRODUCTION IN
CULTURED HUMAN AND RAT HEPATOCYTES

By

Elizabeth Sarah Greene

May 2006

Chair: Lokenga Badinga
Major Department: Animal Sciences

A series of experiments were conducted to examine the short-term effects of

long-chain fatty acids (FA) on acyl CoA oxidase (ACO), 3-hydroxy, 3-methylglutaryl

CoA reductase (HMG-R) and apolipoprotein A-I (Apo A-I) gene expression, and

high-density lipoprotein cholesterol (HDL-C) production in HepG2 (human) and

H-4-II-E (rat) hepatocytes. In the three experiments, the FA studied were 1) FA of

differing saturation and chain length, 2) conjugated linoleic acid (CLA), and 3) cis (c9,

cl 1) and trans (t9, tl 1) isomers of octadecenoic acid.

In HepG2 cells, ACO mRNA was up-regulated by trans-10, cis-12 CLA and

cis-vaccenic acid (cl 1). HMG-R gene expression was increased by stearic acid (ST) and

trans-10, cis-12 CLA. Steady-state levels of Apo A-I mRNA were increased by all FA in

the first experiment, trans-10, cis-12 CLA, and cl1. HDL-C was decreased only by

cis-9, trans-11 CLA. In H-4-II-E cells, ACO mRNA was up-regulated by LA, CLA, ST,









oleic acid, and trans-vaccenic acid (tl 1). HMG-R gene expression was increased by ST,

CLA isomers, and tl 1. Apo A-I was increased by ST and EPA, but decreased by CLA

and cis and trans monounsaturated FA. HDL-C was increased by LNA in the first

experiment.

Based on these findings, we investigated the possibility that the FA effects are

mediated by peroxisome proliferator-activated receptor a (PPARa). In HepG2 cells,

activation or inhibition of PPARa had minimal effects on basal or FA-effects on gene

expression, consistent with the low-levels of endogenous PPARa in this cell line. In

H-4-II-E cells, activation of PPARa increased the abundance of basal ACO mRNA,

enhanced the effect of ST on ACO and Apo A-I mRNA, and enhanced the effects of t 1

on ACO, HMG-R, and Apo A-I gene expression. Inhibition of PPARa decreased basal

expression of ACO and attenuated the effects of ST and tl 1 on ACO and effects of

trans-10, cis-12 CLA on Apo A-I gene expression. These results indicate that specific

FAs may regulate lipid-metabolizing genes in the liver through a PPARa-dependent

mechanism. Because of different responses to FA in human and rat cell lines, however,

net effects are likely species specific.


xviii














CHAPTER 1
INTRODUCTION

Dietary fat is an important nutrient for the function and survival of all organisms.

Historically, body lipids have been considered primarily to serve as an energy source, as

constituents of cell membranes, and as precursors for molecules involved in signal

transduction, such as steroids and prostaglandins. More recently however, fatty acids

(FA) have been shown to affect gene expression, leading to changes in cell

differentiation, growth, and metabolism (Clarke and Jump, 1994; Jump et al., 1996).

Additionally, dietary fat has been implicated in the progression of several chronic

diseases, including type II diabetes, cardiovascular disease, and some types of cancer

(Sanders, 2003), though the effects may depend on the composition of dietary fat

consumed. Therefore, understanding the molecular basis for FA effects on gene

regulation is necessary for further elucidation of the role of fats in human health. To

address this issue, our studies focused on the effects of three general classes of FA that

may play a significant role in health and metabolism: n-3 and n-6 long-chain

polyunsaturated fatty acids, conjugated linoleic acids (CLA), and cis and trans isomers of

fatty acids.

Dietary polyunsaturated fatty acids (PUFA) have been reported to lower blood

triglycerides, alter the blood lipid profile, decrease intramuscular lipid droplet size,

improve insulin sensitivity, and enhance glucose utilization (Jump and Clarke, 1999).

Since the seminal observation that PUFAs could inhibit hepatic lipogenesis in mice

(Allmann and Gibson, 1965), numerous studies have demonstrated that diets rich in









PUFAs influence metabolic changes by coordinately suppressing lipid synthesis in the

liver and enhancing fatty acid oxidation in both liver and skeletal muscle (Jump and

Clarke, 1999). The PUFA induction of genes encoding proteins involved in lipid

oxidation include 3-hydroxy, 3-methylglutaryl-CoA synthase (Rodriguez et al., 1994),

carnitine palmitoyltransferase, fatty acid binding proteins, and peroxisomal acyl-CoA

oxidase (ACO; Reddy and Hashimoto, 2001).

Conjugated linoleic acid (CLA) is a collective term for positional and geometric

isomers of linoleic acid (LA). Though over 16 individual isomers have been identified

(Rickert et al., 1999), only cis-9, trans-11 CLA and trans-10, cis-12 CLA are known to

possess biological activity (Pariza et al., 2000). Cis-9, trans-11 CLA is the predominant

CLA produced as an intermediate in the rumen during biohydrogenation of dietary LA

and is commonly found in dairy products and ruminant meat. Dietary sources of

trans-10, cis-12 CLA derive predominantly from synthetic partial hydrogenation and are

found in margarines, shortenings, and supplements (Gaullier et al., 2002). First identified

in grilled beef as a potential anti-carcinogen (Pariza and Hargraves, 1985), numerous

health benefits have been attributed to CLA mixtures, including actions as an

antiadipogenic (Park et al, 1997), antidiabetogenic (Houseknecht et al., 1998), and

antiatherosclerotic (Kritchevsky et al., 2004) agent. More recently, studies involving

individual isomers have shown that the two main isoforms can have different effects on

metabolism and cell function and may act through different signaling pathways (Wahle et

al., 2004). Metabolic responses to cis-9, trans-11 and trans-10, cis-12 CLA may differ,

but both isomers have implications for human health. Most studies have been performed

in animal models, with species differences observed. In particular, only some of the









findings attributed to animal models pertain to human subjects, and even when

comparing studies in humans, results are often inconclusive (Terpstra, 2004).

Trans-fatty acids are geometrical isomers of unsaturated FA that assume a saturated

fat-like configuration that differs from that of their cis counterparts. The predominant

source of trans fats in the human diet is hydrogenated oils (such as margarine and

partially hydrogenated soybean oil) commonly found in baked goods and deep fat-fried

fast foods (Hu et al., 2001). Metabolic studies in several species have shown that

trans-FA can negatively alter the lipid profile to a greater extent than saturated fats,

because trans-FA not only increase small, dense LDL cholesterol (Mauger et al., 2003),

but also decrease HDL cholesterol in some studies (Judd et al., 1994; de Roos et al.,

2003). Additionally, epidemiological evidence associates trans-FA intake with increased

risk for cardiovascular disease (Ascherio et al., 1999). Few studies, however, have

examined the role of individual trans-FA in modulating lipid metabolism. As with other

FA, it is possible that cis and trans isomers of octadecenoic acid may also have

differential effects on lipid metabolism.

Based on both dietary and in vitro studies of lipid metabolism, we hypothesized

that various FA of differing degree of saturation and double-bond position will have

differing effects on ACO, 3-hydroxy, 3-methylglutaryl CoA reductase (HMG-R), and

apolipoprotein A-I (Apo A-I) gene expression, as well as HDL cholesterol production in

HepG2 and H-4-II-E hepatoma cells (Table 1-1). Also, because several FA and their

derivatives are known ligands for peroxisome proliferator-activated receptors (PPAR;

Schoonjans et al., 1996), we hypothesized that these FA may act on lipid-metabolizing

genes through activation of PPARa, the predominant receptor subtype in the liver









(Braissant et al., 1996). If this hypothesis is correct, activation of PPARa should mimic

the effects of FA, whereas inhibition of PPARa would be expected to block FA effects in

HepG2 and H-4-II-E hepatoma cell lines. The overall aim of our studies was to examine

the differential roles of fatty acids on lipid metabolizing genes involved in peroxisomal

P-oxidation and cholesterol synthesis in human and rat hepatoma cell lines.

Table 1-1. Biological functions of key genes studied


Gene Function

ACO Rate limiting in peroxisomal P-oxidation

HMG-R Rate limiting in cholesterol synthesis; converts HMG-CoA to
mevlonate

Apo A-I Necessary for proper packaging of HDL cholesterol














CHAPTER 2
LITERATURE REVIEW

Structure and Metabolism of Lipids

Structure and Nomenclature of Lipids

Based on physical properties, the term lipid denotes a heterogeneous group of

substances that are insoluble in water, but are soluble in non-polar solvents such as

chloroform and alcohols (Smith, 2000). This definition covers a wide range of

molecules, including FA, phospholipids, sterols, sphingolipids, terpenes, and others

(Christie, 2003). Fatty acids consist of a chain of two or more carbon atoms, with a

methyl group at one end, and a carboxyl group at the other end of the chain. The main

structural features are their chain length, degree of unsaturation (number of double

bonds), and presence of substituent groups. Additionally, the presence of double bonds

allows for positional and geometric isomerism. Positional isomers occur when double

bonds are located at different positions along the carbon chain. The position of

unsaturation is numbered in reference to the first of the pair of carbon atoms between

which the double bond occurs. Geometric isomerism refers to the configuration of the

hydrogen atoms in respect to the double bond. If the hydrogen atoms are on the same

side of the molecule opposite the double bond, it is said to be in the cis configuration.

Alternately, if the hydrogen atoms are on opposite sides, the configuration is trans. Most

naturally occurring unsaturated FA are in the cis configuration, but natural and synthetic

trans isomers do exist.









The naming scheme for FA must be able to clearly define a lipid structure in a

manner that is amenable to scientists and researchers of all fields. Several systems are

currently used, though to different degrees. First, there are the trivial names, such as

stearic acid and linoleic acid, which were assigned as the individual FA were discovered.

Although these may be used for the most-commonly occurring FA, naming and

remembering unusual unsaturated, branched, or hydroxyl-FA becomes unwieldy.

Because of this difficulty, two different systems have been developed. The older system

used Greek letters to identify carbon atoms, beginning at the carboxyl end. Considering

the carboxyl carbon as Cl, C2 is called the a-carbon, C3 the 0-carbon, and so on, ending

with the co-carbon at the methyl end. Though this system is no longer preferred, it is used

to name the co-3 and co-6 FAs, in which the last double bond in the chain occurs three and

six carbons from the co-carbon, respectively. In much of the newer literature, the co is

often replaced by an n, but the meaning remains the same.

Currently, the preferred system for specifying individual FA is the numbering

system standardized by the International Union of Pure and Applied Chemistry (IUPAC)

(IUPAC-IUB, 1977). For linoleic acid, an 18-carbon FA with two cis double bonds in

positions 9 and 12 from the carboxyl end, the systematic name is cis-9, cis-12

octadecadienoic acid. In the shorthand system, FA are identified by two numbers

separated by a colon; the first number indicates the number of carbon atoms, the second

indicates the number of double bonds in the structure. For example, a saturated fat such

as stearic acid would be 18:0, whereas a polyunsaturated fat, such as linoleic acid would

be represented by cis-9, cis-12 18:2.









Biosynthesis of Fatty Acids

Most naturally occurring FA contain an even number of carbon atoms, leading

early researchers to speculate that they were formed by the condensation of two-carbon

units. This was confirmed using rats fed acetic acid labeled with 13C in the carboxyl

group and 2H in the methyl group. When FA were isolated from the rat tissues, the

labeled carbons were found in alternate positions along the chain, showing that the

complete FA could be derived from acetic acid (Rittenberg and Bloch, 1944). When the

details of P-oxidation were elucidated in the 1950s, it led to speculation that FA synthesis

could be the simple reversal FA breakdown. However, several discoveries soon showed

that the pathways were distinctly different. First, NADPH (not NAD+ as in oxidation)

serves as a cofactor. Second, there is a requirement for bicarbonate (Wakil, 1962; Brady

and Gurin, 1952).

Fatty acid synthesis can be broken into two basic processes: condensation of two

carbon units to form 16 tol8-carbon FA and various modifications of these products. In

mammals, the majority of carbon for de novo FA synthesis comes from pyruvate, the

end-product of glycolysis. To be used in FA synthesis, acetyl coenzyme A (CoA; the

activated form of acetic acid) must be generated from pyruvate. To accomplish this, the

pyruvate is transported from the cytosol into the mitochondria, where the enzyme

pyruvate dehydrogenase acts to produce acetyl-CoA. Acetyl-CoA and oxaloacetate

combine to form citrate, which can then be transported back out of the mitochondria via a

tricarboxylate anion carrier, where the cycle is completed, and acetyl-CoA is produced by

the action of ATP:citrate lyase.









The first and rate-limiting reaction in de novo FA synthesis is catalyzed by

acetyl-CoA carboxylase (ACC). In this enzymatic reaction, acetyl-CoA is carboxylated,

leading to the formation of malonyl-CoA (Knowles, 1989). This reaction requires biotin

as a cofactor, as shown by inhibition of carboxylation by avidin, a potent inhibitor of

biotin (Wakil et al., 1958). Acetyl-CoA carboxylase is activated by phosphorylation and

deactivated by dephosphorylation (Shacter et al., 1986). The malonyl-CoA generated by

ACC forms the source of nearly all carbons of the FA. Only the first two carbons arise

from the "primer molecule," acetyl-CoA. In order for individual malonyl-CoA units to

join into the FA chain, they must be attached to the acyl carrier protein (ACP). The ACP

is a small molecular mass protein (8.8 kDa) that is very stable over a range of pH and

temperature values (D'Agnolo et al., 1975).

The enzymatic steps involved in adding successive malonyl-CoA units to the chain

are collectively known as fatty acid synthase (FAS). In animals, FAS is a multifunctional

enzyme, with discrete domains catalyzing the condensation, dehydration, and reduction

reactions. Animal FAS complexes consist of homodimers with molecular weight of

approximately 450-550 kDa (Smith, 1994). Essentially, to elongate the chain,

malonyl-ACP attached to one half of the dimer interacts with the growing acyl chain

attached to the active site of the condensing enzyme on the other half of the dimer (Joshi

et al., 1998). The typical end product of FAS is palmitic acid (16:0). The thioesterase

actions of FAS cleave the product from the enzyme. This specificity for a 16-carbon

product is likely due to stearic hindrance of the condensing domain by the large FA

(Chirala and Wakil, 2004). Although the production of palmitate is most common,

different organisms and tissues can produce FA of shorter chain lengths as necessary.









For example, in the rat mammary gland, where large amounts of 8:0 and 10:0 are

necessary for the formation of milk triacylglycerols (TAG), a second thioesterase is

present, forming medium-chain FA (Smith, 1994).

Although the main product of FAS is palmitate, many tissues contain longer chain

FA, particularly as a component of membrane lipids. The formation of long and very

long-chain FA is catalyzed by Type III synthases, often termed elongases due to their

lengthening of pre-formed and dietary FA. In mammalian tissues, two separate

elongation systems are located in the mitochondria and endoplasmic reticulum (ER). In

the mitochondria, two carbon units in the form of acetyl-CoA (not malonyl-CoA as in de

novo synthesis) are added preferentially to monoenoic over saturated substrates (Moon et

al., 2001). Mitochondrial elongation is essentially a reversal of P-oxidation, with a

requirement for NADPH and NADH (Seubert and Podack, 1973). Formation of the

longer chain FA occurs at the ER. In this case, malonyl-CoA serves as the two carbon

donor and NADPH is the reducing agent. This system can produce FA with chain

lengths in excess of 20 carbons (Suneja et al., 1990).

Once saturated FA have been produced by the organism, they can be used to

produce unsaturated FA, mainly by the process of oxidative desaturation. In this

mechanism, a double bond is introduced directly into a pre-formed saturated long-chain

FA, using 02 and a reducing compound (NADH) as cofactors (Scheuerbrandt and Bloch,

1962). Mammalian enzymes normally introduce new double bonds between an existing

double bond and the carboxyl group, whereas plant enzymes introduce the new bond

between an existing double bond and the terminal methyl group. There are three

components to the desaturation complex: NADH-cytochrome b5 reductase, cytochrome









bs, and the desaturase enzyme (Stritmatter et al., 1974). Most of what is known about

desaturases is derived from early studies showing that A9 desaturase is the rate-limiting

step in the conversion of stearic acid (18:0) to oleic acid (18:1, n-9) (Schroepfer and

Bloch, 1965). Because of its actions, it is also referred to as stearyl-CoA desaturase

(SCD). Mammals contain desaturases able to introduce double bonds in the A5, A6, and

A9 positions. Plants additionally possess the A12 and A15 desaturases necessary for the

formation of n-6 and n-3 FA.

Degradation of Fatty Acids

In the body, FA from dietary or stored TAG are broken down to provide a source of

energy. The main forms of FA oxidation are termed alpha (a), beta (0), and omega (co),

depending on the carbon in the chain that is attacked. Of the three types of oxidation,

P-oxidation is the most prevalent.

The basic mechanism for P-oxidation was originally proposed by Knoop in 1904

after feeding labeled FA to dogs, and was confirmed by Dakin's isolation of the proposed

intermediates in 1912. Fats degraded in this manner liberate two-carbon units in the form

of acetyl-CoA through the introduction of a double bond between the 0- and y-carbons,

hence the name P-oxidation. Until relatively recently, mitochondria were considered the

only cellular site for P-oxidation. Although all the necessary enzymes are present in

mitochondria, the microbodies (peroxisomes in mammals and glyoxysomes in plants) can

also complete the process (Lazarow and de Duve, 1976). The contribution of

microbodies to total P-oxidation depends on the organism and specific tissue considered.

For example, in mammals, peroxisomal P-oxidation of very long-chain FAs is









particularly important in the liver and kidneys, with defects leading to devastating

diseases (Fournier et al., 1994).

Fatty acyl-CoAs formed within the cytosol cannot enter the mitochondrion directly,

providing a major point of control and regulation of FA metabolism (Eaton, 2002). The

observation that carnitine stimulates the P-oxidation of long-chain FA gave the first clue

to its function in the mitochondrial uptake of FA (Bremer, 1962; Fritz and Yue, 1963).

Acyl residues are transferred to camitine by carnitine-palmitoyl transferase (CPT-I) at the

surface of the outer mitochondrial membrane. This allows the FA to transverse the

membrane via porin, where they are then transported through the inner mitochondrial

membrane by a carnitine:acylcarnitine translocase (Pande, 1975). The translocase causes

a one-to-one exchange of carnitine for acylcamitine, ensuring a constant level of carnitine

within the mitochondria (Ramsay and Tubbs, 1975). Once within the mitochondrial

matrix, a second carnitine-palmitoyl transferase, CPT-II, acts to transfer the acyl group

from camitine back to CoA, reforming acyl-CoAs, the substrate for P-oxidation (Bieber,

1988).

The reactions of P-oxidation involve four enzymes in repeated sequence, resulting

in the cleavage of two carbons at a time from the acyl chain. Acyl-CoA dehydrogenase

acts to produce trans-2,3-enoyl-CoA. This step is linked to the respiratory chain via

electron transfer flavoprotein (ETF) and ETF-ubiquinone oxireductase (Parker and Engel,

2000). The 2-enoyl-CoA hydratase then acts on the product of the first reaction,

producing 3-hydroxyacyl-CoA. The next enzyme in the sequence, 3-hydroxyacyl-CoA

dehydrogenase, is linked with NAD+ and produces 3-oxoacyl-CoA. Finally,

3-oxoacyl-CoA thiolase actions produce a saturated acyl-CoA that has been shortened by









two carbons, in the form of acyl-CoA (Eaton et al., 1996). Each of the enzymes is

present in several isoforms with varying chain-length specificities, primarily for short,

medium, long, and very long-chain acyl-CoA. This allows for improved efficiency of

P-oxidation and prevents buildup of intermediates that could lead to inhibition (Bartlett

and Eaton, 2004).

Peroxisomal P-oxidation is important in almost all eukaryotic organisms (Kunau et

al., 1995). The peroxisomal and mitochondrial enzymes of P-oxidation differ in several

ways. Peroxisomes do not have an electron transport system coupled to energy

production as can be found in mitochondria. The first and rate limiting step in

peroxisomal P-oxidation is catalyzed by acyl-CoA oxidase (ACO), which introduces a

trans-2 double bond and produces hydrogen peroxide. Next, a trifunctional enzyme

produces 0-ketoacyl-CoA, which is acted upon by a thiolase, producing acetyl-CoA and a

shortened acyl-CoA (Mannaerts et al., 2000). Due to limited substrate specificities for

ACO, peroxisomes are incapable of oxidizing long-chain FA completely (Singh et al.,

1984). Medium chain products of peroxisomal P-oxidation are transferred to carnitine,

allowing them to be transported into the mitochondria for complete oxidation. Defects in

peroxisomal P-oxidation can lead to the accumulation of very long-chain FA in various

tissues, producing devastating diseases such as Zellweger syndrome and

adrenoleukodystrophy (Kunau et al., 1995).

The above enzymatic cycle assumes that the substrate is a straight-chain, saturated

FA with an even number of carbons. For FA of odd-chain lengths, P-oxidation yields

propionyl-CoA in addition to acetyl-CoA; therefore, the ability of an organism or tissue

to oxidize these FA depends on the ability of that organism or tissue to metabolize









propionate. The liver is well-equipped to oxidize propionate and oxidizes odd-chain FA

well, whereas the heart cannot oxidize the product, and P-oxidation of odd-chain FA

stops with an increase in propionate (Grynberg and Demaison, 1996). Beta-oxidation of

unsaturated FA poses several problems. Most naturally occurring unsaturated FA contain

cis double bonds and the bonds may be in the wrong position along the chain for effective

P-oxidation. Unsaturated FA with odd-numbered double bonds, such as the 9-cis bond of

LA, are shortened to 3-cis-enoyl-CoA and then isomerized to 2-trans-enoyl CoA that can

be further degraded via P-oxidation (Stoffel and Caesar, 1965). Fatty acids with

even-numbered double bonds are shortened to 4-cis-enoyl-CoA, which are then

dehydrogenated to 2-trans, 4-cis-dienoyl-CoA. One double bond is then removed by

NADPH-dependent 2,4-dienoyl-reductase, allowing P-oxidation to continue (Kunau and

Dommes, 1978).

The acyl-CoA produced by chain-shortening can have several different fates,

depending on the tissue. In ketogenic tissues such as the liver, acetyl-CoA is used to

form the ketone bodies, acetoacetate and P-hydroxybutyrate, for export and peripheral

oxidation. In most tissues, however, acetyl-CoA enters the Krebs cycle and generates

energy in the form of ATP (Hiltunen and Qin, 2000).

Nutritional and Biological Properties of the Polyunsaturated Fatty Acids

Dietary Requirements of the Essential Fatty Acids

Mammalian cells can synthesize saturated and omega-9 (n-9) unsaturated FA de

novo from acetyl-CoA, but lack the A 12 and A 15 desaturase enzymes necessary for the

formation of double bonds in the omega-6 (n-6) and omega-3 (n-3) positions. Because of

this enzyme deficiency, the linoleic acid (LA; 18:2, n-6) and a-linolenic acid (LNA; 18:3,

n-3) are considered essential nutrients in the human diet (Innis, 1991). Once ingested,









LA and LNA can be further elongated and desaturated into biologically important

long-chain polyunsaturated fatty acids (PUFA) of 20 or more carbons and three to six

double bonds.

Determining the essential requirements of a nutrient generally begins with

recognition of a deficiency, continues with the study of intakes that can prevent or

reverse the deficiency, and finally concludes with the definition of a range of intakes for

optimal biological function. In 1929, Burr and Burr discovered that rats fed a fat-free

diet developed dermatitis and grew at a slower rate than their fat-fed counterparts. These

deficiencies could only be eliminated by adding certain FAs to the diet, which were later

determined to be LA and arachidonic acid (AA; 20:4). This knowledge was applied to

produce essential FA deficiency in a variety of species, including man. In all species, the

deficiency is characterized by skin symptoms, such as dermatosis or eczema, retarded

growth, impaired reproduction, and degeneration or impairment of function in many

bodily organs, including the heart and kidneys (Sinclair, 1990). These signs are

characterized by changes in the FA composition of many tissues, particularly in

biological membranes and mitochondria.

Well-documented essential FA deficiency in man is rare, but was first seen in the

1940s and 50s in infants receiving formula containing skim milk and sugar as a substitute

for mother's milk. When fed formula containing increasing concentrations of LA,

clinical signs of deficiency disappeared when concentrations in the diet were above 0.1%

of dietary energy (Hansen et al., 1958). Adult essential FA deficiency was most

commonly seen in patients receiving total parenteral nutrition (TPN), in which early

formulas were fat-free (Holman, 1981). In some cases, patients responded to the









application to the skin of fats with a high proportion of LA, showing that the FA do not

necessarily have to be absorbed through the gastrointestinal tract to be effective. More

frequently, LA deficiency may develop as a secondary condition to other disorders such

as severe malnutrition and fat malabsorption.

The n-3 FA can, in part, substitute for a deficiency in n-6 FA, but also have their

own distinct roles (Benatti et al., 2004). The understanding of n-3 FA essentiality lagged

significantly behind that of n-6 FA, partially because of their naturally lower amounts in

the body. The first case of n-3 FA deficiency was induced by an n-3 FA-free TPN

formula. Symptoms of n-3 FA deficiency in the patient included numbness, tingling,

weakness, inability to walk, leg pain, psychological disturbances and blurred vision. The

patient's plasma lipid profile showed the concentration of total n-3 FA to be at 34% of

the control value. When soybean oil, a source of LNA, was added back to the TPN

formula, the signs of deficiency disappeared (Holman et al., 1982).

As essential FA deficiency is usually associated with a disease state, there is little

evidence to determine dietary reference intakes for healthy populations. Therefore, based

on the data that are available on health effects of LA and LNA, adequate intake (AI)

levels have been recommended. The AI is a value based on experimentally derived

intake levels or approximate mean nutrient intakes by a group of healthy individuals.

Based on current estimates, PUFAs contribute approximately 5-6% of energy in the

Western diet (Grundy et al., 1982). For adults, it is recommended that consumption of

LA should be 17 g/d for men and 12 g/d for women. For LNA, recommended intakes are

1.6 g/d for men and 1.1 g/d for women (Food and Nutrition Board, 2005). Also, for

cardiovascular health benefits, the long-chain desaturation and elongation products,









eicosapentaenoic acid (EPA, 20:5) and docosahexaenoic acid (DHA, 22:6) together

should represent 0.3% of dietary energy, with each FA being at least 0.1% of energy

(Simopoulos et al., 2000). These values represent, in general, a decrease in n-6 FA

consumption and an increase in n-3 FA consumption for the typical individual, altering

the current n-6 to n-3 ratio from 10-20:1 to 1-4:1 (Simopoulos, 1999).

Long-Chain Polyunsaturated Fatty Acids of the n-6 Family

Organs and tissues performing storage (adipose tissue), chemical processing (liver),

mechanical work (muscle), and excretion (kidney) have membranes in which the n-6 FA

predominate, particularly with AA as the major component (Innis, 1991). Arachidonic

acid serves an important role as a precursor for biologically important eicosanoids and it

and other n-6 FA may play a role as secondary messengers in the process of signal

transduction. As previously stated, LA can be desaturated and elongated in mammals to

produce biologically important long-chain PUFAs of the n-6 family (Klenk and

Mohrhauer, 1960; Mead, 1968). These include y-linolenic acid (GLA, 18:3) and AA.

Normally, only a small proportion of dietary linoleate can be converted to longer-chain

PUFA. Most of it is P-oxidized to provide energy (Cunnane and Anderson, 1997). To

produce GLA, A6 desaturase acts on LA, introducing a double bond at carbon six in the

FA chain. This product is then elongated to dihomo-y-linoleic acid (20:3, n-6), which is

converted to AA by A5 desaturase. Arachidonic acid can be elongated to form adrenic

and co-6-tetracosatetraenioc acids (22:4 and 24:4), but since there is no evidence of a

functional mammalian A4 desaturase, co-6-docosapentaenoic acid (22:5) must be formed

via an alternate pathway. A double bond is added to tetracosatetraenioc acid by A6









desaturase, forming tetracosapentaenoic acid, which is then oxidized in peroxisomes to

form the 22 carbon product (Sprecher et al., 1995; Ferdinandusse et al., 2001).

Long-Chain Polyunsaturated Fatty Acids of the n-3 Family

Nervous tissue, reproductive organs, and the retina have membranes that contain a

large percentage of long-chain FA, particularly PUFAs of the n-3 series (Innis, 1991). As

with LA and the n-6 PUFAs, dietary LNA can be elongated and desaturated to form

long-chain PUFAs of the n-3 family. The key n-3 PUFAs are EPA and DHA. Within the

body, the amounts of n-3 PUFAs are lower than that of the n-6 PUFAs. This is due to the

small proportion of LNA in the diet, as well as the competition between FA for the A5

and A6 desaturases (Dang et al., 1989). To produce EPA, LNA is desaturated by A6

desaturase to stearidonic acid (18:4), then elongated by 2 carbons, and further desaturated

by A5 desaturase. As with PUFA of the n-6 series, no A4 desaturase is present in

mammals, as in lower eukaryotes, to form DHA directly (Qiu et al., 2001). To form

DHA, EPA undergoes two cycles of chain elongation to produce co-3-tetracosapentaenoic

acid, which is then desaturated to co-3-tetracosahexaenoic acid (24:6) (Sprecher et al.,

1995). In mammals, EPA and DHA can be derived not only from dietary LNA, but also

in the diet directly from sources such as cold-water fish and fish oil.

Digestion and Assimilation of Dietary Fats

In the Western diet fats constitute approximately 40% of energy in the diet. In the

human diet, the majority of fat consumed, whether of animal or plant origin, is in the

form of TAG. Triacylglycerols are the major biological form of storage lipid, composed

of three FA esterified to a glycerol backbone. Long-chain FA, such as oleic acid (18:1)

and palmitic acid (16:0) are the major FA present in dietary TAG, although FA can vary









in chain length from C2 to C24 and from saturated FA to unsaturated FA with six or

more double bonds. In addition to TAG, smaller amounts of phospholipids, cholesterol,

and other sterols are consumed in the diet. An average adult on a Western diet consumes

approximately 150 grams of TAG and 4-8 grams of phospholipids daily. Cholesterol

intake can vary depending on diet, but average daily intake of total cholesterol is 400-500

mg (Rizek et al., 1974).

Although the majority of TAG digestion occurs in the small intestine, digestion

begins in the stomach. Partial hydrolysis of TAG begins with the actions of lingual or

gastric lipase, depending on the species studied (Mu and Hoy, 2004). Lingual lipase is

secreted by the von Ebner's glands of the tongue and is transported with the food bolus to

the stomach (Hamosh and Scow, 1973). Gastric lipase is secreted from the gastric

mucosa. Secretion of either of these lipases can be stimulated mechanically (suckling

and swallowing), neurally (sympathetic agonists), and by diet (high fat) (Hamosh, 1978).

The relative contribution of these lipases to fat hydrolysis is species dependent. For

example, rodents have a relatively high activity of lingual lipase and low activity of

gastric lipase, whereas, in primates, gastric lipase has high activity (Mu and Hoy, 2004).

Both lingual and gastric lipases show a stereo-specific preference for cleaving TAG at the

sn-3 position, regardless of the FA present, although short- and medium-chain FA are

hydrolyzed at a faster rate than long-chain FA (Jensen et al., 1983). This preferential

cleavage gives rise to diglycerides and non-esterified fatty acids (NEFA) as major

digestion products. Approximately 10-30% of dietary fat is partially hydrolyzed in the

stomach which facilitates further digestion in the small intestine (Hamosh and Scow,

1973). In addition, the churning action of the stomach creates a coarse emulsion









stabilized by phospholipids, and proteolytic digestion in the stomach serves to release fats

from food particles where they are generally associated with proteins (Gurr et al., 2002).

The major digestion of dietary TAG results from the actions of pancreatic lipase.

Entry of TAG, TAG degradation products, and acidic stomach contents into the

duodenum causes gall bladder emptying and secretion of pancreatic lipase and

cholecystokinin (Meyer and Jones, 1974). Bile acids serve to emulsify the fats and

increase the available surface area for enzymatic action, where pancreatic lipase and

colipase act to hydrolyze TAG. Colipase attaches to the ester bond of the TAG, which in

turn strongly binds the lipase (Patton, 1981). Pancreatic lipase cleaves the sn-1 and sn-3

bonds specifically, leading to the formation of 2-monoacylglycerols and free FA, with

small amounts of 1,2- and 2,3-diacylglycerols as intermediate products (Mattson and

Volpenhein, 1964). Although pancreatic lipase attacks primarily at stereospecific

locations, the relative rate of hydrolysis depends on the FA present. The lipase has much

slower activity when long-chain FA, particularly the n-3 polyunsaturated FA (20:5 and

22:6), are located in the sn-3 position (Ikeda et al., 1995). Additionally,

2-monoacylglycerols can isomerize to 1-monoglycerides to a small extent in aqueous

conditions, allowing for the formation of a small percentage of glycerol and free FA

(Mattson and Volpenhein, 1962). Phospholipids undergo a similar hydrolysis as TAG,

however the specific enzyme, phospholipase A2, cleaves FA from the sn-2 position of the

phosphoglyceride (van Deenen and deHass, 1963). Dietary cholesterol enters the

duodenum as both free and esterified cholesterol. Prior to absorption, the esterified

cholesterol is hydrolyzed to free cholesterol and NEFA by cholesterol esterase (Hyun et









al., 1969). Cholesterol esterase may also aid in the hydrolysis of TAGs that contain

long-chain PUFA (Carlier et al., 1991).

Lipid absorption in humans begins in the distal duodenum and is completed in the

jejunum. Non-esterified fatty acids and 2-monoacylglycerols, along with phospholipids,

enter into bile micelles, forming mixed micelles. This solubilization allows the non-polar

lipids to travel through the unstirred water layer and reach the brush-border membrane of

the enterocyte (Dietschy et al., 1971; Wilson et al., 1971). The pH of the unstirred water

layer promotes protonation of NEFA, allowing them to more easily leave the micelles

and move to the epithelial cell membrane.

Once in close contact with the brush-border, the 2-monoacylglycerols, NEFA, and

free cholesterol cross the microvillus membrane. In the past, it had been thought that FA

pass into the enterocyte via passive diffusion due to high intraluminal and low cytosolic

concentrations of lipids (Keelan et al., 1992). More recently, however, it has been

proposed that a specific transport protein facilitates the movement of FA into the cell.

Two such proteins that may be involved in intestinal lipid transport are plasma membrane

fatty acid binding protein and fatty acid translocase (Frohnert and Bernlohr, 2000). Bile

salts and some cholesterol are not absorbed and pass to the ileum, where they are

recycled via the portal blood to the liver.

Once within the enterocyte, FAs are re-esterified into TAG and phospholipids in

a multi-step process. First, FAs bind to a cytosolic fatty acid binding protein (FABPc),

allowing for targeting to the ER (Cartwright et al., 2000). There, acyl-CoA synthetase, a

membrane-associated enzyme, activates FAs to their acyl-CoA thioesters via an

ATP-dependent mechanism. This activation effectively traps FA within the cell,









maintaining the concentration gradient and increasing the rate of TAG synthesis

(DiRusso and Black, 1999). Since the major forms of absorbed lipids in humans and

other non-ruminants are 2-monoacylglycerols and NEFAs, resynthesis of approximately

80% of the TAG occurs via the monoacylglycerol pathway (Lehner et al., 1993). In this

pathway, the first step is the acylation of 2-monoglygerides with fatty-acyl-CoA to

diacylglycerols by monoacylglycerol acyltransferase (MGAT). Monoacylglycerol

acyltransferase has a preference for medium-chain saturated and long-chain unsaturated

2-monoacylglycerols (Coleman and Haynes, 1984), but all acyl-CoA studied are

incorporated with similar efficiency (Bugaut et al., 1984). The reaction produces

predominantly 1,2-diacylglycerols, with only about 10% 2,3-diacylglycerols formed

(Lehner and Kuksis, 1996). This stereospecificity allows for the final and rate limiting

step in TAG synthesis. Diacylglycerol acyltransferase (DGAT), which will not act on the

2,3-isomer, acetylates diacylglycerol in an acyl-CoA dependent manner (Coleman, 1988).

Similarly to MGAT, DGAT shows substrate specificity for di-unsaturated or

mixed-diacylglycerols over disaturates.

During fat absorption, the resynthesized TAG are packaged in the enterocyte into

lipoproteins, making the lipids stable for transport in the aqueous environment of the

blood. The human intestine secretes mainly chylomicrons (CM) and very low-density

lipoproteins (VLDL). During fasting, VLDLs are the main lipoproteins secreted by the

small intestine, whereas CMs are secreted during fat feeding (Ockner et al., 1969).

Chylomicrons are the main route of transport for long-chain dietary FAs. Medium-chain

FA (C<12) are absorbed in the non-esterified form, passing directly into the portal blood

system. This occurs because short- and medium-chain FA are more likely to occupy









position three of the TAG and are therefore hydrolyzed in the small intestine and not

retained as 2-monoacylglycerols (Sethi et al., 1993). Soon after dietary lipids enter the

enterocyte, fat droplets can be seen in the ER from the formation of TAG. The rough ER

is the site of synthesis of phospholipids and apolipoproteins, which provide a coat to

stabilize the lipid droplet. Specifically, apolipoprotein B48 (apo B48) associates with the

TAG during its synthesis, forming the immature CM (Cartwright et al., 2000). In the

smooth ER, the immature CM accumulates further TAG via the actions of microsomal

transfer protein (MTP). The CMs then migrate through the Golgi apparatus, where

glycosylation takes place (Leblond and Bennett, 1977) before the fully-formed CM are

exported in secretary vesicles. The CM-containing vesicle travels to the basolateral

surface of the enterocyte, fuses with the plasma membrane, and is secreted into the

extracellular space by exocytosis (Sabesin and Frase, 1977). Very low-density

lipoproteins, as mentioned above, are formed in the small intestine when the levels of

lipids are too low to form CMs. Very low-density lipoproteins differ from CMs in their

density, size, lipid content, and composition, and although both are formed in the same

organelles, the two particles are not mixed in individual Golgi vesicles (Mahley et al.,

1971).

Lipoproteins secreted from the intestine do not enter the blood stream directly.

Instead, they are secreted into minute lymph vessels, known as lacteals, due to their

milky appearance when filled with lipid. From there, the CM and VLDL enter the

circulation in the subclavian vein via the thoracic duct (Mu and Hoy, 2004). Once in the

blood stream, intestinal lipoproteins come into contact with other plasma lipoproteins,

where transfer of protein and TAG occurs (Redgrave and Small, 1979). In particular,









CM and VLDL acquire apolipoprotein CII (apo C-II), which is essential for further

metabolism. As CM and VLDL pass through capillaries, they come into contact with and

bind to lipoprotein lipase (LPL), which is expressed in extrahepatic tissues that use FA,

such as adipose tissue, skeletal and cardiac muscle, and the mammary gland (Ginsberg,

1998). Lipoprotein lipase, with apo C-II as a cofactor, hydrolyzes the TAG in the

particle, generating NEFA that can diffuse into the tissue for further metabolism or

storage (Frayn, 1998). The TAG depleted CM remnant is rapidly removed from plasma

and is metabolized by the liver. Once VLDL have interacted with LPL, they also lose

surface apolipoproteins C and E, and become low-density lipoprotein (LDL) particles

once only apo B remains. The apo B of LDL is recognized by the LDL receptor on the

surface of most cells, allowing for LDL uptake and metabolism within peripheral cells.

The LDL particles are the major carriers of blood cholesterol in humans, pigs, and guinea

pigs; however, in most mammalian species, high-density lipoprotein (HDL) serves this

function.

The reverse transport from peripheral cells to the liver is an important physiological

process necessary to counteract the deposition of cholesterol in tissues from VLDL and

LDL cholesterol. In reverse transport, HDL, primarily synthesized by the liver (Wang

and Briggs, 2004), takes cholesterol from peripheral tissues and transports it to the liver

for metabolism. In 1968, it was first recognized that reverse cholesterol transport

involved the active transport of cholesterol, as cellular free cholesterol was converted to

the insoluble ester outside of the cell. The enzyme involved in this process is

lecithin:cholesterol acyl-transferase (LCAT), and is a component of HDL that increases

the cholesterol esters within this lipoprotein fraction (Glomset, 1968). The rate of LCAT









is affected primarily by the surface properties of individual lecithin molecules (Pownell et

al., 1985). Two additional proteins contribute to HDL remodeling, both by working

down concentration gradients in an energy-independent manner. Phospholipid transfer

protein supplies lecithin to HDL (Tollefson et al., 1988), and a cholesterol ester transfer

protein can move cholesterol esters made by LCAT to other lipoproteins, particularly

LDL (Tall, 1993). The TAG portion of HDL can be catabolized by the extrcellular

hepatic triacylglycerol lipase, and the cholesterol is removed by the liver via several

different mechanisms (Nagata et al., 1988; Wang and Briggs, 2004). It is only tissues

that actively uptake or synthesize cholesterol that contribute to the reverse cholesterol

transport pathway.

Dietary Fats in Relation to Health

Dietary Fats in Relation to Weight Control

According to the World Health Organization (World Health Report, 2002), obesity

rates have risen over three-fold since 1980 in most developed and developing countries

worldwide. Current estimates count more than one billion adults as overweight and at

least 300 million as clinically obese. In the US, approximately 30% of adults are

categorized as obese, which is defined as at least 20% heavier than their ideal weight.

Obesity is associated with increased early mortality and an increased risk for a variety of

diseases, including metabolic, cardiovascular and gastrointestinal diseases. Because of

this, the World Health Organization has listed obesity as one of the top ten global health

problems in Western cultures (World Health Report, 2002).

Analysis of epidemiological data suggests that dietary fat plays a role in obesity,

though the mode of action is not clear (Bray and Popkin, 1998; Astrup et al., 2000). It is

evident however, that it is not a simple relationship. Longititudinal measurements of









food intake in both the US and the UK show that fat intake has not increased as a

proportion of dietary energy over the last 30 years, unlike the rising trends in obesity

(Heini and Weinsier, 1997; Nielsen et al., 2002). Obesity may be due to the types of fat

consumed or the interaction of dietary fats or FAs with other dietary compounds.

If high intakes of dietary fat are a factor in the development of obesity, then

reducing the fat in the diet would be expected to produce weight loss. Studies examining

the relationship between fat in the diet and changes in body weight reported several

conclusions. When animals were fed a high fat diet, almost all species develop obesity,

as demonstrated in primates, rodents, pigs, dogs, and cats (West and York, 1998).

Exceptions include animals with a strong genetic component to obesity, such as C57/BL

mice and Osborne-Mendel rats (Bray et al., 2004). Reductions in body weight of subjects

consuming a low-fat diet are modest and tend to extend over only a short period of time

(Jeffery et al., 1995), and the higher the body mass index (BMI) of the subject, the greater

the weight loss (Astrup et al., 2000). There is also a positive relationship between the

percent reduction in dietary fat and the decrease in energy intake, suggesting that the

major mechanisms for weight loss associated with reduced-fat diets may be primarily

through a lower energy intake (Bray et al., 2002). Dietary fat, therefore, may not be an

independent cause for obesity. Excessive energy intake, whatever the source, and

decreased energy expenditure generally are considered the main causes of obesity (Foreyt

and Poston, 2002). Current recommendations still call for a low-fat intake, due to fat's

higher caloric density than other nutrients, coupled with less energy expenditure of an

increasingly sedentary population (Astrup et al., 2002). When this theory is considered,

it is evident that the type or composition of dietary fat may have little effect on obesity.









However, the type of fat consumed can play significant, differential, and more direct

roles in other health and disease states.

Dietary Fats and Blood Cholesterol

Dyslipidemia is a condition in which plasma concentrations of LDL cholesterol and

TAG are elevated and HDL cholesterol is lower than found in normal, healthy

individuals. According to the most recent guidelines set by the US National Cholesterol

Education Program (NCEP), total cholesterol should be <200 mg/dL, with LDL

cholesterol <100 mg/dL and HDL cholesterol >40 mg/dL (Grundy et al., 2004).

Abnormally high LDL cholesterol and low HDL cholesterol outside the recommended

values are considered significant risk factors for cardiovascular disease, therefore

maintaining optimum blood concentrations is beneficial. Dietary fat has the ability to

modify blood cholesterol components in both a positive and negative manner, depending

on the types of fat consumed.

As a group, consumption of saturated FAs raises total and LDL cholesterol in

blood, but individual saturated fats can have differing effects (Reddy and Katan, 2004).

Several feeding studies have demonstrated that individuals consuming diets high in

saturated fat had increased concentrations of both HDL and LDL cholesterol (Kromhout

et al., 1995). Myristic (14:0) and lauric (12:0) acids have a greater effect on elevating

LDL cholesterol than palmitic acid (16:0), but, among these, palmitic acid is greatest in

the food supply. Stearic acid, in contrast, decreases plasma and liver cholesterol

concentrations, primarily by reducing intestinal cholesterol absorption. The mechanism

by which stearic acid reduces cholesterol is thought to be by reducing solubility of

cholesterol and altering the population of microflora that can synthesize secondary bile

acids (Cowles et al., 2002).









In the human diet, the predominant monounsaturated fatty acid (MUFA) is oleic

acid (18:1, n-9). It is found at high levels in olive oil, canola oil, and nuts. The

Mediterranean diet, which is not low in fat but is associated with a healthy blood lipid

profile, contains a high percentage of fat as oleic acid. When saturated fats in the diet are

replaced with oleic acid, total and LDL cholesterol concentrations are lowered (Gardner

and Kramer, 1995). This seems to be caused by a passive mechanism; when saturated

fats are decreased and MUFA are increased, the fat induced-suppression of LDL receptor

activity is less and LDL uptake into cells is increased (Dietschy et al., 1993). Effects of

MUFA on HDL cholesterol are less clear. Some studies have indicated that MUFA have

no effects on blood HDL concentrations (Delaplanque et al., 1991; Mata et al., 1992).

This combined with the LDL-lowering effects suggest that MUFA can shift the

LDL:HDL ratio towards a healthier profile. However, upon extensive meta-analysis, the

effects of MUFA on HDL cholesterol in blood could not be confirmed (Gardner and

Kramer, 1995).

Polyunsaturated fatty acids of the n-6 family, particularly LA, lower total and LDL

cholesterol when they are supplied in the diet in place of saturated fats (Kris-Etherton and

Yu, 1997). In addition to the passive mechanism described for MUFA, PUFA actively

increase receptor-dependent LDL uptake, although this is a small effect (Dietschy et al.,

1993. Dietschy, 1998). In some studies in which n-6 PUFA replaced saturated fats, a

significant decrease in HDL cholesterol was reported (Shepherd et al., 1978; Jackson et

al., 1984), although this is not a consistent effect (Iacono and Dougherty, 1991). When

n-3 PUFA were supplemented with the regular diet, LDL cholesterol was raised in some

studies (Harris et al., 1988; Fumeron et al., 1991) and HDL cholesterol was either









unchanged or slightly increased (Harris, 1989). These effects tend to be more

pronounced in hyperlipidemic subjects. For example, an increase in LDL cholesterol

occurred in isolated hypertriglyceridemiac subjects when more than 10 g of n-3 FA were

supplemented per day (Schmidt and Dyerberg, 1994). When very long-chain n-3 FA,

such as EPA and DHA were specifically supplemented, they not only have the ability to

significantly lower serum TAG, but also to increase LDL cholesterol more so than

supplementing with LNA or a mixture of n-3 FA (Harris, 1997).

Trans-FA are geometrical isomers of unsaturated FA that assume a saturated fat-

like configuration. The predominant source in the human diet is from hydrogenated oils,

such as margarine and partially hydrogenated soybean oil, commonly found in baked

goods and deep fat-fried fast foods (Hu et al., 2001). Metabolic studies have shown that

consumption of trans-FA has the ability to negatively alter the lipid profile to a greater

extent than saturated fats, because they not only increase small, dense LDL cholesterol

(Mauger et al., 2003), but also decrease HDL cholesterol (Judd et al., 1994; de Roos et

al., 2003). This leads to an increase in the ratio of total to HDL cholesterol that is

approximately double that observed with saturated fats (Willett and Ascherio, 1994).

Additionally, diets high in trans-FA are associated with raised TAG concentrations

(Katan and Zock, 1995), an independent risk marker of cardiovascular disease.

Dietary Fats and Cardiovascular Disease

Most of the FAs in the Western diet are derived from meats, oils, and dairy

products, leading to a large intake of saturated and MUFAs, with a relatively small

proportion of PUFA consumed. Saturated fat and cholesterol represent two of the most

established dietary risk factors for cardiovascular disease (CVD), whereas MUFA and

PUFA are likely to provide beneficial effects with increased amounts in the diet. These









effects are partially due to the effects on the blood lipid profile, but the risks associated

with intake of certain fats are greater than would be expected from cholesterol effects

alone.

As stated above, intake of certain saturated fats can increase LDL and decrease

HDL cholesterol, creating an atherogenic lipid profile. A recent analysis of the Nurses'

Health Study revealed that intake of short to medium chain FA was not associated with

increased coronary heart disease (CHD) risk. In the same analysis, however, intakes of

longer chain saturates, particularly stearic acid, were associated with an increased risk of

CHD (Hu et al., 1999). Additionally, stearic acid may negatively impact other markers of

atherogenesis. Stearic acid can increase lipoprotein(a) concentration (Aro et al., 1997),

and may activate Factor VII (Mitropoulos et al., 1994) and impair fibrinolysis (Ferguson

et al. 1970). On the positive side however, when compared with consumption of palmitic

acid, stearic acid decreases platelet volume, platelet aggregation, and coagulation factor

VII activity (Kelly et al., 2001). Due to the many negative health implications of a diet

high in saturated fats, there is a consensus to reduce the intake of saturated fats to less

than 10% of the total daily energy supply (American Heart Association, 2000).

The Seven Countries Study gave the first epidemiological evidence for a negative

correlation between dietary intake of MUFA and mortality from CHD. Mortality was

noticeably low in Mediterranean countries, where olive oil is the main source of fat (Keys

et al., 1986). In addition to the positive effects on plasma LDL cholesterol levels, diets

rich in olive oil can improve endothelial function, as compared to a high saturated fat diet

(Fuentes et al., 2001), and attenuate postprandial endothelial dysfunction that follows a

fatty meal (Vogel et al., 2000). Intake of MUFA is also protective against LDL









oxidation. Due to their structure, MUFA are more stable and less susceptible to lipid

peroxidation. A high intake of MUFA results in a greater incorporation into LDL

cholesterol (Mata et al., 1996). Oxidation of LDL cholesterol prevents its recognition by

the LDL receptor and subsequent uptake into cells. It is instead taken up by the

scavenger receptors of macrophages, leading to the accumulation of cholesterol and the

formation of fatty streaks. These processes promote the development of atherosclerosis

(Westhuyzen, 1997). Several studies have shown that dietary sources of MUFA other

than olive oil are associated with an increased CHD risk (Posner et al., 1991; Esrey et al.,

1996). However, these studies did not correct for potential confounding effects, such as

the intake of other FAs and antioxidants.

Epidemiological evidence supports a role for dietary LA in reducing the risk of

CHD. High adipose LA in healthy men is associated with lower CHD mortality

(Riemersma et al., 1986), while low dietary intake of LA predisposes to myocardial

infarction (Simpson et al., 1982). A more recent study of Japanese subjects found

reduced serum LA in patients with ischemic stroke as compared to healthy controls (Iso

et al., 2002). Similarly to LA, LNA intake was inversely associated with mortality from

CHD in the Multiple Factor Intervention Trial (Dolecek, 1992). Large prospective

studies in both men and women have found that LNA protected against both cardiac

deaths and nonfatal myocardial infarction (Ascherio et al., 1996; Hu et al., 1999). The

effects of LNA on plasma lipids are not large; therefore the reduction in CHD risk may

have more to do with cardiac function, such as arrhythmia, inflammation, and

thrombosis.









The low rate of CVD seen in several communities consuming a diet rich in fish

(Bang et al., 1980; Kromhout et al., 1985; Hirai et al., 1989; Oomen et al., 2000) has

prompted investigations into how fish and its nutritional components may lower the risk

of CVD. Fish, particularly fatty fish such as tuna, mackerel, and salmon, are rich in the

n-3 FA EPA and DHA (Parkinson et al., 1994). High serum and adipose tissue

long-chain n-3 PUFA have been associated with reduced risk of fatal myocardial

infarction (Simon et al., 1995; Pedersen et al., 2000; Lemaitre et al., 2003), primary

cardiac arrest (Siscovick et al., 1995), and sudden cardiac death (Albert et al., 2002).

Upon meta-analysis of 11 randomized controlled trials comparing long-chain n-3 PUFA

intake to placebo or control diets, intake of long-chain n-3 PUFA was associated with

lower cardiac fatalities in patients with CHD (Bucher et al., 2002). However, these FA

did not protect against nonfatal cardiac events or total morbidity (Erkkila et al., 2003;

Lemaitre et al., 2003), suggesting that the hypolipidemic effects of DHA and EPA on

atherosclerosis are distinct from those effects associated with arrhythmic myocardial

dysfunction. In a canine model with dogs made susceptible to fatal ventricular

fibrillation and sudden cardiac death, infusion of EPA and DHA reduced cardiac deaths

by preventing ventricular fibrillation (Billman et al., 1997; Billman et al., 1999). As the

fat infusion was given only one hour prior to inducing ischemia, the effects are not likely

by membrane incorporation of n-3 PUFA, but rather by direct action of nonesterified

PUFA on the myocytes. In support of this, induction of arrhythmias in cultured neonatal

rat myocytes was abolished by the addition of EPA or DHA to the culture medium (Kang

and Leaf, 1996), and supplementation of four g/d of EPA and DHA increased heart rate

variability in survivors of myocardial infarction, reducing the risk of subsequent









arrhythmic events (Christensen et al., 1996). The anti-arrhythmic actions of EPA and

DHA seem to be associated with the ability of these FA to prevent calcium overload in

cardiac myocytes during periods of stress (Leaf and Kang, 1997).

As stated previously, trans-FA negatively impact the blood lipid profile in several

ways. However, the relationship between consumption of trans-FA and cardiovascular

risk is greater than is predicted based on these lipid changes (Ascherio et al., 1999),

suggesting effects on other distinct risk markers for CVD. In humans, it has been shown

that trans-FA increase lipoprotein (a) levels (Nestel, et al., 1992; Sundram et al., 1997),

which are positively associated with increased risk of CHD (Utermann, 1989).

Additional studies have examined the effects of trans-FA on markers of low-grade

chronic inflammation. The Nurses' Health Study showed that a high intake of trans-FA

was positively associated with concentrations of tumor necrosis factor a receptors 1 and 2

(Mozaffarian et al., 2004). A dietary intervention study in which 8% of dietary energy

from carbohydrates, oleic acid, or stearic plus trans-FA was replaced with trans-FA

supported this epidemiological data, and additionally found increases in plasma

C-reactive protein, IL-6 and E-selectin with the trans-FA diet (Baer et al., 2004).

Conjugated Linoleic Acid

Conjugated linoleic acid (CLA) is the collective term for a group of positional and

geometric conjugated dienoic isomers of LA. These FA are considered conjugated

because, unlike other FA, the double bonds occur on adjacent carbons, and are not

separated by a methylene group. To date, 16 CLA isomers have been identified (Rickert

et al., 1999), with double bonds ranging in position from carbons 6 and 8 to carbons 12

and 14. The double bonds can occur in pairs of geometric isomers as cis-cis, cis-trans,

trans-cis, and trans-trans. However, only two isomers (cis-9, trans-11 CLA and









trans-10, cis-12 CLA) are known to possess biological activity (Pariza et al., 2000).

Sources of CLA in the human diet are ruminant products and synthetic supplements,

though the specific makeup of CLA differs among sources. In milk, cheese, and

ruminant meat, which can contain 2-8 mg of CLA/g lipid depending on the source (Lin et

al., 1995; Chin et al., 1992), approximately 80% of the CLA is cis-9, trans-11 CLA and

10% is trans-10, cis-12 CLA (Fogerty et al., 1988). Based on this, recent studies suggest

average intakes of 150-200 mg of CLA per day (Jiang et al., 1999; Ritzenthaler et al.,

2001), with intakes as high as 650 mg/day on a diet rich in animal fats (Park et al.,

1999a). Conjugated linoleic acid dietary supplements, produced by the chemical

isomerization of LA, contain predominantly cis-9, trans- 1 CLA and trans-10, cis-12

CLA in equal amounts (Gaullier et al., 2002).

The cis-9, trans-11 CLA isomer is produced as an intermediate in the rumen during

the biohydrogenation of dietary LA. A key anaerobic bacterium in this process is

Butyrivibriofibrisolvens (Kepler et al., 1966). The cis-12 bond of LA is acted upon by

the microbial isomerase, forming cis-9, trans-11 CLA. In some of the literature, this

isomer is also referred to as rumenic acid. This CLA product can leave the rumen and be

directly absorbed, or it can be further metabolized by ruminal microbial hydrogenases,

forming trans-vaccenic acid (trans-11, 18:1) before being completely hydrogenated to

stearic acid (Kepler et al., 1966). This product may also exit the rumen and be absorbed

and transported to peripheral tissues. In the mammary tissue and muscle, a A9-desaturase

is present that can act on trans-vaccenic acid to produce cis-9, trans-11 CLA (Holman

and Mahfouz, 1980; Pollard et al., 1980). This has been shown to occur in several

mammalian species, including ruminants (Griinari et al., 2000), mice (Santora et al.,









2000), and humans (Turpeinen et al., 2002). Certain ruminal bacteria also have the

capability to convert LA to trans-10, cis-12 CLA by isomerizing the cis-9 bond (Griinari

and Bauman, 1999). This can be hydrogenated to form trans-10 octadecenoic acid,

which may be absorbed and transported to peripheral tissues, but since mammals do not

possess a A12-desaturase, it would not be converted back to trans-10, cis-12 CLA.

Numerous beneficial physiological effects have been attributed to CLA. The

seminal observation came when CLA isolated from grilled beef inhibited

chemically-induced skin neoplasia in mice (Ha et al., 1987). This discovery led to

research examining the effects of CLA on cancer (Ha et al., 1990), immune function

(Miller et al., 1994), atherosclerosis (Lee et al., 1994), weight gain and food intake (Chin

et al., 1994), and body composition (Park et al., 1997). As previously stated, the two

biologically active isomers of CLA are cis-9, trans-11 and trans-10, cis-12 CLA.

Though derived from the same parent molecule, the two isomers are structurally and

functionally distinct. Both isomers contain a trans double bond, creating a straighter

carbon chain, as opposed to the "kink" created by the cis configuration. Many enzymes

recognize specific configurations in FA; therefore it is not surprising that differences in

bond position and orientation of CLA isomers give them differing biological activities.

Numerous studies now indicate that the various physiological and biological effects of

CLA may be due to the separate actions of the cis-9, trans-11 and trans-10, cis-12

isomers (Pariza et al., 2000).

Dietary CLA modulates body composition through decreases in adiposity and

increases in lean mass in various animal models. The effects in mice are the most

dramatic, with a 50-60% reduction in total adipose mass in animals fed mixed isomers of









CLA over a 4-5 week period, as compared to mice fed the control diet (Park et al., 1997).

The effect in mice can be sustained, even after removal of CLA from the diet (Park et al.

2001). Additionally, when the trans-10, cis-12 isomer was fed to mice, it was more

effective in lowering adipose tissue mass than cis-9, trans-11 CLA (Park et al., 1999b).

Similar reductions in adipose mass have been noted in Sprague-Dawley and Zucker lean

rats fed CLA, although the effects are not as large as in mice (25-30% reduction) (Sisk et

al., 2001). In contrast to lean rats, obese Zucker rats exhibit an adipose-enhancing effect

of dietary CLA (Szymczyk et al., 2000). In pigs, CLA-feeding decreased fat deposition

and increased lean tissue (Dugan et al., 1997; Thiel-Cooper et al., 2001). In humans,

however, the results are not as clear. Several studies have shown no effects of mixed

CLA or individual isomers in the diet on changes in body composition in human subjects

(Terpstra, 2004), and to date, no studies have shown changes in body weight (Larsen et

al., 2003). Conversely, studies feeding mixed CLA and the trans-10, cis-12 isomer have

reported reductions in body fat mass but no changes in body mass index (Blankson et al.,

2000; Smedman and Vessby, 2001; Riserus et al., 2004). These changes are much less

than those observed in pigs and mice; however, pigs and mice are generally fed at least

five times more CLA per kilogam of body weight than humans (House et al., 2005).

Comparable doses as used in animal studies would correspond to a daily intake of 130 g

in humans (Larsen et al., 2003). Long term supplementation with mixed CLA isomers by

healthy overweight individuals also seems to be well tolerated, although reductions of

body fat mass may or may not be maintained (Gaullier et al., 2005; Larsen et al., 2006).

Mechanisms by which CLA reduces adiposity may involve pathways that involve

energy expenditure. This is shown by increased metabolic rates and reduced nighttime









respiratory quotients in mice fed CLA for six weeks (West and York, 1998). Effects of

CLA also have been linked with the induction of adipocyte apoptosis, both in vivo

(Tsuboyama-Kasaoka et al., 2000) and in vitro (Evans et al., 2000), and with decreased

uptake of TAG into adipocytes, particularly due to the suppression of lipoprotein lipase

activity by trans-10, cis-12 CLA (Park et al., 2001).

Obesity puts an individual at a greater risk for other diseases, including type II

diabetes. It would be expected then, that reduction in fat mass due to CLA intake would

help decrease this risk. Evidence, however supports an additional direct effect of CLA on

diabetes, which can vary, depending on the species studied. In animal models of

diabetes, such as the Zucker diabetic fatty rat, CLA-enriched diets reduce fasting glucose,

insulinemia (Houseknecht et al., 1998), tryglyceridemia and blood NEFA concentrations

(Belury and Vanden Huvel, 1999) as compared with controls. These beneficial effects

may be due, in part, to enhanced muscle uptake of glucose (Ryder et al., 2001). It is

important to note that these effects are seen when a mixture of CLA isomers are fed.

When fed butter enriched with cis-9, trans-11 CLA, little or no effect was seen,

indicating that the effects on glucose tolerance are likely due to the trans-10, cis-12

isomer (Ryder et al., 2001). In contrast with diabetic animals, CLA modestly increases

fasting serum insulin in nondiabetic pigs (Stangl et al., 1999), mice (Tsuboyama-Kasaoka

et al., 2000), and humans (Medina et al., 2000). These negative effects on insulin

resistance may result from decreased plasma leptin concentrations (Wang and Jones,

2004) or an increase in TAG concentration in muscle due to feeding trans-10, cis-12

CLA (Terpstra, 2004).









Similar to other classes of FA, CLA can affect the blood lipid profile and

cardiovascular risk factors. In rabbits fed an atherogenic diet, supplementation with CLA

lowered serum TAG and LDL cholesterol concentrations, as compared to controls (Lee et

al., 1994). These animals fed CLA also showed a decrease in atherosclerotic plaque

formation. Another study with rabbits showed a regression of established atherosclerosis,

despite an increase in total cholesterol and decrease in HDL cholesterol (Kritchevsky et

al., 2000). In a similar model in hamsters fed the cis-9, trans-11 isomer, there was no

effect on plasma lipids (Gavino et al., 2000). Culturing of human platelets with either

CLA isomer inhibited induced platelet aggregation (Truitt et al., 1999), but in human

subjects supplemented with a mixture of CLA isomers, no difference was observed in

platelet aggregation or prothrombin time (Benito et al., 2001). Together, these findings

potentially implicate trans-10, cis-12 CLA to have positive effects on the blood lipid

profile and coronary risk factors.

In addition to the effects on disease states, CLA can alter lipid metabolism. When

consumed, CLA is incorporated into membrane phospholipids and alters FA homeostasis,

particularly in the liver (Belury, 2002). Conjugated linoleic acid that is not broken down

via P-oxidation is desaturated and elongated to other conjugated metabolites (Belury and

Kempa-Steczko, 1997). The competition of CLA with LA for A6-desaturase may result

in decreased AA, and can explain the reduced eicosanoid production in several systems

(Belury, 2002; Brown and McIntosh, 2003). It has also been found that mice (Degrace et

al., 2003) and hamsters (de Deckere et al., 1999) supplemented with CLA, particularly

the trans-10, cis-12 isomer, develop enlarged, fatty livers. This effect has been attributed

to an increase in liver TAG, cholesterol, cholesterol esters, and NEFA (Kelley et al.,









2004), though the mechanism is unclear. Supplementation with trans-10, cis-12 CLA in

vivo and in vitro in various animal and human models leads to an increase in the ratio of

saturated to monounsaturated fats (House et al., 2005). This is likely due to a reduction

in stearoyl-CoA desaturase, which catalyzes the biosynthesis of MUFA from stearic and

palmitic acids (Lee et al., 1998). Trans-10, cis-12 CLA also inhibits transcription of

other genes involved in de novo FA synthesis, desaturation, and TAG synthesis, which

may partially explain its effects on changes in lipid metabolism in the liver (Baumgard et

al., 2002).

Roles of the Peroxisome Proliferator-Activated Receptors in Lipid Metabolism

Peroxisome proliferator-activated receptors (PPAR) belong to the steroid hormone

receptor superfamily that are ligand-activated transcription factors (Wahli and Martinez,

1991), and act by modulating a network of responsive genes. They have been identified

in many species, including Xenopus (Dreyer et al., 1992), mouse (Issemann and Green,

1990), rat (Gottlicher et al., 1992), and human (Sher et al., 1993). The name PPAR

derives from the ability of the first-identified member to induce hepatic peroxisome

proliferation in mice, but this phenomenon seems to be rodent-specific, and does not

occur in other mammals. The PPARs consist of a family of three isoforms: PPARa, -y,

and -P/5 (Issemann and Green, 1990; Dreyer et al., 1992; Kliewer et al., 1994). Though

encoded by separate genes, and different in their tissue distribution and metabolic actions,

all three isoforms are structurally similar and can be activated by FA and their metabolic

derivatives, making them the first recognized lipid sensors in the body (Schoonjans et al.,

1996). Genes whose expression is modified by PPARs are numerous and control glucose

homeostasis, cell cycle, inflammation, immune response, and lipid metabolism

(Desvergne and Wahli, 1999).









Similar to other nuclear receptors, the PPARs possess structural features composed

of functional domains. The DNA-binding domain consists of two zinc fingers that

specifically bind peroxisome proliferator response elements (PPRE) in enhancer sites of

regulated genes (Wahli and Martinez, 1991). The PPRE are specific DNA sequences

formed by the direct repeat of a hexanucleotide sequence (AGGTCA), separated by one

or two nucleotides (Torra et al., 2001). Unlike other steroid receptors which function as

homodimers, to bind to the PPRE, PPAR must form a heterodimer with the retinoid X

receptor (RXR) in the cytoplasm, allowing for transport to the nucleus (Miyata et al.,

1994). The ligand binding domain appears to be quite large in comparison with other

nuclear receptors (Nolte et al., 1998; Xu et al., 1999), potentially allowing PPARs to

interact with a broad range of structurally distinct natural and synthetic ligands.

As PPARs play a critical role in lipid metabolism, the search for natural ligands

began with the FAs and eicosanoids. Cell-based transactivation assays and direct binding

studies have identified and characterized the endogenous receptor effectors. In general,

all isoforms of PPAR are more responsive to n-6 and n-3 PUFA than to saturated or

monounsaturated FAs (Krey et al., 1997). However, the affinities for the receptor vary,

suggesting a role for site-specific availability and metabolism of particular FA, as well as

different affinities for the specific PPAR isoforms (Sampath and Ntambi, 2005). It has

been shown that FA such as LA, LNA, and AA can activate PPARa at a concentration of

100 pM (Lehmann et al., 1997). Additionally, EPA is a much more potent activator of

PPARa than arachidonic acid in primary hepatocytes (Ren et al., 1997). Since the

concentration of NEFA in human blood can be greater than 1 mM, these FA can be

considered potent endogenous ligands for PPARa. It is important to note, however, that









the intracellular concentrations of PUFA are not known. Like PPARa, PPARy has

affinity for the PUFAs, as well as metabolic derivatives of PUFAs, such as

9-hydroxyoctadecadienoic acid (HODE) and 13-HODE (Nagy et al., 1998), and CLA

(Hontecillas et al., 2002). Peroxisome proliferator-activated receptor 6 also interacts with

saturated and unsaturated FA, but with a ligand specificity that is intermediate between

that of PPARy and PPARa (Berger and Moller, 2002). Even with the abundance of

natural ligands for PPARs, the emphasis in recent years has been on the development of

synthetic ligands, due to their greater therapeutic and commercial value. Fibrates, which

are ligands for PPARa, and thiazolidinediones (TZD), which are ligands for PPARy, are

two classes of drugs used to treat hypocholesterolemia and type II diabetes.

PPARa

The first PPAR discovered (Issemann and Green, 1990), PPARa is expressed

predominantly in the liver, kidney, heart, brown fat, and skeletal muscle (Braissant et al.,

1996; Auboeuf et al., 1997), as well as in monocytic (Chinetti et al., 1998), vascular

endothelial (Inuoe et al., 1998), and vascular smooth muscle cells (Staels et al., 1998). It

plays an important role in lipid metabolism via regulation of the expression of genes

involved in cellular free FA uptake, P-oxidation, and cellular cholesterol trafficking (Li et

al., 2002). It has been reported that PPARa is greatly induced during fasting or starvation

in which a switch from carbohydrates and fats to mostly fats as an energy source is

required. During fasting, FA released from the adipose tissue are taken up by the liver,

where they are re-esterified to TAG or broken down via P-oxidation to ketones.

Peroxisome proliferator-activated receptor a induces expression of fatty acid translocase

(Motojima et al., 1998) and fatty acid transport protein (Martin et al., 1997), genes

involved in transport of FA into the cell, as well as CPT-I (Brady et al., 1999), which









catalyzes the rate limiting step for transport of FA into the mitochondria for oxidation.

Activation of PPARa also directly upregulates genes involved in peroxisomal

P-oxidation, including acetyl-CoA synthase (Schoonjans et al., 1995) and ACO

(Tugwood et al., 1992). The importance of PPARa in this response has been

demonstrated by studies involving PPARa-null mice, which are unable to induce the

change in energy source, resulting in hypoglycemia, hyperlipidemia, hypoketonemia and

fatty liver (Kersten et al., 1999). In rodents, activation of PPARa induces peroxisome

proliferation, hepatomegaly, and hepatocarcinogenesis (Issemann and Green, 1990).

Fortunately, these effects are not present in humans, possibly due to the 10-fold greater

concentrations of PPARa in rodent as compared to human liver (Palmer et al., 1998) or to

differences in the PPREs of responsive genes, such as ACO (Lambe et al., 1999).

Fibrates have been a commonly prescribed drug to treat dyslipidemia in humans for

over 30 years, but the direct role of PPARa in the lipid-lowering actions of fibrates has

only recently been established. In humans, fibrate administration lowers plasma TAG

and increases plasma concentrations of HDL and its major constituents, apolipoproteins

A-I (apo A-I) and A-II (apo A-II) (Malmendier and Delcroix, 1985; Mellies et al., 1987).

Peroxisome proliferator-activated receptor a activation affects several key genes in HDL

metabolism, including apo A-I, apo A-II, ABCA1, LPL, and scavenger receptor class B

type I (Fruchart, 2001). Peroxisome proliferator-activated receptor a also has been

shown to down-regulate apo C-III (Hertz et al., 1995; Staels et al., 1995), a protein that

inhibits TAG hydrolysis by LPL, further contributing to the lipid-lowering effects of

fibrates.









Surprisingly, the role of PPARa in cardiovascular disease appears to be negative.

In a mouse model, over-expression of PPARa in the heart increases FA oxidation and

decreases glucose use, similar to that seen in the diabetic heart. Upon fibrate

administration, these mice develop greater cardiomyopathy than the wild-type controls

(Finck et al., 2002). Peroxisome proliferator-activated receptor a null mice do not show

this effect (Finck et al., 2003). This knowledge, when combined with research indicating

that PPARa and apolipoprotein E double knockout mice are resistant to insulin-resistance

and atherosclerotic lesions induced by a high-fat diet (Tordjman et al., 2001), suggests

that PPARa senses FAs and induces their use, thereby playing a potential causative role

in cardiovascular disease. Unlike humans, in rodent models, fibrate administration

decreases apo A-I and apo A-II expression, suggesting differential regulation in the

different species (Berthou et al., 1995). Overall reduction in TAG and increase in HDL

cholesterol in humans, even with potential for negative cardiovascular events, would still

result in less fat accumulation in the vessel walls, and would be beneficial to heart health.

PPARp/6

SPeroxisome proliferator-activated receptor P/6 (hereafter referred to as PPARS) has

been slighted in its importance in the body because of its ubiquitous expression and

unavailability of selective ligands, despite the fact that it is the predominant isoform in

skeletal muscle one of the most insulin responsive and metabolically demanding tissues

of the body. The importance of PPAR6 in FA metabolism was first realized from studies

using knockout animals. Most PPAR6 null mice die during early embryogenesis, and the

numbers that do survive show a marked decrease in fat mass (Peters et al., 2000). In

exercised or fasted PPARa null mice, the liver, but not the muscle glycogen levels









deplete as compared to wild-type litter mates, indicating that a factor other than PPARa

may be in control of energy homeostasis (Muoio et al., 2002).

Recently, synthetic, highly selective PPAR6-agonists have been developed, and its

role in FA catabolism and energy homeostasis has been further elucidated (Peters et al.,

2000; Barak et al., 2002). Activation of PPAR6 increases FA oxidation in human and

rodent myocytes, showing the redundancy of PPARs a and 6 in FA homeostasis (Muoio

et al., 2002). In genetically obese ob/ob mice, PPAR6 activation not only enhances

P-oxidation in skeletal muscle, but protects against diet-induced obesity, improves

glucose tolerance, and improves insulin sensitivity, showing its potential as a target in

treating and preventing obesity and type II diabetes (Tanaka et al., 2003).

Similarly to PPARa, PPAR6 activation up-regulates adenosine

triphosphate-binding cassette transporter-Al (ABCA1) gene expression and increases

cholesterol efflux from cells and increases HDL cholesterol in mice (Leibowitz et al.,

2000) and non-human primates (Oliver et al., 2001). The PPAR6 selective agonist

GW501516, in particular, has shown therapeutic potential for the treatment of

dyslipidemia, by dramatically improving the serum lipid profile of insulin-resistant

rhesus monkeys. This occurs through decreases in concentrations of blood TAG and

insulin and increases in HDL to a greater extent than is achieved with fibrates in fasting

individuals. Activation of PPAR6 has the added normal-lipidemic effect of lowering the

blood concentrations of small dense LDL cholesterol (Oliver et al., 2001).

Recent research suggests a role for PPAR6 in the heart as well. In cultured human

cardiomyocytes, PPAR6 is highly expressed, and its activation leads to an increase in FA

oxidation (Cheng et al., 2004), leading to a potential increase in energy to an energy









demanding organ, and implicating PPAR6 as a modulator of cardiac energy homeostasis.

Regarding foam cell formation, research with different PPAR6 activators have given

different results. In one study, activation increased cholesterol efflux through the ABCAI

pathway (Oliver et al., 2001), whereas another study demonstrated enhanced lipid

accumulation (Vosper et al., 2001). Though these discrepancies may be due to different

experimental models and structurally different agonists, further research can help

elucidate the role of PPARS.

PPARy

Because it is primarily found in adipose tissue, PPARy is a prime suspect in the

regulation of lipid metabolism. In support of this, many studies have shown the

importance of PPARy in the formation and functioning of adult fat cells (Rosen et al.,

2000). As obesity is a primary risk factor for incidence of the metabolic syndrome, it is

highly likely that PPARy plays a role in the associated diseases and their treatment.

Thiazolidinediones are pharmacologic activators of PPARy, which significantly improve

insulin sensitivity in humans with type II diabetes (Sood et al., 2000). The mechanism of

action, however, still remains unclear, especially considering the fact that muscle is the

major insulin responsive tissue, and PPARy is present at very low levels in muscle and

liver and high in adipose. Resolving this apparent paradox had proved difficult. Most

research has stemmed from clinical trials and rodent models of obesity and diabetes.

Unlike the readily available PPARa null mice, PPARy knockout mice die early in

gestation, preventing valuable loss-of-function studies (Barak et al., 1999). By the use of

microarrays for gene expression profiling, several key metabolic genes were identified,

all primarily in the adipocyte. Changes induced by TZD administration to Zucker









diabetic fatty rats include modulation of genes involved in glucose uptake, lipid uptake

and storage, and energy expenditure (Way et al., 2001). The small increases in glucose

disposal by the adipose, coupled with greater sequestration of fat into adipose, thereby

relieving some of the metabolic burden of muscle and liver and allowing for greater

glucose use by these tissues, is a potential explanation for the profound activity of the

TZD class of drugs.

In addition to the action of PPARy ligands on adipose, there is mounting evidence

that these compounds can exert some effects on other tissues. aP2/DTA mice, whose

white and brown adipose tissue has been eliminated by fat-specific expression of

diphtheria toxin A chain, develop hyperglycemia, hyperinsulinemia, and hyperlipidemia

indicative of insulin-resistant diabetes (Ross et al., 1993). Thiazolidinedione

administration to these animals improves the serum lipid profile, but results are

conflicting on the effects on glucose tolerance, with one study showing decreases in

insulin (Burant et al., 1997) and another showing no change (Chao et al., 2000). Using

tissue-specific PPARy knock-out mice, the question of whether TZDs directly or

indirectly affect insulin resistance has been researched. Targeted deletion of PPARy in

adipose results in adipose hypertrophy, elevated plasma NEFA and TAG, increased

hepatic gluconeogenesis and insulin resistance, without changes in insulin sensitivity of

muscle (He et al., 2003). These observations indicate that changes in adipose function

via PPARy result in changes in hepatic function with minimal effects in muscle.

In addition to the effects on adipose tissue and insulin resistance, activation of

PPARy seems to play a role in atherosclerosis. Again, the positive effects of TZD

treatment on decreased risk of atherosclerosis may be secondary to the improvement in






46


lipid profile, but PPARy activation may also have a direct effect on the formation and

progression of atherosclerotic lesions. Peroxisome proliferator-activated receptor y

activation inhibits leukocyte-endothelial cell interaction, a critical inflammatory response

in the formation of atherosclerotic plaques (Jackson et al., 1999). Activation by TZDs

also inhibits the expression of vascular cell adhesion molecule (Pasceri et al., 2000) and

E-selectin (Nawa et al., 2000), which would reduce the "homing" of monocyte and

macrophage cells to atherosclerotic plaques.














CHAPTER 3
EFFECTS OF N-3 AND N-6 FATTY ACIDS ON LIPID METABOLIZING GENES
AND HIGH-DENSITY LIPOPROTEIN CHOLESTEROL PRODUCTION IN
CULTURED HUMAN AND RAT HEPATOCYTES

Introduction

Dietary polyunsaturated fatty acids (PUFA) have been shown to lower blood

triglycerides, alter the blood lipid profile, decrease intramuscular lipid droplet size,

improve insulin sensitivity, and enhance glucose utilization (Jump and Clarke, 1999).

Since the observation that PUFAs could inhibit hepatic lipogenesis in mice (Allmann and

Gibson, 1965), numerous studies have demonstrated that diets rich in PUFAs influence

metabolic changes by coordinately suppressing lipid synthesis in the liver and enhancing

fatty acid oxidation in both liver and skeletal muscle (Jump and Clarke, 1999). The

PUFA induction of genes encoding proteins involved in lipid oxidation include

3-hydroxy, 3-methylglutaryl CoA synthase (Rodriguez et al., 1994), carnitine

palmitoyltransferase, fatty acid binding proteins and peroxisomal acyl-CoA oxidase

(ACO; Reddy and Hashimoto, 2001). With the discovery of a new member of the steroid

hormone receptor superfamily, the peroxisome proliferator-activated receptor (PPAR;

Issemann and Green, 1990) and the discovery that certain fatty acids (FA) and their

derivatives can specifically bind PPARs (Gottlicher et al., 1992), the possibility arose that

PUFAs mediate metabolic effects via alteration of PPAR activity. In the liver, the

predominant isoform is PPARa; therefore this isoform has become the primary focus of

studies involving the liver.









The objective of this study was to examine the short term effects of FAs of

differing levels of saturation and bond position on lipid metabolizing gene expression and

high-density lipoprotein (HDL) cholesterol production in HepG2 and H-4-II-E cells.

Based on both dietary and in vitro studies of lipid metabolism, we hypothesized that FAs

of differing saturation and double bond position may have differing effects on ACO,

3-hydroxy, 3-methylglutaryl CoA reductase (HMG-R), and apolipoprotein A-I (Apo A-I)

gene expression. Also, because several fatty acids and their derivatives are known

ligands for PPARs, we hypothesized that fatty acids may act on lipid metabolizing genes

through activation of PPARa in the liver.

Materials and Methods

Materials

Polystyrene tissue culture dishes (100 x 20 mm) were purchased from Corning

(Corning Glass Works, Corning, NY). The antibiotic/antimycotic (ABAM), sodium

pyruvate, fatty acid-free bovine serum albumin (BSA), stearic acid (ST), WY 14,643, and

MK886 were from Sigma Chemical Co. (St. Louis, MO). Minimum Essential Medium

(MEM), phenol red-free MEM, Hanks Balanced Salt Solution (HBSS) and TriZol reagent

were from GIBCO BRL (Carlsbad, CA). The fetal bovine serum (FBS) was from Atlanta

Biologicals (Norcross, GA). Linoleic, linolenic, and eicosapentaenoic acids were from

Cayman Chemicals (Ann Arbor, MI). BioTrans nylon membrane and [a-32P]

deoxycytidine triphosphate (SA 3000 Ci/nmol) were from MP Biolomedicals (Atlanta,

GA). The Enzyme Color Solution, Reacting Solution, and HDL Calibrator were from

Wako Diagnostics (Richmond, VA).









Cell Culture and Treatment

HepG2 (ATCC # HB-8065; Manassas, VA) and H-4-II-E (ATCC # CRL-1548;

Manassas, VA) cells were suspended in 10 mL of growth medium (MEM), containing 2.2

g/L sodium bicarbonate, 1.0 mM sodium pyruvate, 1% (v/v) ABAM and 10% FBS. Cells

were cultured at 370C in a humidified atmosphere containing 95% 02 and 5% CO2.

Cultures were replenished with fresh medium every 2 d until cells were approximately

90% confluent. Cells were washed twice with HBSS, and cultured in fresh serum-free

medium containing appropriate treatments for an additional 24 h.

Stock solutions of fatty acids were stored at -200C. At preparation of treatments,

fatty acids were mixed with serum-free culture medium containing 33 mg/mL of fatty

acid-free BSA to a concentration of 1 mM. This mixture was incubated for 2 h at 370C to

allow complexation of the fatty acids with BSA and then further diluted in culture

medium to a final treatment concentration of 100 pM of fatty acids.

To investigate the effects of supplemental PUFAs on hepatic gene expression and

cholesterol synthesis, HepG2 and H-4-II-E cells were treated with stearic (ST), linoleic

(LA), linolenic (LNA) or eicosapentoenoic (EPA) acid (100 pM). Sub-confluent cells

were incubated with serum-free medium alone (Control) or with appropriate treatments

(listed above) completed with BSA, for a period of 24 h. Cells were rinsed twice with 10

mL of HBSS. The remaining cell monolayer was then lysed in 3 mL of TriZol reagent,

and stored at -800C for subsequent mRNA analysis. The same fatty acid treatments were

repeated, using phenol red-free MEM. After incubation, conditioned media were

collected and stored at -200C until lipid extraction and HDL cholesterol analysis.

To determine whether fatty acid effects on gene expression involves PPARa

activation, confluent HepG2 and H-4-II-E cells were treated with ST (100 pM), the









PPARa agonist WY 14,643 (10 [M), or a combination of fatty acid and WY 14,643.

Additional sets of culture dishes were incubated with ST alone, the PPARa inhibitor

MK886 (10 aM; Kehrer et al., 2001), or a combination of ST and MK886. After a 24 h

incubation, cells were washed twice with 10 mL of HBSS, lysed with TriZol, and stored

at -80C until mRNA analysis.

RNA Isolation and Analysis

Total cellular RNA was isolated from cells using TriZol reagent according to the

manufacturer's instructions. Ten micrograms of total RNA was fractioned in a 1.0%

agarose formaldehyde gel following previously described protocols (Ing et al., 1996)

using the MOPS buffer (Fisher Scientific, Pittsburgh, PA) and transferred to a Biotrans

nylon membrane by downward capillary transfer in 20X SSC (3 M NaC1, 0.3 M sodium

citrate, pH 7.0) using the TurboBlotting system (Schleicher and Schuel, Keene, NH).

Nylon membranes were cross-linked by exposure to a UV light source for 90 sec and

baked at 800C for 1 h. Membranes were incubated for 2 h at 500C in ultrasensitive

hybridization buffer (ULTRAhyb; Ambion, Austin, TX) followed by an overnight

incubation at 500C in the same ULTRAhyb solution containing the 32P-labeled acyl-CoA

oxidase (ACO), 3-hydroxy, 3-methylglutaryl CoA reductase (HMG-R) and

apolipoprotein A-I (Apo A-I) cDNA probes. Probes were generated by RT-PCR for

ACO (forward 5'-CCGGAGCTGCTTACACACAT-3'; reverse 5'-GGTCATACGTGGC

TGTGGTT-3'), HMG-R (forward 5'-TCCTTGGTGATGGGAGCTTGTTGTG-3';

reverse 5'-TGCGAACCCTTCAGATGTTTCGAGC-3'), human Apo A-I (forward

5'-AAGACAGCGGCAGAGACTAT-3'; reverse 5'-ATCTCCTCCTGCCACTTCTT-3'),

and rat Apo A-I (forward 5'-AAGGACAGCGGCAGAGACTA-3'; reverse

5'-CCACAACCTTTAGATGCCTT-3'). The sizes and sequences of these cDNA probes









were verified by DNA sequencing prior to their use in Northern blot analysis. Filters

were sequentially washed in 2X SSC (1X= 0.15 M sodium chloride, 0.015 M sodium

citrate)-0.1% SDS and in 0.lx SSC-0.1% SDS twice each at 500C and then exposed to

X-ray film to detect radiolabeled bands. Equal loading of total RNA for each

experimental sample was verified by comparison to 18S rRNA ethidium bromide

staining.

Lipid Extraction

Total lipids were extracted from conditioned media as described by Bligh and Dyer

(1959), with modifications. For each sample, 2 mL of conditioned media was aliquotted

into a 20 mL glass screw-top vial. Fourteen mL of chloroform:methanol (2:1, v/v) was

then added and the vials were vortexed for 5 minutes. The vials were then centrifuged at

1700 rpm for 5 minutes. The bottom lipid-containing chloroform layer was transferred to

a clean, dry, pre-weighed vial, placed in a 37C water bath, and dried under nitrogen gas.

Dry samples were placed in a 50C oven for 10 minutes and placed in a desiccator to cool

to room temperature. Samples were weighed, and lipid weight was determined by

difference. The sample was resuspended in chloroform and stored at -200C until HDL

cholesterol analysis.

HDL Cholesterol Assay

Lipid extracts from conditioned media were analyzed using a commercially

available L-Type HDL-C kit, following the manufacturer's directions. Briefly, using a

96-well plate, 3 [tL of sample was pipetted into each well. Two hundred seventy [tL of

Enzyme Color Solution (R1) was added, and the plate was incubated for 5 minutes at

37C. Ninety [tL of Reacting Solution (R2) was then added, and the plate was incubated

another 5 minutes at 370C. The absorbance at 600 nm was measured using the









SpectraMax 340 PC microplate reader (Molecular Devices, Sunnyvale, CA), and the

concentration of the samples was calculated by plotting against a standard curve.

Statistical Analysis

All hybridization signals as measured by densitometry were evaluated by least

squares analysis of variance (ANOVA) using the General Linear Model (GLM)

procedure of the SAS software package (SAS Institute Inc, Cary, NC). In each

experiment, treatments were run in duplicate, and the whole experiment was also

duplicated, giving n=4 plates per treatment. The general model for mRNA analysis

included experiment, treatment, and experiment x treatment interaction. In mRNA

analyses, densitometric values for target genes were expressed as ratios of target gene

densitometric values over the corresponding 18S rRNA densitometric values. For HDL

cholesterol concentration, the sources of variation included experiment, treatment,

experiment x treatment interaction, and plate (experiment x treatment). The plate, nested

within experiment and treatment, was considered a random variable, and therefore the

plate variance was used as an error term to test the effects of experiment, treatment, and

experiment x treatment interaction. Treatment means were further compared using

preplanned orthogonal contrasts. These contrasts were control vs. fat treatment (ST, LA,

LNA, EPA), saturated fat (ST) vs. PUFA (LA, LNA, EPA), n-6 (LA) vs. n-3 (LNA,

EPA); and LNA vs. EPA. For all responses, the two cell lines were analyzed separately.

Results

Effects of Fatty Acids on HepG2 Cells

Steady-state levels of ACO mRNA were not affected by any FA treatment in

HepG2 cells (P = 0.3; Figure 3-1). Concentrations of HMG-R mRNA transcript were

greater (+24%, P = 0.006) in HepG2 cells treated with ST than in PUFA-treated cells









(Figure 3-2). Concentrations of Apo A-I mRNA transcript were greater (+15%, P = 0.05)

in HepG2 cells treated with FA than in control cells (Figure 3-3). There were no

differences in HDL cholesterol concentration among any of the treatments (P = 0.9;

Figure 3-4).

Effects of Fatty Acids on H-4-II-E Cells

In the H-4-II-E cells, ACO mRNA expression was greater (+26%, P = 0.004) in

ST-treated cells as compared to PUFA-treated cells (Figure 3-5). Concentrations of

HMG-R mRNA were greater in ST-treated as compared to PUFA-treated cells (+27%;

P = 0.002), in n-3 (EPA and LNA)-treated as compared to n-6 (LA)-treated cells (+30%;

P = 0.004), and in EPA-treated as compared to LNA-treated cells (+49%; P < 0.001),

with the EPA treatment showing the greatest induction of HMG-R mRNA transcript

(Figure 3-6). Similarly, steady-state levels of Apo A-I mRNA were increased in

ST-treated cells as compared to PUFA-treated cells (+39%; P < 0.001) and in

EPA-treated cells as compared to LNA-treated cells (+31%; P = 0.008; Figure 3-7). As

compared to n-6 FA, n-3 FA increased (+79%; P = 0.0002) HDL cholesterol

concentration by H-4-II-E cells, with the effect predominantly deriving from the large

increase (+84%; P < 0.0001) in production with LNA as compared to EPA (Figure 3-8).

Role of PPARa in Stearic Acid-Induced Effects on Gene Expression

Co-incubation of HepG2 cells with ST and 10 atM WY 14,643, a specific PPARa

agonist, decreased (-9%; P = 0.04) ACO mRNA expression as compared to ST alone.

There was no detectible effect on ACO mRNA with the use of the agonist alone (P = 0.8;

Figure 3-9). WY 14,643 decreased both basal (-32%; P = 0.0002) and ST-induced

(-10%; P = 0.02) expression of HMG-R mRNA (Figure 3-10). Use of the PPARaagonist









alone (P = 0.5) or in combination with ST (P = 0.4) had no effects on Apo A-I mRNA

(Figure 3-11).

In HepG2 cells, incubation with 10 tM MK886, a specific PPARainhibitor

increased (P < 0.05) basal production of all three gene transcripts (Figures 3-12, 3-13,

and 3-14). Co-incubation with MK886 had no effects on ST-induced expression of any

of the genes.

Co-incubation of H-4-II-E cells with WY 14,643 increased (+22%; P = 0.04) basal

levels and enhanced (+38%; P = 0.0003) the effect of ST on ACO gene expression

(Figure 3-15). Both basal (-45%; P = 0.01) and ST-induced (-32%; P = 0.03) HMG-R

mRNA expression were decreased with the use of the PPARa agonist (Figure 3-16). The

abundance of ST-induced Apo A-I mRNA transcript was enhanced (+29%; P = 0.001) by

the use of WY 14,643 (Figure 3-17). Basal levels of Apo A-I mRNA were unaffected

(P = 0.2).

In H-4-II-E cells, incubation with MK886 attenuated (-28%; P = 0.001) the effects

of ST on ACO mRNA expression (Figure 3-18). The PPARa inhibitor increased (+29%;

P = 0.003) the basal concentration of HMG-R mRNA transcript, but had no effects

(P = 0.8) on ST-induced gene expression (Figure 3-19). The concentration of both basal

(-96%; P = 0.01) and ST-induced (-39%; P = 0.03) Apo A-I mRNA transcript was

reduced by the use of MK886 (Figure 3-20).

Discussion

Dietary fat has been implicated as a major factor in many areas of health and

disease. However, it has been suggested by numerous studies that all fats may not have

the same effects. In this study, both human and rat hepatoma cells were used as models,

as it also has been suggested that species differences exist in fat metabolism (Bergen and









Mersmann, 2005). In HepG2 (human) liver cells, ACO mRNA expression was

unaffected by any FA treatment. In contrast, in the H-4-II-E (rat) liver cells, ACO

mRNA expression was induced by ST only. Other studies, however, have shown

up-regulation of ACO mRNA in rat liver by dietary PUFAs as well as by saturated fats

(Berthou et al., 1995). In HepG2 cells, it has been shown that PUFAs of differing

saturation and length can regulate ACO mRNA in a dose-dependent and differential

manner (Rise and Galli, 1999). In a human retinoblastoma cell line, low concentrations

of supplemental n-3 PUFA increased ACO mRNA, whereas high concentrations of the

FA decreased it (Langelier et al., 2003). Consistent with our findings in rat cells, pigs fed

a tallow-based diet high in saturated fat had an increased concentration of ACO mRNA

as compared to fish-oil fed animals (Ding et al., 2003).

3-hydroxy, 3-methylglutaryl CoA reductase is the rate limiting enzyme in

cholesterol synthesis, and its inhibition is the target of the station class of drugs, used in

the treatment of hyperlipidemias. In this study, we showed that in HepG2 cells, HMG-R

mRNA was up-regulated by ST as compared to the PUFAs, whereas in the H-4-II-E cells,

it was up-regulated by both ST and EPA. Consistent with our findings in rodent cells, in

C3H mice fed diets differing in fat composition, HMG-R mRNA was increased to a

greater extent in mice fed the PUFA diet than in those fed the saturated fat diet (Cheema

and Agellon, 1999). In Reuber H35 rat hepatoma cells, incubation with either saturated

fats or PUFAs increased HMG-R enzyme activity (Garcia-Pelayo et al., 2003). Enzyme

activity of HMG-R also has been shown to be increased in mice fed a diet high in PUFAs

(Kuan and Dupont, 1989).









Apolipoprotein A-I is the predominant lipoprotein associated with HDL cholesterol

and is essential for its normal metabolism. Deletion of the Apo A-I gene in humans

results in very low plasma concentrations of HDL cholesterol and premature coronary

artery disease (Schaefer et al., 1982). Dietary fat has the ability to modulate plasma

lipids, and may act, in part, by effects on apolipoproteins. In this study, we showed that,

in HepG2 cells, Apo A-I mRNA was up-regulated by all FA. However, no effects were

seen in HDL cholesterol concentration in the culture media. This is supported in a study

by Dashti and coworkers (2002) in which HDL concentration was not different between

LA- and saturated fat-treated HepG2 cells. In Golden-Syrian hamsters, an effective

model for human diet and blood lipid interactions, canola and soybean oils increased Apo

A-I mRNA as compared to a butter diet, though HDL concentrations were lowered in the

diets containing unsaturated as compared to saturated fats (Dorfman et al., 2005). In the

H-4-II-E cells, ST increased Apo A-I mRNA concentration as compared to the

PUFA-treated cells. In contrast to current findings, Sprague-Dawley rats fed diets high in

saturated fat or PUFAs showed no differences in Apo A-I amounts (Hatahet et al., 2003).

However, the saturated fat diet contained primarily palmitic acid, not stearic acid, as in

this study.

As fatty acids and their derivatives have been identified as potential ligands for

peroxisome proliferator-activated receptors (PPAR), we investigated the possibility that

fatty acid effects in the two cell lines may be mediated by PPARa. Incubation of HepG2

cells with WY 14,643, a PPARa agonist, had no effects on basal expression of ACO or

Apo A-I mRNA. In the H-4-II-E cells, however, incubation with the agonist not only

enhanced ST-induced ACO and Apo A-I mRNA expression but also increased basal









expression of ACO mRNA. Not unexpectedly, use of the PPARa inhibitor, MK886, was

able to cause the opposite effect, blocking the effects of ST on ACO and Apo A-I mRNA

expression in H-4-II-E cells. Although ACO is an established PPARa responsive gene

(Tugwood, et al., 1992), species differences do exist. It is questionable whether the

PPAR response element of human ACO is active (Woodyatt et al., 1999). Dietary studies

have shown that rodents are responsive to the effects of PPARa activation, but

non-rodent species, such as primates and guinea pigs, are resistant or unresponsive to

some of the negative effects (Bentley et al., 1993; Cattley et al., 1998). In a

comprehensive analysis of gene expression in human and rat hepatoma cells by

microarray analysis, only rat ACO mRNA was responsive to WY 14,643 (Vanden

Heuvel et al., 2003). Other genes that may be differentially regulated in human and rat

liver include cytosolic aspartate aminotransferase (Tomkiewicz et al., 2004), peroxisomal

3-oxoacyl-CoA thiolase (Lawrence et al., 2001), and catalase (Ammerschlaeger et al.,

2004). Additionally, different PPARa agonists may regulate lipid metabolism in a

compound-dependent manner. A recent study by Duez and coworkers (2005) showed

that, in mice, fenofibrate and gemfibrozil, both stimulate ACO mRNA expression, but

only fenofibrate greatly induces Apo A-I gene expression. Interestingly, although effects

of PPARa activation or inhibition on ACO and Apo A-I mRNA were different between

the human and rat cell lines, effects on HMG-R mRNA were similar. In both cell lines,

activation of PPARa by WY 14,643 caused a decrease in basal and ST-induced HMG-R

mRNA expression. Inhibition of PPARa by MK886 increased HMG-R mRNA

expression to a level similar to that induced by ST treatment alone, suggesting that ST









effects are mediated by PPARa. The findings of this study, in combination with other

reports, strongly suggest a species-specific role for PPARa in gene regulation.

Summary

In HepG2 cells, ST up-regulated HMG-R gene expression as compared to PUFAs.

As compared to control, in this cell line, all FA in this experiment up-regulated Apo A-I

gene expression. When PPARa was selectively activated, the effect of ST on ACO gene

expression was decreased, whereas both basal and ST-induced HMG-R gene expression

were decreased. Incubation with the PPARa inhibitor was able to decrease the basal

production of all three genes, but had no effects on ST-induced gene expression.

In H-4-II-E cells, ST up-regulated ACO, HMG-R, and Apo A-I gene expression as

compared to the PUFAs. Selective activation of PPARa increased basal levels of ACO

and further enhanced the effect of ST on ACO and Apo A-I mRNA. Conversely,

selective activation of PPARa decreased basal levels of HMG-R and blocked the effect of

ST on HMG-R mRNA. Incubation with the PPARa inhibitor was able to decrease the

effects of ST-induced ACO and Apo A-I mRNA, as well as decrease the basal

concentration of Apo A-I mRNA and increase the basal concentration of HMG-R

mRNA. Together, these results indicate that FAs likely regulate lipid metabolizing genes

in the liver through a PPARa-dependent mechanism. However, due to different

responses in the human and rat hepatoma cell lines, the net effects are likely species

specific.










(A)
(A)C ST LA LNA EPA


ACO 3.2 kb

rRNA 18S



(B) 0.7

0.6

W 0.5
o
Z 0
S0.4

@ 0.3
E
0
z 0.2

0.1

0.0
Control ST LA LNA EPA
Treatments

Figure 3-1. Effect of long-chain FA on ACO mRNA expression in HepG2 cells. Ten
micrograms of total cellular RNA isolated from control and FA-treated
HepG2 cells were subjected to Northern blot analysis, and resulting
densitometric values were analyzed by the GLM procedure of SAS. A) A
representative Northern blot. B) Means SEM calculated over two
experiments (n = 4 for each treatment). To further examine treatment effects,
means were separated using orthogonal contrasts. Contrast 1: Control vs. (ST
+ LA + LNA + EPA), P = 0.9; Contrast 2: ST vs. (LA + LNA + EPA),
P = 0.09; Contrast 3: LA vs. (LNA + EPA), P = 0.7; Contrast 4: LNA vs.
EPA, P = 0.2.








C ST LA


HMG-R
rRNA

(B) 1.0

0.8 -


o0
E'o
nN
E
v1


Control ST LA LNA


4.1-4.7kb
18S


EPA


Treatments
Figure 3-2. Effect of long-chain FA on HMG-R mRNA expression in HepG2 cells. Ten
micrograms of total cellular RNA isolated from control and FA-treated
HepG2 cells were subjected to Northern blot analysis, and resulting
densitometric values were analyzed by the GLM procedure of SAS. A) A
representative Northern blot. B) Means SEM calculated over two
experiments (n = 4 for each treatment). To further examine treatment effects,
means were separated using orthogonal contrasts. Contrast 1: Control vs. (ST
+ LA + LNA + EPA), P = 0.8; Contrast 2: ST vs. (LA + LNA + EPA),
P = 0.006; Contrast 3: LA vs. (LNA + EPA), P = 0.6; Contrast 4: LNA vs.
EPA, P = 0.9.


~irrI


LNA EPA







(A)
Apo A-I

rRNA

(B) 0.6

0.5 -


E,
E'-o
<0
z


Treatments


Figure 3-3. Effects of long-chain FA on Apo A-I mRNA expression in HepG2 cells. Ten
micrograms of total cellular RNA isolated from control and FA-treated
HepG2 cells were subjected to Northern blot analysis, and resulting
densitometric values were analyzed by the GLM procedure of SAS. A) A
representative Northern blot. B) Means SEM calculated over two
experiments (n = 4 for each treatment). To further examine treatment effects,
means were separated using orthogonal contrasts. Contrast 1: Control vs. (ST
+ LA + LNA + EPA), P = 0.05; Contrast 2: ST vs. (LA + LNA + EPA),
P = 0.7; Contrast 3: LA vs. (LNA + EPA), P = 0.3; Contrast 4: LNA vs. EPA,
P = 0.4.


LNA


EPA


0.9 kb
18S


Control


- ~ im usN amumn i iui n u a ai


LNA


EPA


I






















5




E
'-3



.2

S1



0
Control ST LA LNA EPA

Treatment

Figure 3-4. Effects of long-chain FA on HDL cholesterol production in HepG2 cells.
Data represents least square means SEM calculated over two experiments.
To further examine treatment effects, means were separated using orthogonal
contrasts. Contrast 1: Control vs. (ST + LA + LNA + EPA), P = 0.8; Contrast
2: ST vs. (LA + LNA + EPA), P = 0.4; Contrast 3: LA vs. (LNA + EPA),
P = 0.9; Contrast 4: LNA vs. EPA, P = 0.9.










C ST


LA LNA EPA


(B)


0.14

0.12

0.10

0.08

0.06

0.04

0.02

0.00


Control ST LA LNA EPA


Treatments

Figure 3-5. Effects of long-chain FA on ACO mRNA expression in H-4-II-E cells. Ten
micrograms of total cellular RNA isolated from control and FA-treated
H-4-II-E cells were subjected to Northern blot analysis, and resulting
densitometric values were analyzed by the GLM procedure of SAS. A) A
representative Northern blot. B) Means SEM calculated over two
experiments (n = 4 for each treatment). To further examine treatment effects,
means were separated using orthogonal contrasts. Contrast 1: Control vs. (ST
+ LA + LNA + EPA), P = 0.2; Contrast 2: ST vs. (LA + LNA + EPA),
P = 0.004; Contrast 3: LA vs. (LNA + EPA), P = 0.6; Contrast 4: LNA vs.
EPA, P = 0.07.


(A)


ACO

rRNA


3.2 kb

18S










C ST LA


LNA EPA


0.20


0.15


0.10


0.05


0.00


Control ST LA LNA EPA


Treatments

Figure 3-6. Effects of long-chain FA on HMG-R mRNA expression in H-4-II-E cells.
Ten micrograms of total cellular RNA isolated from control and FA-treated
H-4-II-E cells were subjected to Northern blot analysis, and resulting
densitometric values were analyzed by the GLM procedure of SAS. A) A
representative Northern blot. B) Means SEM calculated over two
experiments (n = 4 for each treatment). To further examine treatment effects,
means were separated using orthogonal contrasts. Contrast 1: Control vs. (ST
+ LA + LNA + EPA), P = 0.9; Contrast 2: ST vs. (LA + LNA + EPA),
P = 0.002; Contrast 3: LA vs. (LNA + EPA), P = 0.004; Contrast 4: LNA vs.
EPA, P < 0.001.


(A)


HMG-R

rRNA


(B)


0.25


4.5 kb

18S










(A) C ST LA LNA EPA

Apo A-I 0.9 kb

rRNA 18S


(B) 0.07

0.06

u) 0.05

nO
0.04
E'o



z 0.02

0.01

0.00
Control ST LA LNA EPA
Treatments

Figure 3-7. Effects of long-chain FA on Apo A-I mRNA expression in H-4-II-E cells.
Ten micrograms of total cellular RNA isolated from control and FA-treated
H-4-II-E cells were subjected to Northern blot analysis, and resulting
densitometric values were analyzed by the GLM procedure of SAS. A) A
representative Northern blot. B) Means SEM calculated over two
experiments (n = 4 for each treatment). To further examine treatment effects,
means were separated using orthogonal contrasts. Contrast 1: Control vs. (ST
+ LA + LNA + EPA), P = 0.6; Contrast 2: ST vs. (LA + LNA + EPA),
P < 0.001; Contrast 3: LA vs. (LNA + EPA), P = 0.6; Contrast 4: LNA vs.
EPA, P = 0.008.

















14


_. 12

I-c
E 10




S6

-J 4




0
Control ST LA LNA EPA

Treatment

Figure 3-8. Effects of long-chain FA on HDL cholesterol production in H-4-II-E cells.
Data represents least square means SEM calculated over two experiments.
To further examine treatment effects, means were separated using orthogonal
contrasts. Contrast 1: Control vs. (ST + LA + LNA + EPA), P = 0.3; Contrast
2: ST vs. (LA + LNA + EPA), P = 0.06; Contrast 3: LA vs. (LNA + EPA),
P = 0.0002; Contrast 4: LNA vs. EPA, P < 0.0001.








C ST


PPARca
Agon


Agonist ST + A


Treatments
Figure 3-9. Effect of WY 14,643 on ACO mRNA response to ST in HepG2 cells. Ten
micrograms of total cellular RNA isolated from control and treated HepG2
cells were subjected to Northern blot analysis, and resulting densitometric
values were analyzed by the GLM procedure of SAS. A) A representative
Northern blot. B) Means SEM calculated over two experiments (n = 4 for
each treatment). To further examine treatment effects, means were separated
using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (ST + ST+A),
P = 0.2; Contrast 2: ST vs. ST+A, P = 0.04; Contrast 3: Control vs. Agonist,
P= 0.8.


(A)


ACO
rRNA


ST +
Agon


3.2 kb
18S


(B)


CO
r0

EZ
O
Z


0.30

0.25

0.20

0.15

0.10

0.05

0.00


Control


1EBe










PPARao
C ST Agon


ST Agonist


Treatments

Figure 3-10. Effect of WY 14,643 on HMG-R mRNA response to ST in HepG2 cells.
Ten micrograms of total cellular RNA isolated from control and treated
HepG2 cells were subjected to Northern blot analysis, and resulting
densitometric values were analyzed by the GLM procedure of SAS. A) A
representative Northern blot. B) Means SEM calculated over two
experiments (n = 4 for each treatment). To further examine treatment effects,
means were separated using orthogonal contrasts. Contrast 1: (Control +
Agonist) vs. (ST + ST+A), P = 0.003; Contrast 2: ST vs. ST+A, P = 0.02;
Contrast 3: Control vs. Agonist, P = 0.002.


HMG-R

rRNA


ST +
Agon


4.1-4.7kb

18S


(B)


oo
Z 0

E o


SE
I -


0.30


0.25


0.20


0.15


0.10


0.05


0.00


Control


ST + A








(A)


PPARao
C ST Agon


Apo A-I

rRNA


(B)


0.25


0.20 -


0.15 [


0.00


Control


Agonist


ST +
Agon


0.9 kb

18S


ST + A


Treatments
Figure 3-11. Effect of WY 14,643 on Apo A-I mRNA response to ST in HepG2 cells.
Ten micrograms of total cellular RNA isolated from control and treated
HepG2 cells were subjected to Northern blot analysis, and resulting
densitometric values were analyzed by the GLM procedure of SAS. A) A
representative Northern blot. B) Means SEM calculated over two
experiments (n = 4 for each treatment). To further examine treatment effects,
means were separated using orthogonal contrasts. Contrast 1: (Control +
Agonist) vs. (ST + ST+A), P = 0.6; Contrast 2: ST vs. ST+A, P = 0.4;
Contrast 3: Control vs. Agonist, P = 0.5.


Ev
E-o


z


0.10

0.05


SI" MINIS S EMI IS "I""IM S "SI""I


%111








(A)


PPARa
C ST Inhib


ACO
rRNA


(B)


0.35


0.30

0.25

0.20

0.15


0.10

0.05

0.00


ST +
Inhib


3.2 kb
18S


Control


Inhib


ST+I


Treatments
Figure 3-12. Effect of MK886 on ACO mRNA response to ST in HepG2 cells. Ten
micrograms of total cellular RNA isolated from control and FA treated HepG2
cells were subjected to Northern blot analysis, and resulting densitometric
values were analyzed by the GLM procedure of SAS. A) A representative
Northern blot. B) Means SEM calculated over two experiments (n = 4 for
each treatment). To further examine treatment effects, means were separated
using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (ST + ST+I),
P = 0.15; Contrast 2: ST vs. ST+I, P = 0.6; Contrast 3: Control vs. Inhib,
P = 0.03.


aIsMIistsMMhi~tIsIMMIs ~1ISM


iCI








(A)


PPARa
Inhib


ST +
Inhib


HMG-R

rRNA


(B) o.s


4.1-4.7kb

18S


0.4 F


0.3 F


0.2 h


Control


Inhib


ST + I


Treatments


Figure 3-13. Effect of MK886 on HMG-R mRNA response to ST in HepG2 cells. Ten
micrograms of total cellular RNA isolated from control and treated HepG2
cells were subjected to Northern blot analysis, and resulting densitometric
values were analyzed by the GLM procedure of SAS. A) A representative
Northern blot. B) Means SEM calculated over two experiments (n = 4 for
each treatment). To further examine treatment effects, means were separated
using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (ST + ST+I),
P = 0.5; Contrast 2: ST vs. ST+I, P = 0.4; Contrast 3: Control vs. Inhib,
P= 0.01.


hikIMaMM wMhiI IMMsMahIwiME


I I1










(A) PPARa ST +
C ST Inhib Inhib

Apo A-I 0.9 kb

rRNA 18S



(B) 0.6


0.5

VU
0.4

Evo
0.3


c 0.2
z

0.1

0.0
Control ST Inhib ST + I

Treatments

Figure 3-14. Effect of MK886 on Apo A-I mRNA response to ST in HepG2 cells. Ten
micrograms of total cellular RNA isolated from control and treated HepG2
cells were subjected to Northern blot analysis, and resulting densitometric
values were analyzed by the GLM procedure of SAS. A) A representative
Northern blot. B) Means SEM calculated over two experiments (n = 4 for
each treatment). To further examine treatment effects, means were separated
using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (ST + ST+I),
P = 0.01; Contrast 2: ST vs. ST+I, P = 0.6; Contrast 3: Control vs. Inhib,
P = 0.02.







PPARao
Agon


Control


ST Agonist


Treatments
Figure 3-15. Effect of WY14,643 on ACO mRNA response to ST in H-4-II-E cells. Ten
micrograms of total cellular RNA isolated from control and treated H-4-II-E
cells were subjected to Northern blot analysis, and resulting densitometric
values were analyzed by the GLM procedure of SAS. A) A representative
Northern blot. B) Means SEM calculated over two experiments (n = 4 for
each treatment). To further examine treatment effects, means were separated
using orthogonal contrasts. Contrast 1: (Control + Agonist) vs. (ST + ST+A),
P = 0.007; Contrast 2: ST vs. ST+A, P = 0003; Contrast 3: Control vs.
Agonist, P = 0.04.


ACO
rRNA


ST +
Agon


(B) 0.08


3.2 kb
18S


0.06 1


<
OS
0


0.04


0.02 -


0.00


IUiI i "IE KM I Ci^i


ST + A


(A)


I I1







(A)


PPARa
Agon


ST +
Agon


4.5 kb
18S


HMG-R

rRNA

(B) 0.10 -

0.08


0.06

0.04


0.02

0.00


Control


3T Agonist
Treatments


ST + A


Figure 3-16. Effect of WY 14,643 on HMG-R mRNA response to ST in H-4-II-E cells.
Ten micrograms of total cellular RNA isolated from control and treated
H-4-II-E cells were subjected to Northern blot analysis, and resulting
densitometric values were analyzed by the GLM procedure of SAS. A) A
representative Northern blot. B) Means SEM calculated over two
experiments (n = 4 for each treatment). To further examine treatment effects,
means were separated using orthogonal contrasts. Contrast 1: (Control +
Agonist) vs. (ST + ST+A), P = 0.12; Contrast 2: ST vs. ST+A, P = 0.03;
Contrast 3: Control vs. Agonist, P = 0.01.


I^I^A ^I^^I^M I^I^A^^I EEMIM


I~rI








PPARa
C ST Agon


ST Agonist ST + A


Treatments
Figure 3-17. Effect of WY 14,643 on Apo A-I mRNA response to ST in H-4-II-E cells.
Ten micrograms of total cellular RNA isolated from control and treated
H-4-II-E cells were subjected to Northern blot analysis, and resulting
densitometric values were analyzed by the GLM procedure of SAS. A) A
representative Northern blot. B) Means SEM calculated over two
experiments (n = 4 for each treatment). To further examine treatment effects,
means were separated using orthogonal contrasts. Contrast 1: (Control
Agonist) vs. (ST + ST+A), P < 0.0001; Contrast 2: ST vs. ST+A, P = 0.001;
Contrast 3: Control vs. Agonist, P = 0.2.


(A)


Apo A-I

rRNA


ST +
Agon


(B)


0.05


0.9 kb

18S


0.04 1


Ev

E'-o
WQ0
'N
QE
<0


0.03

0.02


0.01

0.00


Control


^^I^I^ ^I^^SI I^i^M^ IMSIMII"M


I I1






76


(A) PPARa ST +
C ST Inhib Inhib

ACO 3.2 kb

rRNA 18S



(B) 0.30


0.25

v0
o 0.20


E D o.15
0

L0.10
Z

0.05


0.00
Control ST Inhib ST +I

Treatments

Figure 3-18. Effect of MK886 on ACO mRNA response to ST in H-4-II-E cells. Ten
micrograms of total cellular RNA isolated from control and treated H-4-II-E
cells were subjected to Northern blot analysis, and resulting densitometric
values were analyzed by the GLM procedure of SAS. A) A representative
Northern blot. B) Means SEM calculated over two experiments (n = 4 for
each treatment). To further examine treatment effects, means were separated
using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (ST + ST+I),
P = 0.5; Contrast 2: ST vs. ST+I, P = 0.001; Contrast 3: Control vs. Inhib,
P = 0.06.






77


(A) PPARa ST +
C ST Inhib Inhib

HMG-R 4.5 kb

rRNA 18s


(B) 0.12


0.10


0.08
zo
E'
0.06


Z 0.04
Z

0.02


0.00
Control ST Inhib ST + I

Treatments

Figure 3-19. Effect of MK886 on HMG-R mRNA response to ST in H-4-II-E cells. Ten
micrograms of total cellular RNA isolated from control and treated H-4-II-E
cells were subjected to Northern blot analysis, and resulting densitometric
values were analyzed by the GLM procedure of SAS. A) A representative
Northern blot. B) Means SEM calculated over two experiments (n = 4 for
each treatment). To further examine treatment effects, means were separated
using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (ST + ST+I),
P = 0.004; Contrast 2: ST vs. ST+I, P = 0.8; Contrast 3: Control vs. Inhib,
P= 0.003.








(A)


PPARa
C S Inhib


Apo A-I

rRNA


(B)


0.05


ST +
Inhib


0.9 kb

18s


0.04 1


Ev
E-o
N
0- Q
0
z


0.03 1


0.02 -


0.01

0.00


Control


Inhib


ST+ I


Treatments
Figure 3-20. Effect of MK886 on Apo A-I mRNA response to ST in H-4-II-E cells. Ten
micrograms of total cellular RNA isolated from control and treated H-4-II-E
cells were subjected to Northern blot analysis, and resulting densitometric
values were analyzed by the GLM procedure of SAS. A) A representative
Northern blot. B) Means SEM calculated over two experiments (n = 4 for
each treatment). To further examine treatment effects, means were separated
using orthogonal contrasts. Contrast 1: (Control + Inhib) vs. (ST + ST+I),
P = 0.003; Contrast 2: ST vs. ST+I, P = 0.03; Contrast 3: Control vs. Inhib,
P= 0.01.


" mY "%Y W%" A0A""P


I i I









ST LA LNA EPA


I
Activation PPARa Inhibition


HDL-C


HepG2 H-4-11-E


Figure 3-21. Regulation of lipid metabolizing genes and HDL cholesterol production by
long-chain fatty acids. In HepG2 cells, HMG-R mRNA was up-regulated by
ST as compared to the PUFAs. All fatty acids up-regulated Apo A-I mRNA
as compared to control. Activation of PPARa attenuated the effects of ST on
ACO and HMG-R gene expression. In H-4-II-E cells, ST up-regulated ACO,
HMG-R, and Apo A-I gene expression as compared to the PUFAs. Activation
of PPARa increased basal expression of ACO and enhanced ST effects on
ACO and Apo A-I mRNA. Both basal and ST-induced HMG-R mRNA levels
were decreased by PPARa activation. Inhibition of PPARa decreased basal
expression of Apo A-I and attenuated ST-induced expression of ACO and
Apo A-I mRNA. Basal concentrations of HMG-R mRNA were increased by
PPARa inhibition. As compared to n-6 PUFA, n-3 PUFA increased HDL
cholesterol production, with the effect predominantly deriving from the
increase due to LNA.














CHAPTER 4
EFFECTS OF ISOMERS OF CONJUGATED LINOLEIC ACID ON LIPID
METABOLIZING GENES AND HIGH-DENSITY LIPOPROTEIN CHOLESTEROL
PRODUCTION IN CULTURED HUMAN AND RAT HEPATOCYTES

Introduction

Conjugated linoleic acid (CLA) is a collective term for positional and geometric

isomers of linoleic acid (LA). Though over 16 individual isomers have been identified

(Rickert et al., 1999), only cis-9, trans-11 CLA and trans-10, cis-12 CLA are known to

possess biological activity (Pariza et al., 2000). Cis-9, trans-11 CLA is the predominant

CLA produced as an intermediate in the rumen during the biohydrogenation of dietary

LA and is commonly found in dairy products and ruminant meat. Dietary sources of

trans-10, cis-12 CLA derive predominantly from synthetic partial biohydrogenation and

is found in margarines, shortenings, and supplements (Gaullier et al., 2002). First

identified in grilled beef as a potential anti-carcinogen (Pariza and Hargraves, 1985),

numerous health benefits have been attributed to CLA mixtures, including actions as an

antiadipogenic (Park et al, 1997), antidiabetogenic (Houseknecht et al., 1998), and

antiatherosclerotic (Kritchevsky et al., 2004) agent. More recently, studies involving

individual isomers have shown that the two main isoforms can have different effects on

metabolism and cell function and may act through different signaling pathways (Wahle et

al., 2004). The metabolic responses to cis-9, trans-11 and trans-10, cis-12 CLA may

differ, but both isomers have implications for human health. Most studies have been

performed in animal models, with species differences observed. In particular, only some









of the findings attributed to animal models pertain to human subjects, and even when

comparing studies in humans, results are often inconclusive (Terpstra, 2004).

The objective of this study was to examine the short term effects of the two

biologically active isomers of CLA on lipid metabolizing gene expression and

high-density lipoprotein (HDL) cholesterol production in HepG2 (human) and H-4-II-E

(rat) hepatoma cell lines. Based on both dietary and in vitro studies of lipid metabolism,

we hypothesized that the different isomers of CLA may have differing effects on acyl-

CoA oxidase (ACO), 3-hydroxy, 3-methylglutaryl CoA reductase (HMG-R), and

apolipoprotein A-I (Apo A-I) gene expression. Also, because several fatty acids and their

derivatives are known ligands for peroxisome proliferator-activated receptors (PPAR),

we hypothesized that CLA isomers may act on lipid-metabolizing genes through

activation of PPARa in the liver.

Materials and Methods

Materials

Polystyrene tissue culture dishes (100 x 20 mm) were purchased from Corning

(Corning Glass Works, Corning, NY). The antibiotic/antimycotic (ABAM), sodium

pyruvate, fatty acid-free bovine serum albumin (BSA), WY 14,643, and MK886 were

from Sigma Chemical Co. (St. Louis, MO). Minimum Essential Medium (MEM), phenol

red-free MEM, Hanks Balanced Salt Solution (HBSS) and TriZol reagent were from

GIBCO BRL (Carlsbad, CA). The fetal bovine serum (FBS) was from Atlanta

Biologicals (Norcross, GA). Linoleic acid, cis-9, trans-11 CLA, and trans-10, cis-12

CLA were from Cayman Chemicals (Ann Arbor, MI). BioTrans nylon membrane and

[a-32P]deoxycytidine triphosphate (SA 3000 Ci/nmol) were from MP Biolomedicals









(Atlanta, GA). The Enzyme Color Solution, Reacting Solution, and HDL Calibrator were

from Wako Diagnostics (Richmond, VA).

Cell Culture and Treatment

HepG2 (ATCC # HB-8065; Manassas, VA) and H-4-II-E (ATCC # CRL-1548;

Manassas, VA) cells were cultured and fatty acids were completed as described in

chapter 3. To investigate the effects of supplemental CLA on hepatic gene expression

and cholesterol synthesis, HepG2 and H-4-II-E cells were treated with LA, cis-9,

trans-11 CLA, or trans-10, cis-12 CLA (100 [tM). Sub-confluent cells were incubated

with serum-free medium alone (Control) or with appropriate treatments (listed above)

completed with BSA, for a period of 24 h. Cells were then rinsed twice with 10 mL

HBSS. The remaining cell monolayer was then lysed in 3 mL TriZol reagent, and stored

at -800C for subsequent mRNA analysis. The same fatty acid (FA) treatments were

repeated, using phenol red-free MEM. After incubation, conditioned media were

collected and stored at -200C until lipid extraction and HDL cholesterol analysis.

To investigate whether CLA effects on gene expression involves PPARa

activation, confluent HepG2 and H-4-II-E cells were treated with trans-10, cis-12 CLA

isomer (100 tM), the PPARa agonist WY 14,643 (10 pM), or a combination of trans-10,

cis-12 CLA and WY 14,643. Additional sets of culture dishes were incubated with

trans-10, cis-12 CLA alone, the PPARa antagonist MK886 (10 lM; Kehrer et al., 2001),

or a combination of trans-10, cis-12 CLA and MK886. After 24 h of incubation, cells

were washed twice with 10 mL HBSS, lysed with TriZol, and stored at -800C until

mRNA analysis.